Introduction to Plant Physiology

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Introduction to Plant Physiology

Fourth Edition

William G. Hopkinsand

Norman P. A. HunerThe University of Western Ontario

John Wiley & Sons, Inc.

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Library of Congress Cataloging-in-Publication Data:

Hopkins, William G.Introduction to plant physiology / William G. Hopkins and Norman P. A. Huner. –4th ed.

p. cm.Includes index.ISBN 978-0-470-24766-2 (cloth)

1. Plant physiology. I. Huner, Norman P. A. II. Title.QK711.2.H67 2008571.2–dc22

2008023261

Printed in the United States of America

10 9 8 7 6 5 4 3 2 1

Preface

When the first edition of this text appeared thirteenyears ago, its writing was guided by several of objectives.

• The text should be suited for a semester course forundergraduate students encountering the subject ofplant physiology for the first time. It was assumedthat the student would have completed a first coursein botany or biology with a strong botanical compo-nent. The book should provide a broad frameworkfor those interested in pursuing advanced study inplant physiology, but it should also provide the gen-eral understanding of plant function necessary forstudents of ecology or agriculture.

• In keeping with the above objective, the text shouldfocus on fundamental principles of how plants workwhile attempting to balance the demands of bio-chemistry and molecular biology on the one hand,and traditional ‘‘whole-plant’’ physiology on theother.

• The text should be interesting and readable. Itshould include some history so the student appreci-ates how we arrived at our current understanding. Itshould also point to future directions and challengesin the field.

• The sheer breadth of plant physiology and therapidly expanding volume of literature in the fieldmake it impossible to include all of the relevantmaterial in an entry-level text. Consequently, thetext must be selective and focused on those topicsthat form the core of the discipline. At the sametime, the student should be introduced to the sig-nificance of physiology in the role that plants playin the larger world outside the laboratory.

While we have made every effort to retain thereadability and overall approach of previous editions;we have also introduced a number of significant changesin this fourth edition. Those changes include:

• For this edition the illustration program has beencompletely revised. Some figures have been deleted,others have been revised, and many new figures havebeen introduced. With the help of the publisher, wehave also introduced color into the illustrations. Theuse of color improves the clarity of the figures, drawsattention to important elements in the figure, andhelps students visualize the relationships between

the figure and the concepts described in the text.At the same time, we are mindful of costs and hopethat this has been done in a way that does not addsignificant cost to the student.

• The number of complex chemical structures inmany figures has been reduced and biosyntheticpathways have been simplified in order to providegreater emphasis on fundamental principles.

• We have removed the traditional introductorychapter on Cells, Tissues, and Organs anddistributed some of this information in chapters towhich it pertains directly.

• The list of references at the end of each chapter hasbeen updated throughout the new edition.

• All life depends on energy and water. Unlike previ-ous editions, the fourth edition begins with fourchapters that focus on the properties of water,osmosis, water potential, and plant–water relations,followed by a series of eight chapters dealing withbioenergetics, primary plant metabolism, and plantproductivity.

• A major change in this edition is the presence ofthree new chapters (13, 14, and 15). Using the basicinformation and concepts developed in chapters 1to 12, these chapters focus on the inherent plastic-ity of plants to respond to environmental changeon various time scales. This includes a discussionof abiotic and biotic stress, plant acclimation tostress, and finally, long-term, heritable adaptationsto environmental stress.

• We have revised the treatment of hormones becausemany instructors have told us that the separatetreatment of each hormone fits their syllabus better.The coverage of each hormone concludes with ageneral description of the current status of receptorsand signal transduction pathways.

• A new chapter focuses on the molecular genetics offlower and fruit development.

• A Glossary has been created for the new edition.

William G. HopkinsNorman P. A. HunerLondon, OntarioApril 2008

v

vi Preface

To the Student

This is a book about how plants work. It is about thequestions that plant physiologists ask and how they goabout seeking answers to those questions. Most of all,this book is about how plants do the things they do intheir everyday life.

The well-known conservationist John Muir oncewrote: When we try to pick out anything by itself, we findit hitched to everything in the universe. Muir might wellhave been referring to the writing of a plant physiologytextbook. The scope of plant physiology as a scienceis very broad, ranging from biophysics and moleculargenetics to environmental physiology and agronomy.Photosynthetic metabolism not only provides carbonand energy for the growing plant, but also determinesthe capacity of the plant to withstand environmentalstress. The growth and development of roots, stems,leaves, and flowers are regulated by a host of interactingfactors such as light, temperature, hormones, nutrition,and carbon metabolism. As a matter of practical necessitymore than scientific reality, we have treated many ofthese topics in separate chapters. To get the most out ofthis book, we suggest you be aware of these limitationsas you read and think about how various mechanismsare integrated to form a functional plant.

Plant physiology is also a very active field of studyand new revelations about how plants work are reported

in the literature almost daily. Many models and expla-nations contained in this book may have been revised bythe time the book appears on the market. If you find aparticular topic interesting and wish to learn more aboutit, the listed publications at the end of each chapter areyour gateway into the relevant research literature. Youcan learn what has happened since this book was writtenby seeking out reviews and opinions published in themore recent editions of those same journals.

In spite of its presumed objectivity, scienceultimately relies on the interpretation of experimentalresults by scientists—interpretations that are oftenfound to be inadequate and filled with uncertainty.However, as results and observations accumulate,interpretations are refined and the degree of uncertaintydiminishes. This is the nature of scientific discoveryand the source of the real excitement of doing science.In this book, we have attempted to convey some senseof this scientific process.

We hope that, through this book, we are able toshare with you some of our own fascination with theexcitement, mystery, and challenge of learning aboutplant physiology.

William G. HopkinsNorman P. A. Huner

Contents

Chapter 1 • Plant Cells and Water 1

1.1 Water has Unique Physical and ChemicalProperties 2

1.2 The Thermal Properties of Water are BiologicallyImportant 3

1.2.1 Water Exhibits a Unique ThermalCapacity 3

1.2.2 Water Exhibits a High Heat of Fusion and Heat ofVaporization 3

1.3 Water is the Universal Solvent 41.4 Polarity of Water Molecules Results in Cohesion

and Adhesion 41.5 Water Movement may be Governed by Diffusion

or by Bulk Flow 5

1.5.1 Bulk Flow is Driven by HydrostaticPressure 5

1.5.2 Fick’s First Law Describes the Process ofDiffusion 5

1.6 Osmosis is the Diffusion of Water Across aSelectively Permeable Membrane 61.6.1 Plant Cells Contain an Array of Selectively

Permeable Membranes 71.6.2 Osmosis in Plant Cells is Indirectly Energy

Dependent 81.6.3 The Chemical Potential of Water has an Osmotic

as Well as a Pressure Component 91.7 Hydrostatic Pressure and Osmotic Pressure are

Two Components of Water Potential 111.8 Water Potential is the Sum of its Component

Potentials 111.9 Dynamic Flux of H2O is Associated with Changes

in Water Potential 12

1.10 Aquaporins Facilitate the Cellular Movement ofWater 13

1.11 Two-Component Sensing/Signalling Systems areInvolved in Osmoregulation 15Summary 17Chapter Review 17Further Reading 17

Chapter 2 • Whole Plant WaterRelations 19

2.1 Transpiration is Driven by Differences in VaporPressure 20

2.2 The Driving Force of Transpiration is Differencesin Vapor Pressure 21

2.3 The Rate of Transpiration is Influenced byEnvironmental Factors 222.3.1 What are the Effects of Humidity? 232.3.2 What is the Effects of Temperature? 232.3.3 What is the Effect of Wind? 24

2.4 Water Conduction Occurs via TrachearyElements 24

2.5 The Ascent of Xylem SAP is Explained byCombining Transpiration with the CohesiveForces of Water 272.5.1 Root Pressure is Related to Root

Structure 282.5.2 Water Rise by Capillarity is due to Adhesion and

Surface Tension 292.5.3 The Cohesion Theory Best Explains the Ascent of

Xylem Sap 30

vii

viii Contents

2.6 Water Loss due to Transpiration must beReplenished 332.6.1 Soil is a Complex Medium 33

2.7 Roots Absorb and Transport Water 342.8 The Permeability of Roots to Water

Varies 352.9 Radial Movement of Water Through the Root

Involves Two Possible Pathways 36Summary 37Chapter Review 37Further Reading 37

BOX 2.1 • Why Transpiration? 25

Chapter 3 • Roots, Soils, and NutrientUptake 39

3.1 The Soil as a Nutrient Reservoir 403.1.1 Colloids are a Significant Component of Most

Soils 403.1.2 Colloids Present a Large, Negatively Charged

Surface Area 403.1.3 Soil Colloids Reversibly Adsorb Cations from the

Soil Solution 413.1.4 The Anion Exchange Capacity of Soil Colloids is

Relatively Low 413.2 Nutrient Uptake 42

3.2.1 Nutrient Uptake by Plants Requires Transport ofthe Nutrient Across Root CellMembranes 42

3.2.2 Simple Diffusion is a Purely PhysicalProcess 42

3.2.3 The Movement of Most Solutes Across MembranesRequires the Participation of Specific TransportProteins 43

3.2.4 Active Transport Requires the Expenditure ofMetabolic Energy 43

3.3 Selective Accumulation of Ions by Roots 463.4 Electrochemical Gradients and Ion

Movement 463.4.1 Ions Move in Response to Electrochemical

Gradients 463.4.2 The Nernst Equation Helps to Predict Whether an

Ion is Exchanged Actively or Passively 473.5 Electrogenic Pumps are Critical for Cellular

Active Transport 493.5.1 Active Transport is Driven by ATPase-Proton

Pumps 493.5.2 The ATPase-Proton Pumps of Plasma Membranes

and Vacuolar Membranes are Different 503.5.3 K+ Exchange is Mediated by Two Classes of

Transport Proteins 51

3.6 Cellular Ion Uptake Processes areInteractive 52

3.7 Root Architecture is Important to Maximize IonUptake 523.7.1 A First Step in Mineral Uptake by Roots is

Diffusion into the Apparent Free Space 533.7.2 Apparent Free Space is Equivalent to the Apoplast

of the Root Epidermal and CorticalCells 54

3.8 The Radial Path of Ion Movement ThroughRoots 543.8.1 Ions Entering the Stele Must First be Transported

from the Apparent Free Space into theSymplast 54

3.8.2 Ions are Actively Secreted into the XylemApoplast 55

3.8.3 Emerging Secondary Roots may Contribute to theUptake of Some Solutes 55

3.9 Root-Microbe Interactions 563.9.1 Bacteria Other than Nitrogen Fixers Contribute to

Nutrient Uptake by Roots 563.9.2 Mycorrhizae are Fungi that Increase the Volume of

the Nutrient Depletion Zone AroundRoots 57

Summary 58Chapter Review 58Further Reading 59

BOX 3.1 • Electrophysiology—Exploring IonChannels 44

Chapter 4 • Plants and InorganicNutrients 61

4.1 Methods and Nutrient Solutions 624.1.1 Interest in Plant Nutrition is Rooted in the Study of

Agriculture and Crop Productivity 624.1.2 The Use of Hydroponic Culture Helped to Define

the Mineral Requirements of Plants 624.1.3 Modern Techniques Overcome Inherent

Disadvantages of Simple SolutionCulture 63

4.2 The Essential Nutrient Elements 654.2.1 Seventeen Elements are Deemed to be Essential for

Plant Growth and Development 654.2.2 The Essential Nutrients are Generally Classed as

Either Macronutrients orMicronutrients 65

4.2.3 Determining Essentiality of MicronutrientsPresents Special Problems 65

Contents ix

4.3 Beneficial Elements 664.3.1 Sodium is an Essential Micronutrient for C4

Plants 664.3.2 Silicon May be Beneficial for a Variety of

Species 674.3.3 Cobalt is Required by Nitrogen-Fixing

Bacteria 674.3.4 Some Plants Tolerate High Concentrations of

Selenium 674.4 Nutrient Functions and Deficiency

Symptoms 674.4.1 A Plant’s Requirement for a Particular Element is

Defined in Terms of CriticalConcentration 67

4.4.2 Nitrogen is a Constituent of Many CriticalMacromolecules 68

4.4.3 Phosphorous is Part of the Nucleic Acid Backboneand has a Central Function in IntermediaryMetabolism 69

4.4.4 Potassium Activates Enzymes and Functions inOsmoregulation 69

4.4.5 Sulfur is an Important Constituent of Proteins,Coenzymes, and Vitamins 70

4.4.6 Calcium is Important in Cell Division, CellAdhesion, and as a Second Messenger 70

4.4.7 Magnesium is a Constituent of the ChlorophyllMolecule and an Important Regulator of EnzymeReaction 70

4.4.8 Iron is Required for Chlorophyll Synthesisand Electron TransferReactions 71

4.4.9 Boron Appears to have a Role in CellDivision and Elongation and Contributes to theStructural Integrity of the CellWall 73

4.4.10 Copper is a Necessary Cofactor for OxidativeEnzymes 73

4.4.11 Zinc is an Activator of NumerousEnzymes 73

4.4.12 Manganese is an Enzyme Cofactor as Well asPart of the Oxygen-Evolving Complex in theChloroplast 74

4.4.13 Molybdenum is a Key Component of NitrogenMetabolism 74

4.4.14 Chlorine has a Role in Photosynthetic OxygenEvolution and Charge Balance Across CellularMembranes 74

4.4.15 The Role of Nickel is not Clear 74

4.5 Toxicity of Micronutrients 75Summary 75Chapter Review 76Further Reading 76

Chapter 5 • Bioenergetics and ATPSynthesis 77

5.1 Bioenergetics and Energy Transformations inLiving Organisms 785.1.1 The Sun is a Primary Source of Energy 785.1.2 What is Bioenergetics? 785.1.3 The First Law of Thermodynamics Refers to

Energy Conservation 795.1.4 The Second Law of Thermodynamics Refers to

Entropy and Disorder 795.1.5 The Ability to do Work is Dependent on the

Availability of Free Energy 805.1.6 Free Energy is Related to Chemical

Equilibria 805.2 Energy Transformations and Coupled

Reactions 815.2.1 Free Energy of ATP is Associated with Coupled

Phosphate Transfer Reactions 815.2.2 Free Energy Changes are Associated with Coupled

Oxidation–Reduction Reactions 835.3 Energy Transduction and the Chemiosmotic

Synthesis of ATP 855.3.1 Chloroplasts and Mitochondria Exhibit Specific

Compartments 855.3.2 Chloroplasts and Mitochondria Synthesize ATP by

Chemiosmosis 90Summary 91Chapter Review 91Further Reading 91

BOX 5.1 • Plastid Biogenesis 86

Chapter 6 • The Dual Role of Sunlight:Energy andInformation 93

6.1 The Physical Nature of Light 936.1.1 Light is Electromagnetic Energy, Which Exists in

Two Forms 936.1.2 Light can be Characterized as a Wave

Phenomenon 946.1.3 Light Can be Characterized as a Stream of Discrete

Particles 946.1.4 Light Energy can Interact with Matter 956.1.5 How Does One Illustrate the Efficiency of Light

Absorption and its PhysiologicalEffects? 97

6.1.6 Accurate Measurement of Light is Important inPhotobiology 98

6.2 The Natural Radiation Environment 996.3 Photoreceptors Absorb Light for use in a

Physiological Process 100

x Contents

6.3.1 Chlorophylls are Primarily Responsible forHarvesting Light Energy forPhotosynthesis 100

6.3.2 Phycobilins Serve as Accessory Light-HarvestingPigments in Red Algae andCyanobacteria 102

6.3.3 Carotenoids Account for the AutumnColors 103

6.3.4 Cryptochrome and Phototropin are PhotoreceptorsSensitive to Blue Light and UV-Aradiation 103

6.3.5 UV-B Radiation May Act as a DevelopmentalSignal 105

6.3.6 Flavonoids Provide the Myriad Flower Colors andAct as a Natural Sunscreen 105

6.3.7 Betacyanins and Beets 106Summary 107Chapter Review 107Further Reading 107

Chapter 7 • Energy Conservation inPhotosynthesis: HarvestingSunlight 109

7.1 Leaves are Photosynthetic Machines thatMaximize the Absorption of Light 110

7.2 Photosynthesis is an Oxidation–ReductionProcess 112

7.3 Photosynthetic Electron Transport 1147.3.1 Photosystems are Major Components of the

Photosynthetic Electron TransportChain 114

7.3.2 Photosystem II Oxidizes Water to ProduceOxygen 117

7.3.3 The Cytochrome Complex and Photosystem IOxidize Plastoquinol 119

7.4 Photophosphorylation is the Light-DependentSynthesis of ATP 120

7.5 Lateral Heterogeneity is the Unequal Distributionof Thylakoid Complexes 122

7.6 Cyanobacteria are Oxygenic 1237.7 Inhibitors of Photosynthetic Electron Transport

are Effective Herbicides 124Summary 127Chapter Review 127Further Reading 128

BOX 7.1 • Historical Perspective—The Discoveryof Photosynthesis 113

BOX 7.2 • The Case for TwoPhotosystems 125

Chapter 8 • Energy Conservation inPhotosynthesis: CO2Assimilation 129

8.1 Stomatal Complex Controls Leaf Gas Exchangeand Water Loss 130

8.2 CO2 Enters the Leaf by Diffusion 1328.3 How Do Stomata Open and Close? 1338.4 Stomatal Movements are Also Controlled by

External Environmental Factors 1358.4.1 Light and Carbon Dioxide Regulate Stomatal

Opening 1358.4.2 Stomatal Movements Follow Endogenous

Rhythms 1368.5 The Photosynthetic Carbon Reduction (PCR)

Cycle 1368.5.1 The PCR Cycle Reduces CO2 to Produce a

Three-Carbon Sugar 1378.5.2 The Carboxylation Reaction Fixes the

CO2 1378.5.3 ATP and NADPH are Consumed in the PCR

Cycle 1388.5.4 What are the Energetics of the PCR

Cycle? 1398.6 The PCR Cycle is Highly Regulated 139

8.6.1 The Regeneration of RuBP isAutocatalytic 140

8.6.2 Rubisco Activity is Regulated Indirectly byLight 140

8.6.3 Other PCR Enzymes are also Regulated byLight 141

8.7 Chloroplasts of C3 Plants also Exhibit CompetingCarbon Oxidation Processes 1428.7.1 Rubisco Catalyzes the Fixation of Both CO2 and

O2 1428.7.2 Why Photorespiration? 1438.7.3 In Addition to PCR, Chloroplasts Exhibit an

Oxidative Pentose Phosphate Cycle 145Summary 149Chapter Review 149Further Reading 150

BOX 8.1 • Enzymes 146

Chapter 9 • Allocation, Translocation, andPartitioning ofPhotoassimilates 151

9.1 Starch and Sucrose are Biosynthesized in TwoDifferent Compartments 1529.1.1 Starch is Biosynthesized in the Stroma 1529.1.2 Sucrose is Biosynthesized in the Cytosol 153

Contents xi

9.2 Starch and Sucrose Biosynthesis are CompetitiveProcesses 154

9.3 Fructan Biosynthesis is An Alternative PathwayFor Carbon Allocation 156

9.4 Photoassimilates are Translocated Over LongDistances 1569.4.1 What is the Composition of the Photoassimilate

Translocated by the Phloem? 1589.5 Sieve Elements are the Principal Cellular

Constituents of the Phloem 1599.5.1 Phloem Exudate Contains a Significant Amount of

Protein 1609.6 Direction of Translocation is Determined by

Source-Sink Relationships 1619.7 Phloem Translocation Occurs by Mass

Transfer 1619.8 Phloem Loading and Unloading Regulate

Translocation and Partitioning 1639.8.1 Phloem Loading can Occur Symplastically or

Apoplastically 1649.8.2 Phloem Unloading May Occur Symplastically or

Apoplastically 1669.9 Photoassimilate is Distributed Between

Different Metabolic Pathways and PlantOrgans 1669.9.1 Photoassimilates May be Allocated to a Variety of

Metabolic Functions in the Source or TheSink 167

9.9.2 Distribution of Photoassimilates BetweenCompeting Sinks is Determined by SinkStrength 168

9.10 Xenobiotic Agrochemicals are Translocated in thePhloem 170Summary 170Chapter Review 171Further Reading 171

Chapter 10 • Cellular Respiration:Unlocking the Energy Storedin Photoassimilates 173

10.1 Cellular Respiration Consists of a Series ofPathways by Which Photoassimilates areOxidized 174

10.2 Starch Mobilization 17510.2.1 The Hydrolytic Degradation of Starch Produces

Glucose 17510.2.2 α-Amylase Produces Maltose and Limit

Dextrins 17610.2.3 β-Amylase Produces Maltose 176

10.2.4 Limit Dextrinase is a DebranchingEnzyme 176

10.2.5 α-Glucosidase Hydrolyzes Maltose 17710.2.6 Starch Phosphorylase Catalyzes the

Phosphorolytic Degradation ofStarch 177

10.3 Fructan Mobilization is Constitutive 17810.4 Glycolysis Converts Sugars to Pyruvic

Acid 17810.4.1 Hexoses Must be Phosphorylated to Enter

Glycolysis 17810.4.2 Triose Phosphates are Oxidized to

Pyruvate 18010.5 The Oxidative Pentose Phosphate Pathway is an

Alternative Route for GlucoseMetabolism 180

10.6 The Fate of Pyruvate Depends on the Availabilityof Molecular Oxygen 181

10.7 Oxidative Respiration is Carried out by theMitochondrion 18210.7.1 In The Presence of Molecular Oxygen, Pyruvate

is Completely Oxidized to CO2 and Water bythe Citric Acid Cycle 182

10.7.2 Electrons Removed from Substrate in the CitricAcid Cycle are Passed to Molecular OxygenThrough the Mitochondrial Electron TransportChain 183

10.8 Energy is Conserved in the Form of ATP inAccordance with Chemiosmosis 185

10.9 Plants Contain Several Alternative ElectronTransport Pathways 18610.9.1 Plant Mitochondria Contain External

Dehydrogenases 18610.9.2 Plants have a Rotenone-Insensitive NADH

Dehydrogenase 18610.9.3 Plants Exhibit Cyanide-Resistant

Respiration 18710.10 Many Seeds Store Carbon as Oils that are

Converted to Sugar 18810.11 Respiration Provides Carbon Skeletons for

Biosynthesis 18910.12 Respiratory Rate Varies with Development and

Metabolic State 19110.13 Respiration Rates Respond to Environmental

Conditions 19210.13.1 Light 19210.13.2 Temperature 19210.13.3 Oxygen Availability 193Summary 193Chapter Review 194Further Reading 194

xii Contents

Chapter 11 • Nitrogen Assimilation 195

11.1 The Nitrogen Cycle: A Complex Pattern ofExchange 19511.1.1 Ammonification, Nitrification, and

Denitrification are Essential Processes in theNitrogen Cycle 196

11.2 Biological Nitrogen Fixation is ExclusivelyProkaryotic 19611.2.1 Some Nitrogen-Fixing Bacteria are Free-Living

Organisms 19611.2.2 Symbiotic Nitrogen Fixation Involves Specific

Associations Between Bacteria andPlants 197

11.3 Legumes Exhibit Symbiotic NitrogenFixation 19711.3.1 Rhizobia Infect the Host Roots, Which Induces

Nodule Development 19811.4 The Biochemistry of Nitrogen Fixation 200

11.4.1 Nitrogen Fixation is Catalyzed by the EnzymeDinitrogenase 200

11.4.2 Nitrogen Fixation is EnergeticallyCostly 201

11.4.3 Dinitrogenase is Sensitive to Oxygen 20211.4.4 Dinitrogenase Results in the Production of

Hydrogen Gas 20211.5 The Genetics of Nitrogen Fixation 203

11.5.1 NIF Genes Code for Dinitrogenase 20311.5.2 NOD Genes and NIF Genes Regulate

Nodulation 20311.5.3 What is the Source of Heme For

Leghemoglobin? 20411.6 NH3 Produced by Nitrogen Fixation is

Converted to Organic Nitrogen 20411.6.1 Ammonium is Assimilated by

GS/GOGAT 20411.6.2 PII Proteins Regulate GS/GOGAT 20511.6.3 Fixed Nitrogen is Exported as Asparagine and

Ureides 20611.7 Plants Generally Take up Nitrogen in the Form

of Nitrate 20711.8 Nitrogen Cycling: Simultaneous Import and

Export 20811.9 Agricultural and Ecosystem Productivity is

Dependent on Nitrogen Supply 209Summary 211Chapter Review 211Further Reading 211

Chapter 12 • Carbon and NitrogenAssimilation and PlantProductivity 213

12.1 Productivity Refers to an Increase inBiomass 213

12.2 Carbon Economy is Dependent on the BalanceBetween Photosynthesis andRespiration 214

12.3 Productivity is Influenced by a Variety ofEnvironmental Factors 21512.3.1 Fluence Rate 21512.3.2 Available CO2 21612.3.3 Temperature 21812.3.4 Soil Water Potential 21912.3.5 Nitrogen Supply Limits Productivity 21912.3.6 Leaf Factors 220Summary 221Chapter Review 222Further Reading 222

Chapter 13 • Responses of Plants toEnvironmental Stress 223

13.1 What is Plant Stress? 22313.2 Plants Respond to Stress in Several Different

Ways 22413.3 Too Much Light Inhibits

Photosynthesis 22513.3.1 The D1 Repair Cycle Overcomes Photodamage

to PSII 22713.4 Water Stress is a Persistent Threat to Plant

Survival 22913.4.1 Water Stress Leads to Membrane

Damage 23013.4.2 Photosynthesis is Particularly Sensitive to Water

Stress 23013.4.3 Stomata Respond to Water Deficit 230

13.5 Plants are Sensitive to Fluctuations inTemperature 23313.5.1 Many Plants are Chilling Sensitive 23313.5.2 High-Temperature Stress Causes Protein

Denaturation 23413.6 Insect Pests and Disease Represent Potential

Biotic Stresses 23513.6.1 Systemic Acquired Resistance

Represents a Plant ImmuneResponse 236

13.6.2 Jasmonates Mediate Insect and DiseaseResistance 237

13.7 There are Features Common to allStresses 237Summary 238Chapter Review 238Further Reading 238

BOX 13.1 • Monitoring Plant Stress byChlorophyll Fluorescence 228

Contents xiii

Chapter 14 • Acclimation to EnvironmentalStress 241

14.1 Plant Acclimation is a Time-DependentPhenomenon 242

14.2 Acclimation is Initiated by Rapid, Short-TermResponses 24214.2.1 State Transitions Regulate Energy Distribution

in Response to Changes in SpectralDistribution 242

14.2.2 Carotenoids Serve a Dual Function: LightHarvesting and Photoprotection 244

14.2.3 Osmotic Adjustment is a Response to WaterStress 247

14.2.4 Low Temperatures Induce Lipid Unsaturationand Cold Regulated Genes in Cold TolerantPlants 248

14.2.5 Q10 for Plant Respiration Varies as a Function ofTemperature 248

14.3 Long-Term Acclimation AltersPhenotype 24914.3.1 Light Regulates Nuclear Gene Expression and

Photoacclimation 24914.3.2 Does the Photosynthetic Apparatus Respond to

Changes in Light Quality? 25214.3.3 Acclimation to Drought Affects Shoot–Root

Ratio and Leaf Area 25314.3.4 Cold Acclimation Mimics

Photoacclimation 25414.4 Freezing Tolerance in Herbaceous Species is a

Complex Interaction Between Light and LowTemperature 25514.4.1 Cold Acclimated Plants Secrete Antifreeze

Proteins 25614.4.2 North Temperate Woody Plants Survive

Freezing Stress 25614.5 Plants Adjust Photosynthetic Capacity in

Response to High Temperature 25714.6 Oxygen may Protect During Accimation to

Various Stresses 258Summary 259Chapter Review 259Further Reading 260

Chapter 15 • Adaptations to theEnvironment 261

15.1 Sun and Shade Adapted Plants RespondDifferentially to Irradiance 262

15.2 C4 Plants are Adapted to High Temperature andDrought 26315.2.1 The C4 Syndrome is Another Biochemical

Mechanism to Assimilate CO2 263

15.2.2 The C4 Syndrome is Usually Associated withKranz Leaf Anatomy 265

15.2.3 The C4 Syndrome has EcologicalSignificance 265

15.2.4 The C4 Syndrome is Differentially Sensitive toTemperature 265

15.2.5 The C4 Syndrome is Associated with WaterStress 266

15.3 Crassulacean Acid Metabolism is an Adaptationto Desert Life 26715.3.1 Is CAM a Variation of the C4

Syndrome? 26815.3.2 CAM Plants are Particularly Suited to Dry

Habitats 26915.4 C4 and CAM Photosynthesis Require Precise

Regulation and Temporal Integration 26915.5 Plant Biomes Reflect Myriad Physiological

Adaptations 27015.5.1 Tropical Rain Forest Biomes Exhibit the

Greatest Plant Biodiversity 27015.5.2 Evapotranspiration is a Major Contributor to

Weather 27115.5.3 Desert Perennials are Adapted to Reduce

Transpiration and Heat Load 27215.5.4 Desert Annuals are Ephemeral 273Summary 273Chapter Review 274Further Reading 274

Chapter 16 • Development: AnOverview 275

16.1 Growth, Differentiation, andDevelopment 27516.1.1 Development is the Sum of Growth and

Differentiation 27516.1.2 Growth is an Irreversible Increase in

Size 27616.1.3 Differentiation Refers To Qualitative Changes

That Normally Accompany Growth 27616.2 Meristems are Centers of Plant

Growth 277

16.3 Seed Development and Germination 27916.3.1 Seeds are Formed in the Flower 27916.3.2 Seed Development and Maturation 28016.3.3 Seed Germination 28116.3.4 The Level and Activities of Various Hormones

Change Dramatically During SeedDevelopment 283

16.3.5 Many Seeds Have Additional Requirements forGermination 284

16.4 From Embryo to Adult 285

xiv Contents

16.5 Senescence and Programmed Cell Death are theFinal Stages of Development 286Summary 287Chapter Review 287Further Reading 288

BOX 16.1 • Development in a MutantWeed 282

Chapter 17 • Growth and Development ofCells 289

17.1 Growth of Plant Cells is Complicated by thePresence of a Cell Wall 28917.1.1 The Primary Cell Wall is a Network of Cellulose

Microfibrils and Cross-LinkingGlycans 289

17.1.2 The Cellulose–Glycan Lattice is Embedded in aMatrix of Pectin and Protein 290

17.1.3 Cellulose Microfibrils are Assembled at thePlasma Membrane as they are Extruded into theCell Wall 292

17.2 Cell Division 29217.2.1 The Cell Cycle 29217.2.2 Cytokinesis 29317.2.3 Plasmodesmata are Cytoplasmic Channels that

Extend Through the Wall to Connect theProtoplasts of Adjacent Cells 294

17.3 Cell Walls and Cell Growth 29417.3.1 Cell Growth is Driven by Water Uptake and

Limited by the Strength and Rigidity of the CellWall 296

17.3.2 Extension of the Cell Wall RequiresWall-Loosening Events that EnableLoad-Bearing Elements in the Wall to Yield toTurgor Pressure 296

17.3.3 Wall Loosening and Cell Expansion isStimulated by Low Ph and Expansins 297

17.3.4 In Maturing Cells, a Secondary Cell Wall isDeposited on the Inside of the PrimaryWall 298

17.4 A Continuous Stream of Signals ProvidesInformation that Plant Cells Use to ModifyDevelopment 29817.4.1 Signal Perception and Transduction 29917.4.2 The G-Protein System is a Ubiquitous Receptor

System 29917.5 Signal Transduction Includes a Diverse Array of

Second Messengers 30017.5.1 Protein Kinase-Based Signaling 30017.5.2 Phospholipid-Based Signaling 300

17.5.3 Calcium-Based Signaling 30117.5.4 Transcriptional-Based Signaling 303

17.6 There is Extensive Crosstalk Among SignalPathways 303Summary 304Chapter Review 304Further Reading 304

BOX 17.1 • Cytoskeleton 295

BOX 17.2 • Ubiquitin andProteasomes—Cleaning up UnwantedProteins 302

Chapter 18 • Hormones I: Auxins 305

18.1 The Hormone Concept in Plants 30518.2 Auxin is Distributed Throughout the

Plant 30618.3 The Principal Auxin in Plants is Indole-3-Acetic

Acid (IAA) 30718.4 IAA is Synthesized from the Amino Acid

l-Tryptophan 30918.5 Some Plants do not Require Tryptophan for IAA

Biosynthesis 31018.6 IAA may be Stored as Inactive

Conjugates 31018.7 IAA is Deactivated by Oxidation and Conjugation

with Amino Acids 31118.8 Auxin is Involved in Virtually Every Stage of

Plant Development 31118.8.1 The Principal Test for Auxins is the Stimulation

of Cell Enlargement in ExcisedTissues 311

18.8.2 Auxin Regulates VascularDifferentiation 311

18.8.3 Auxin Controls the Growth of AxillaryBuds 313

18.9 The Acid-Growth Hypothesis Explains AuxinControl of Cell Enlargement 314

18.10 Maintenance of Auxin-Induced Growth andOther Auxin Effects Requires GeneActivation 316

18.11 Many Aspects of Plant Development are Linkedto the Polar Transport of Auxin 317Summary 320Chapter Review 321Further Reading 321

BOX 18.1 • Discovering Auxin 307

BOX 18.2 • Commercial Applications ofAuxins 314

Contents xv

Chapter 19 • Hormones II:Gibberellins 323

19.1 There are a Large Number ofGibberellins 323

19.2 There are Three Principal Sites for GibberellinBiosynthesis 324

19.3 Gibberellins are Terpenes, Sharing a CorePathway with Several Other Hormones and aWide Range of Secondary Products 325

19.4 Gibberellins are Synthesized fromGeranylgeranyl Pyrophosphate(GGPP) 327

19.5 Gibberellins are Deactivated by2β-Hydroxylation 329

19.6 Growth Retardants Block the Synthesis ofGibberellins 329

19.7 Gibberellin Transport is PoorlyUnderstood 330

19.8 Gibberellins Affect Many Aspects of PlantGrowth and Development 33019.8.1 Gibberellins Stimulate Hyper-elongation of

Intact Stems, Especially in Dwarf and RosettePlants 330

19.8.2 Gibberellins Stimulate Mobilization of NutrientReserves During Germination of CerealGrains 332

19.9 Gibberellins Act by Regulating GeneExpression 333Summary 336Chapter Review 336Further Reading 337

BOX 19.1 • Discovery of Gibberellins 325

BOX 19.2 • Commercial Applications ofGibberellins 330

BOX 19.3 • Della Proteins and the GreenRevolution 335

Chapter 20 • Hormones III:Cytokinins 339

20.1 Cytokinins are Adenine Derivatives 33920.1.1 Cytokinin Biosynthesis Begins with the

Condensation of an Isopentenyl Group with theAmino Group of AdenosineMonophosphate 339

20.1.2 Cytokinins may be Deactivated by Conjugationor Oxidation 340

20.2 Cytokinins are Synthesized Primarily in the Rootand Translocated in the Xylem 341

20.3 Cytokinins are Required for CellProliferation 34320.3.1 Cytokinins Regulate Progression through the

Cell Cycle 34320.3.2 The Ratio of Cytokinin to Auxin

Controls Root and Shoot Initiation in CallusTissues and the Growth of AxillaryBuds 344

20.3.3 Crown Gall Tumors are Genetically Engineeredto Overproduce Cytokinin and Auxin 345

20.3.4 Cytokinins Delay Senescence 34620.3.5 Cytokinins Have an Important Role in

Maintaining the Shoot Meristem 34720.3.6 Cytokinin Levels in the Shoot Apical Meristem

Are Regulated by Master ControlGenes 348

20.4 Cytokinin Receptor and Signaling 35020.4.1 The Cytokinin Receptor is a Membrane-Based

Histidine Kinase 35020.4.2 The Cytokinin Signaling Chain Involves a

Multistep Transfer of Phosphoryl Groups toResponse Regulators 351

Summary 353Chapter Review 353Further Reading 354

BOX 20.1 • The Discovery of Cytokinins 341

BOX 20.2 • Tissue Culture has Made PossibleLarge-Scale Cloning of Plants byMicropropagation 345

Chapter 21 • Hormones IV: Abscisic Acid,Ethylene, andBrassinosteroids 355

21.1 Abscisic Acid 35521.1.1 Abscisic Acid is Synthesized from a Carotenoid

Precursor 35521.1.2 Abscisic Acid is Degraded to Phaseic Acid by

Oxidation 35721.1.3 Abscisic Acid is Synthesized in Mesophyll Cells,

Guard Cells, and Vascular Tissue 35721.1.4 Abscisic Acid Regulates Embryo Maturation and

Seed Germination 35821.1.5 Abscisic Acid Mediates Response to Water

Stress 35821.1.6 Other Abscisic Acid Responses 35921.1.7 ABA Perception and Signal

Transduction 359

xvi Contents

21.2 Ethylene 36221.2.1 Ethylene is Synthesized from the Amino Acid

Methionine 36221.2.2 Excess Ethylene is Subject to

Oxidation 36421.2.3 The Study of Ethylene Presents a Unique Set of

Problems 36421.2.4 Ethylene Affects Many Aspects of Vegetative

Development 36421.2.5 Ethylene Receptors and Signaling 365

21.3 Brassinosteroids 36721.3.1 Brassinosteroids are Polyhydroxylated Sterols

Derived from the TriterpeneSqualene 367

21.3.2 Several Routes for Deactivation ofBrassinosteroids have been Identified 369

21.3.3 Brassinolide receptors and Signaling 369Summary 369Chapter Review 370Further Reading 370

BOX 21.1 • The Discovery of AbscisicAcid 356

BOX 21.2 • The Discovery of Ethylene 363

BOX 21.3 • Mitogenactivated Protein Kinase: AWidespread Mechanism for SignalTransduction 366

Chapter 22 • Photomorphogenesis:Responding to Light 373

22.1 Photomorphogenesis is Initiated byPhotoreceptors 373

22.2 Phytochromes: Responding to Red and Far-RedLight 37422.2.1 Photoreversibility is the Hallmark of

Phytochrome Action 37622.2.2 Conversion of Pr to Pfr in Etiolated Seedlings

Leads to a Loss of Both Pfr and TotalPhytochrome 377

22.2.3 Light Establishes a State of DynamicPhotoequilibrium Between Pr andPfr 378

22.2.4 Phytochrome Responses can be GroupedAccording to their FluenceRequirements 378

22.3 Cryptochrome: Responding to Blue and UV-ALight 379

22.4 Phytochrome and Cryptochrome MediateNumerous Developmental Responses 37922.4.1 Seed Germination 37922.4.2 De-Etiolation 38022.4.3 Shade Avoidance 38122.4.4 Detecting End-of-day Signals 38122.4.5 Control of Anthocyanin Biosynthesis 38222.4.6 Rapid Phytochrome Responses 38222.4.7 PhyA may Function to Detect the Presence of

Light 38322.5 Chemistry and Mode of Action of Phytochrome

and Cryptochrome 38322.5.1 Phytochrome is a Phycobiliprotein 38322.5.2 Phytochrome Signal Transduction 38422.5.3 Cryptochrome Structure is Similar to DNA

Repair Enzymes 38622.5.4 Cryptochrome Signal Transduction 386

22.6 Some Plant Responses are Regulated by UV-BLight 387

22.7 De-Etiolation in Arabidopsis: A Case Study inPhotoreceptor Interactions 387Summary 388Chapter Review 389Further Reading 389

BOX 22.1 • Historical Perspectives—TheDiscovery of Phytochrome 375

Chapter 23 • Tropisms and NasticMovements: Orienting Plantsin Space 391

23.1 Phototropism: Reaching for the Sun 39223.1.1 Phototropism is a Response to a Light

Gradient 39223.1.2 Phototropism is a Blue-Light

Response 39323.1.3 Phototropism Orients a Plant for Optimal

Photosynthesis 39323.1.4 Fluence Response Curves Illustrate the

Complexity of PhototropicResponses 394

23.1.5 The Phototropic Response is Attributed to aLateral Redistribution of DiffusibleAuxin 395

23.1.6 Phototropism and Related Responses areRegulated by a Family of Blue-SensitiveFlavoproteins 396

23.1.7 A Hybrid Red/Blue Light Photoreceptor hasbeen Isolated from a Fern 397

Contents xvii

23.1.8 Phototropin Activity and SignalChain 397

23.1.9 Phototropism in Green Plants is Not WellUnderstood 398

23.2 Gravitropism 39823.2.1 Gravitropism is More than Simply Up and

Down 39923.2.2 The Gravitational Stimulus is the Product of

Intensity and Time 39923.2.3 Root Gravitropism Occurs in Four

Phases 40123.3 Nastic Movements 405

23.3.1 Nyctinastic Movements are RhythmicMovements Involving Reversible TurgorChanges 406

23.3.2 Nyctinastic Movements are due to Ion Fluxesand Resulting Osmotic Responses in SpecializedMotor Cells 407

23.3.3 Seismonasty is a Response to MechanicalStimulation 409

Summary 410Chapter Review 411Further Reading 411

BOX 23.1 • Methods in the Study ofGravitropism 400

Chapter 24 • Measuring Time: ControllingDevelopment by Photoperiodand EndogenousClocks 413

24.1 Photoperiodism 41424.1.1 Photoperiodic Responses may be Characterized

by a Variety of Response Types 41524.1.2 Critical Daylength Defines Short-Day and

Long-Day Responses 41524.1.3 Plants Actually Measure the Length of the Dark

Period 41724.1.4 Phytochrome and Cryptochrome are the

Photoreceptors for Photoperiodism 41824.1.5 The Photoperiodic Signal is Perceived by the

Leaves 41924.1.6 Control of Flowering by Photo-

period Requires a TransmissibleSignal 420

24.1.7 Photoperiodism Normally Requires a Period ofHigh Fluence Light Before or After the DarkPeriod 421

24.2 The Biological Clock 42324.2.1 Clock-Driven Rhythms Persist Under Constant

Conditions 423

24.2.2 Light Resets the Biological Clock on a DailyBasis 425

24.2.3 The Circadian Clock isTemperature-Compensated 426

24.2.4 The Circadian Clock is a Significant Componentin Photoperiodic TimeMeasurement 427

24.2.5 Daylength Measurement Involves an InteractionBetween an External Light Signal and aCircadian Rhythm 428

24.2.6 The Circadian Clock is a Negative FeedbackLoop 429

24.3 Photoperiodism in Nature 430Summary 431Chapter Review 432Further Reading 432

BOX 24.1 • Historical Perspectives: TheDiscovery of Photoperiodism 414

BOX 24.2 • Historical Perspectives: TheBiological Clock 422

Chapter 25 • Flowering and FruitDevelopment 433

25.1 Flower Initiation and Development Involves theSequential Action of Three Sets ofGenes 43325.1.1 Flowering-Time Genes Influence the Duration

of Vegetative Growth 43425.1.2 Floral-Identity Genes and Organ-Identity Genes

Overlap in Time and Function 43625.2 Temperature can Alter the Flowering Response

to Photoperiod 43725.2.1 Vernalization Occurs most Commonly in

Winter Annuals and Biennials 43825.2.2 The Effective Temperature for Vernalization is

Variable 43925.2.3 The Vernalization Treatment is Perceived by the

Shoot Apex 44025.2.4 The Vernalized State is

Transmissible 44025.2.5 Gibberellin and Vernalization Operate through

Independent Genetic Pathways 44025.2.6 Threee Genes Determine the Vernalization

Requirement in Cereals 44125.3 Fruit Set and Development is Regulated by

Hormones 44225.3.1 The Development of Fleshy Fruits can be

Divided into Five Phases 442

xviii Contents

25.3.2 Fruit Set is Triggered by Auxin 44225.3.3 Ripening is Triggered by Ethylene in

Climacteric Fruits 444Summary 445Chapter Review 446Further Reading 446

BOX 25.1 • Ethylene: It’s a Gas! 445

Chapter 26 • Temperature: PlantDevelopment andDistribution 447

26.1 Temperature in the PlantEnvironment 447

26.2 Bud Dormancy 44926.2.1 Bud Dormancy is Induced by

Photoperiod 45026.2.2 A Period of Low Temperature is Required to

Break Bud Dormancy 45126.3 Seed Dormancy 451

26.3.1 Numerous Factors Influence SeedDormancy 451

26.3.2 Temperature has a Significant Impactl on SeedDormancy 453

26.4 Thermoperiodism is a Response to AlternatingTemperature 454

26.5 Temperature Influences PlantDistribution 454Summary 457Chapter Review 457Further Reading 457

BOX 26.1 • Bulbs and Corms 450

Chapter 27 • Secondary Metabolites 459

27.1 Secondary Metabolites: A.K.A NaturalProducts 459

27.2 Terpenes 46027.2.1 The Terpenes are a Chemically and Functionally

Diverse Group of Molecules 46027.2.2 Terpenes are Constituents of Essential

Oils 46027.2.3 Steroids and Sterols are Tetracyclic

Triterpenoids 46227.2.4 Polyterpenes Include the Carotenoid Pigments

and Natural Rubber 46227.3 Glycosides 463

27.3.1 Saponins are Terpene Glycosides withDetergent Properties 464

27.3.2 Cardiac Glycosides are Highly Toxic SteroidGlycosides 465

27.3.3 Cyanogenic Glycosides are A Natural Source ofHydrogen Cyanide 466

27.3.4 Glucosinolates are Sulfur-Containing Precursorsto Mustard Oils 466

27.4 Phenylpropanoids 46727.4.1 Shikimic Acid is a Key Intermediate in the

Synthesis of Both Aromatic Amino Acids andPhenylpropanoids 468

27.4.2 The Simplest Phenolic Molecules are EssentiallyDeaminated Versions of the CorrespondingAmino Acids 468

27.4.3 Coumarins and Coumarin Derivatives Functionas Anticoagulants 468

27.4.4 Lignin is a Major Structural Component ofSecondary Cell Walls 470

27.4.5 Flavonoids and Stilbenes have ParallelBiosynthetic Pathways 471

27.4.6 Tannins Denature Proteins and Add anAstringent Taste to Foods 472

27.5 Secondary Metabolites are Active Against Insectsand Disease 47427.5.1 Some Terpenes and Isoflavones have Insecticidal

and Anti-Microbial Activity 47427.5.2 Recognizing Potential Pathogens 47527.5.3 Salicylic Acid, a Shikimic Acid Derivative,

Triggers Systemic AcquiredResistance 475

27.6 Jasmonates are Linked to Ubiquitin-RelatedProtein Degradation 476

27.7 Alkaloids 47627.7.1 Alkaloids are a Large Family of Chemically

Unrelated Molecules 47627.7.2 Alkaloids are Noted Primarily for their

Pharmacological Properties and MedicalApplications 476

27.7.3 Like Many Other Secondary Metabolites,Alkaloids Serve as Preformed Chemical DefenseMolecules 479

Summary 479Chapter Review 480Further Reading 480

Appendix • Building Blocks: Lipids, Proteins,and Carbohydrates 481

I.1 Lipids 481I.2 Proteins 483I.3 Carbohydrates 485

I.3.1 Monosaccharides 485I.3.2 Polysaccharides 486

Index/Glossary 489

Cytosol

Vacuole

PIP

TIP

H2O H2O

H2O

H2O

1Plant Cells and Water

Without water, life as we know it could not exist. Wateris the most abundant constituent of most organisms.The actual water content will vary according to tissueand cell type and it is dependent to some extenton environmental and physiological conditions, butwater typically accounts for more than 70 percent byweight of non-woody plant parts. The water contentof plants is in a continual state of flux, dependingon the level of metabolic activity, the water statusof the surrounding air and soil, and a host of otherfactors. Although certain desiccation-tolerant plantsmay experience water contents of only 20 percent anddry seeds may contain as little as 5 percent water,both are metabolically inactive, and resumption ofsignificant metabolic activity is possible only after thewater content has been restored to normal levels.

Water fills a number of important roles in thephysiology of plants; roles for which it is uniquely suitedbecause of its physical and chemical properties. Thethermal properties of water ensure that it is in the liq-uid state over the range of temperatures at which mostbiological reactions occur. This is important becausemost of these reactions can occur only in an aqueousmedium. The thermal properties of water also con-tribute to temperature regulation, helping to ensure thatplants do not cool down or heat up too rapidly. Wateralso has excellent solvent properties, making it a suit-

able medium for the uptake and distribution of mineralnutrients and other solutes required for growth. Manyof the biochemical reactions that characterize life,such as oxidation, reduction, condensation, and hydrol-ysis, occur in water and water is itself either a reactantor a product in a large number of those reactions. Thetransparency of water to visible light enables sunlightto penetrate the aqueous medium of cells where it can beused to power photosynthesis or control development.

Water in land plants is part of a very dynamic sys-tem. Plants that are actively carrying out photosynthesisexperience substantial water loss, largely through evap-oration from the leaf surfaces. Equally large quantitiesof water must therefore be taken up from the soil andmoved through the plant in order to satisfy deficienciesthat develop in the leaves. For example, it is estimatedthat the turnover of water in plants due to photosynthesisand transpiration is about 1011 tonnes per year.

This constant flow of water through plants is amatter of considerable significance to their growth andsurvival. The uptake of water by cells generates a pres-sure known as turgor; in the absence of any skeletalsystem, plants must maintain cell turgor in order toremain erect. As will be shown in later chapters, theuptake of water by cells is also the driving force forcell enlargement. Few plants can survive desiccation.There is no doubt that the water relations of plants and

1

2 Chapter 1 / Plant Cells and Water

plant cells are fundamental to an understanding of theirphysiology.

This chapter is concerned with the water relationsof cells. Topics to be addressed include the following:

• a review of the unique physical and chemical prop-erties of water that make it particularly suitable as amedium for life,

• physical processes that underlie water movement inplants, including diffusion, osmosis, and bulk flowas mechanisms for water movement, and

• the chemical potential of water and the concept ofwater potential.

These concepts provide the basis for understandingwater movement within the plant and between the plantand its environment, to be discussed in Chapter 2.

1.1 WATER HAS UNIQUEPHYSICAL AND CHEMICALPROPERTIES

The key to understanding many of the unique propertiesof water is found in the structure of the water moleculeand the strong intermolecular attractions that resultfrom that structure. Water consists of an oxygen atomcovalently bonded to two hydrogen atoms (Figure 1.1).The oxygen atom is strongly electronegative, whichmeans that it has a tendency to attract electrons. Oneconsequence of this strong electronegativity is that, inthe water molecule, the oxygen tends to draw electronsaway from the hydrogen. The shared electrons thatmake up the O—H bond are, on the average, closer tothe oxygen nucleus than to hydrogen. As a consequence,the oxygen atom carries a partial negative charge, and acorresponding partial positive charge is shared betweenthe two hydrogen atoms. This asymmetric electron

A.

O

HH

+ +

B.

O

O

O

O

O

H

HH

H

H

H

HH

A.

O

HH

+ +

− −

B.

O

O

O

O

O

H

HH

H

H

H

HH

FIGURE 1.1 (A) Schematic structure of a water molecule.(B) The hydrogen bond (dashed line) results from theelectrostatic attraction between the partial positivecharge on one molecule and the partial negative chargeon the next.

distribution makes water a polar molecule. Overall,water remains a neutral molecule, but the separation ofpartial negative and positive charges generates a strongmutual (electrical) attraction between adjacent water

TABLE 1.1 Some physical properties of water compared with other molecules of similar molecular size.Because thermal properties are defined on an energy-per-unit mass basis, values are given in units of joulesper gram

Specific Melting Heat of Boiling Heat ofMolecular heat point fusion point vaporizationmass (Da) (J/g/˚C) (˚C) (J/g) (˚C) (J/g)

Water 18 4.2 0 335 100 2452Hydrogen sulphide 34 — −86 70 −61 —Ammonia 17 5.0 −77 452 −33 1234Carbon dioxide 44 — −57 180 −78 301Methane 16 — −182 58 −164 556Ethane 30 — −183 96 −88 523Methanol 32 2.6 −94 100 65 1226Ethanol 46 2.4 −117 109 78 878

1.2 The Thermal Properties of Water are Biologically Important 3

molecules or between water and other polar molecules.This attraction is called hydrogen bonding (Figure 1.1).The energy of the hydrogen bond is about 20 kJ mol−1.The hydrogen bond is thus weaker than either covalentor ionic bonds, which typically measure several hundredkJ mol−1, but stronger than the short-range, transientattractions known as Van der Waals forces (about 4 kJmol−1). Hydrogen bonding is largely responsible for themany unique properties of water, compared with othermolecules of similar molecular size (Table 1.1).

In addition to interactions between water molecules,hydrogen bonding also accounts for attractions betweenwater and other molecules or surfaces. Hydrogen bond-ing, for example, is the basis for hydration shells thatform around biologically important macromoleculessuch as proteins, nucleic acids, and carbohydrates.These layers of tightly bound and highly oriented watermolecules are often referred to as bound water. Ithas been estimated that bound water may account foras much as 30 percent by weight of hydrated proteinmolecules. Bound water is important to the stability ofprotein molecules. Bound water ‘‘cushions’’ protein,preventing the molecules from approaching closeenough to form aggregates large enough to precipitate.

Hydrogen bonding, although characteristic ofwater, is not limited to water. It arises whereverhydrogen is found between electronegative centers.This includes alcohols, which can form hydrogen bondsbecause of the—OH group, and macromolecules suchas proteins and nucleic acids where hydrogen bonds

between amino (−NH2) and carbonyl (|

C|

= O) groups

help to stabilize structure.

1.2 THE THERMAL PROPERTIESOF WATER AREBIOLOGICALLY IMPORTANT

Perhaps the single most important property of water isthat it is a liquid over the range of temperatures mostcompatible with life. Boiling and melting points aregenerally related to molecular size, such that changesof state for smaller molecules occur at lower temper-atures than for larger molecules. On the basis of sizealone, water might be expected to exist primarily inthe vapor state at temperatures encountered over mostof the earth. However, both the melting and boilingpoints of water are higher than expected when comparedwith other molecules of similar size, especially ammonia(NH3) and methane (CH4) (Table 1.1). Molecules suchas ammonia and the hydrocarbons (methane and ethane)are associated only through weak Van der Waals forcesand relatively little energy is required to change theirstate. Note, however, that the introduction of oxygenraises the boiling points of both methanol (CH3 —OH)

and ethanol (CH3CH2OH) to temperatures much closerto that of water. This is because the presence of oxy-gen introduces polarity and the opportunity to formhydrogen bonds.

1.2.1 WATER EXHIBITS A UNIQUETHERMAL CAPACITY

The term specific heat1 is used to describe the thermalcapacity of a substance or the amount of energy that canbe absorbed for a given temperature rise. The specificheat of water is 4.184 J g−1 ◦C−1, higher than that ofany other substance except liquid ammonia (Table 1.1).Because of its highly ordered structure, liquid water alsohas a high thermal conductivity. This means that itrapidly conducts heat away from the point of application.The combination of high specific heat and thermal con-ductivity enables water to absorb and redistribute largeamounts of heat energy without correspondingly largeincreases in temperature. For plant tissues that consistlargely of water, this property provides for an excep-tionally high degree of temperature stability. Localizedoverheating in a cell due to the heat of biochemicalreactions is largely prevented because the heat maybe quickly dissipated throughout the cell. In addition,large amounts of heat can be exchanged between cellsand their environment without extreme variation in theinternal temperature of the cell.

1.2.2 WATER EXHIBITS A HIGHHEAT OF FUSION AND HEATOF VAPORIZATION

Energy is required to cause changes in the state of anysubstance, such as from solid to liquid or liquid to gas,without a change in temperature. The energy requiredto convert a substance from the solid to the liquid stateis known as the heat of fusion. The heat of fusion forwater is 335 J g−1, which means that 335 J of energy arerequired to convert 1 gram of ice to 1 gram of liquidwater at 0◦C (Table 1.1). Expressed on a molar basis,the heat of fusion of water is 6.0 kJ mol−1 (18 g of waterper mole × 335 J g−1). The heat of fusion of water isone of the highest known, second only to ammonia.The high heat of fusion of water is attributable tothe large amount of energy necessary to overcome the

1Specific heat is defined as the amount of energy required toraise the temperature of one gram of substance by 1◦C(usually at 20◦C). The specific heat of water is the basis forthe definition of a quantity of energy called the calorie. Thespecific heat of water was therefore assigned the value of 1.0calorie. In accordance with the International System of Units(Systeme Internationale d’Unites, or SI), the preferred unitfor energy is the joule (J). 1 calorie = 4.184 joules.

4 Chapter 1 / Plant Cells and Water

strong intermolecular forces associated with hydrogenbonding.

The density of ice is another important property.At 0◦C, the density of ice is less than that of liquidwater. Thus water, unlike other substances, reachesits maximum density in the liquid state (near 4◦C),rather than as a solid. This occurs because moleculesin the liquid state are able to pack more tightly than inthe highly ordered crystalline state of ice. Consequently,ice floats on the surface of lakes and ponds ratherthan sinking to the bottom where it might remainyear-round. This is extremely important to the survivalof aquatic organisms of all kinds.

Just as hydrogen bonding increases the amount ofenergy required to melt ice, it also increases the energyrequired to evaporate water. The heat of vaporizationof water, or the energy required to convert one mole ofliquid water to one mole of water vapor, is about 44 kJmol−1 at 25◦C. Because this energy must be absorbedfrom its surroundings, the heat of vaporization accountsfor the pronounced cooling effect associated with evap-oration. Evaporation from the moist surface cools thesurface because the most energetic molecules escape thesurface, leaving behind the lower-energy (hence, cooler)molecules. As a result, plants may undergo substantialheat loss as water evaporates from the surfaces of leafcells. Such heat loss is an important mechanism for tem-perature regulation in the leaves of terrestrial plants thatare often exposed to intense sunlight.

1.3 WATER IS THE UNIVERSALSOLVENT

The excellent solvent properties of water are due to thehighly polar character of the water molecule. Water has

the ability to partially neutralize electrical attractionsbetween charged solute molecules or ions by surround-ing the ion or molecule with one or more layers oforiented water molecules, called a hydration shell.Hydration shells encourage solvation by reducing theprobability that ions can recombine and form crystalstructures (Figure 1.2).

The polarity of molecules can be measured bya quantity known as the dielectric constant. Waterhas one of the highest known dielectric constants(Table 1.2). The dielectric constants of alcohols aresomewhat lower, and those of nonpolar organic liquidssuch as benzene and hexane are very low. Water isthus an excellent solvent for charged ions or molecules,which dissolve very poorly in nonpolar organic liquids.Many of the solutes of importance to plants are charged.On the other hand, the low dielectric constants ofnonpolar molecules helps to explain why chargedsolutes do not readily cross the predominantly nonpolar,hydrophobic lipid regions of cellular membranes.

1.4 POLARITY OF WATERMOLECULES RESULTS INCOHESION AND ADHESION

The strong mutual attraction between water moleculesresulting from hydrogen bonding is also known as cohe-sion. One consequence of cohesion is that water hasan exceptionally high surface tension, which is mostevident at interfaces between water and air. Surface ten-sion arises because the cohesive force between watermolecules is much stronger than interactions betweenwater and air. The result is that water molecules at thesurface are constantly being pulled into the bulk water(Figure 1.3). The surface thus tends to contract andbehaves much in the manner of an elastic membrane. A

H

Na+

OO

OO

O

OO

OO

O

O

O

O

O

H

H

H

H

H

H

H H

H

HH

H

H H

H

HH

H

H

HH

H

H

H

H

H

HH

Cl-

O

H

FIGURE 1.2 Solvent properties of water. The orientation of water molecules aroundthe sodium and chloride ions screens the local electrical fields around each ion. Thescreening effect reduces the probability of the ions reuniting to form a crystallinestructure.

1.5 Water Movement may be Governed by Diffusion or by Bulk Flow 5

TABLE 1.2 Dielectric constants for somecommon solvents at 25◦C

Water 78.4Methanol 33.6Ethanol 24.3Benzene 2.3Hexane 1.9

high surface tension is the reason water drops tend to bespherical or that a water surface will support the weightof small insects.

Cohesion is directly responsible for the unusuallyhigh tensile strength of water. Tensile strength is themaximum tension that an uninterrupted column of anymaterial can withstand without breaking. High tensilestrength is normally associated with metals but, underthe appropriate conditions, water columns are also capa-ble of withstanding extraordinarily high tensions—onthe order of 30 megapascals (MPa).2

The same forces that attract water molecules to eachother will also attract water to solid surfaces, a processknown as adhesion. Adhesion is an important factor inthe capillary rise of water in small-diameter conduits.

The combined properties of cohesion, adhesion,and tensile strength help to explain why water risesin capillary tubes and are exceptionally important inmaintaining the continuity of water columns in plants.Cohesion, adhesion, and tensile strength will be dis-cussed in greater detail in Chapter 2, when evaporativewater loss from plants and water movement in the xylemare examined.

Air

Water

FIGURE 1.3 Schematic demonstration of surface ten-sion in a water drop. Intermolecular attractions betweenneighboring water molecules (heavy arrows) are greaterthan attractions between water and air (light arrows),thus tending to pull water molecules at the surface intothe bulk water.

2The pascal (Pa), equal to a force of 1 newton per squaremeter, is the standard SI unit for pressure.

MPa = Pa × 106.

1.5 WATER MOVEMENT MAY BEGOVERNED BY DIFFUSIONOR BY BULK FLOW

One objective of plant physiology is to understand thedynamics of water as it flows into and out of cells orfrom the soil, through the plant, into the atmosphere.Movement of substances from one region to another iscommonly referred to as translocation. Mechanisms fortranslocation may be classified as either active or passive,depending on whether metabolic energy is expendedin the process. It is sometimes difficult to distinguishbetween active and passive transport, but the transloca-tion of water is clearly a passive process. Although inthe past many scientists argued for an active component,the evidence indicates that water movement in plantsmay be indirectly dependent upon on expenditure ofmetabolic energy. Passive movement of most substancescan be accounted for by one of two physical processes:either bulk flow or diffusion. In the case of water, aspecial case of diffusion known as osmosis must also betaken into account.

1.5.1 BULK FLOW IS DRIVEN BYHYDROSTATIC PRESSURE

Movement of materials by bulk flow (or mass flow) ispressure-driven. Bulk flow occurs when an externalforce, such as gravity or pressure, is applied. As a result,all of the molecules of the substance move in a mass.Movement of water by bulk flow is a part of our everydayexperience. Water in a stream flows in response to thehydrostatic pressure established by gravity. It flows fromthe faucet in the home or workplace because of pres-sure generated by gravity acting on standing columnsof water in the municipal water tower. Bulk flow alsoaccounts for some water movement in plants, such asthrough the conducting cells of xylem tissue or themovement of water into roots. In Chapter 9, we discusshow bulk flow is a major component of the most widelyaccepted hypothesis for transport of solutes through thevascular tissue.

1.5.2 FICK’S FIRST LAW DESCRIBESTHE PROCESS OF DIFFUSION

Like bulk flow, diffusion is also a part of our everydayexperience. When a small amount of sugar is placedin a cup of hot drink, the sweetness soon becomesdispersed throughout the cup. The scent of perfumefrom a bottle opened in the corner of a room will soonbecome uniformly distributed throughout the air. Ifthe drink is not stirred and there are no mass move-ments of air in the room, the distribution of thesesubstances occurs by diffusion. Diffusion can be inter-preted as a directed movement from a region of a high

6 Chapter 1 / Plant Cells and Water

concentration to a region of lower concentration, butit is accomplished through the random thermal motionof individual molecules (Figure 1.4). Thus, while bulkflow is pressure-driven, diffusion is driven principallyby concentration differences. Diffusion is a significantfactor in the uptake and distribution of water, gases, andsolutes throughout the plant. In particular, diffusion is animportant factor in the supply of carbon dioxide for pho-tosynthesis as well as the loss of water vapor from leaves.

The process of diffusion was first examined quan-titatively by A. Fick. Fick’s first law, formulated in1855, forms the basis for the modern-day quantitativedescription of the process.

J = −D · A · �C · l−1 (1.1)

J is the flux or the amount of material crossing a unitarea per unit time (for example, mol m−2 s−1). D is thediffusion coefficient, a proportionality constant that isa function of the diffusing molecule and the mediumthrough which it travels. A and l are the cross-sectionalarea and the length of the diffusion path, respectively.The term �C represents the difference in concentrationbetween the two regions, also known as the concen-tration gradient. �C is the driving force for simplediffusion. In the particular case of gaseous diffusion, itis more convenient to use the difference in density (gmm−3) or vapor pressure (KPa, kilopascal) in place ofconcentration. The negative sign in Fick’s law accountsfor the fact that diffusion is toward the lower concentra-tion or vapor pressure. In summary, Fick’s law tells usthat the rate of diffusion is directly proportional to thecross-sectional area of the diffusion path and to the con-centration or vapor pressure gradient, and it is inverselyproportional to the length of the diffusion path.

1.6 OSMOSIS IS THE DIFFUSIONOF WATER ACROSS ASELECTIVELY PERMEABLEMEMBRANE

Fick’s law is most readily applicable to the diffusion ofsolutes and gases. In the general model illustrated inFigure 1.4, for example, the diffusing molecules couldbe glucose dissolved in water, carbon dioxide dissolvedin water, or carbon dioxide in air. We note that thediffusion of solute molecules from chamber A to cham-ber B from the time t0 to t1 does not affect the volumein either chamber, as indicated by no difference in theheight of the liquid in the columns of chambers A andB (Figure 1.4). While Fick’s law theoretically applies tothe diffusion of solvent molecules as well, it can at firstbe difficult to imagine a situation in which diffusion ofsolvent molecules could occur. Consider what wouldhappen if, for example, water were added to one of thechambers in Figure 1.4. As soon as the water level in the

t0

Solution Pure solvent

A B

A B

t1

FIGURE 1.4 Diffusion in solutions is usually associatedwith the directed movement of a solute molecule from aregion of high concentration to a region of lower concen-tration, due to the random thermal motion of the solutemolecules. Initially at time 0 (t0), there is a much higherprobability that a solute molecule in chamber A will passthrough the open window into chamber B. After a certaintime (t1), the number of solute molecules in chamber Bwill increase and the number in chamber A will decrease.This will continue until the molecules are uniformlydistributed between the two chambers. At that point,the probability of solute molecules passing between thechambers in either direction will be equal and net diffu-sion will cease. Note, the dotted line indicates that thereis no change in volume in either chamber A or B, due tothe diffusion of the solute molecule.

first chamber reached the open window, it would flowover into the second chamber—an example of bulk flow.

Alternatively, suppose that we separated chambersA and B by a selectively permeable membrane that

1.6 Osmosis is the Diffusion of Water Across a Selectively Permeable Membrane 7

allowed solvent molecules to pass freely between thetwo chambers but not the solute molecules (Figure 1.5).At time t0, the heights of the liquids in the columns ofchambers A and B are the same, indicating comparablevolumes. However, after time t1, the height of the liquidin the column of chamber A has increased while theheight of the liquid in the column of chamber B hasdecreased, indicating an increased volume in chamber Aand a decreased volume in chamber B. The differencein the height of the columns (�h) is a measure ofthe difference in volume between the two chambersseparated by a selectively permeable membrane. Theincreased volume in chamber A is due to the diffusion ofsolvent (water) from chamber B to chamber A. Diffusionof water, a process known as osmosis, will occur onlywhen the two chambers are separated from one anotherby a selectively permeable membrane. A selectivelypermeable membrane allows virtually free passage ofwater and certain small molecules, but restricts themovement of large solute molecules. Thus, all cellularmembranes are selectively permeable. Osmosis, then, issimply a special case of diffusion through a selectivelypermeable membrane.

1.6.1 PLANT CELLS CONTAIN ANARRAY OF SELECTIVELYPERMEABLE MEMBRANES

Although plants, like all multicellular organisms, exhibita wide variation in cellular morphology and function,these disparate cells are, in fact, remarkably alike. Allcells are built according to a common basic plan andat least start out with the same fundamental structures.In its simplest form, a cell is an aqueous solution ofchemicals called protoplasm surrounded by a plasmamembrane. The membrane and the protoplasm it con-tains are collectively referred to as a protoplast. Ofcourse, all of the components that make up protoplasmhave important roles to play in the life of a cell, butthe plasma membrane is particularly significant becauseit represents the boundary between the living and non-living worlds. The plasma membrane is also selectivelypermeable, which means that it allows some materialsto pass through but not others. The plasma membranethus not only physically defines the limits of a cell; it alsocontrols the exchange of material and serves to maintainessential differences between the cell and its environ-ment. The plant protoplast is, in turn, surrounded by acell wall. The cell wall defines the shape of the cell and,through adhesion to the walls of adjacent cells, providessupport for the plant as a whole.

In an electron micrograph (an image seen throughthe electron microscope), membranes are a singularlyprominent feature (Figure 1.6). In addition to the plasmamembrane, other membranes are found throughout the

t0

Solution Pure solvent

A

A

B

A B

t1

B

Δh

FIGURE 1.5 Osmosis is the directed movement of the sol-vent molecule (usually water) across a selectively perme-able membrane. Chamber A is separated from chamberB by a selectively permeable membrane. The selectivelypermeable membrane allows the free movement of thesolvent (water) molecules between chambers A and B,but restricts the movement of the solute molecules. Attime zero (t0), all the solute molecules are retained inchamber A and chambers A and B exhibit identical vol-umes, as indicated by the broken line. After a certaintime t1, all solute molecules are still retained in cham-ber A, but the volume of chamber A has increased whilethe volume in chamber B has decreased due to the diffu-sion of water across the selectively permeable membranefrom chamber B to chamber A. This change in volume isrepresented by �h.

protoplast where they form a variety of subcellular struc-tures called organelles (‘‘little organs’’). Organellesserve to compartmentalize major activities within thecell. For example, photosynthesis (Chapters 7 and 8)is localized to the chloroplasts whereas respiration

8 Chapter 1 / Plant Cells and Water

Cellwall

Mitochondrion

Mitochondrion

Peroxisome

Chloroplast

Chloroplast

Intercellularspace

Tonoplast

Centralvacuole

Nucleus

Plasma membrane

Ribosomesin thecytoplasm

FIGURE 1.6 The plant cell. A mature mesophyll cell froma Coleus leaf, as seen in the electron microscope. Notethe prominent, large central vacuole surrounded by thetonoplast, chloroplasts, and mitochondria. (Electronmicrograph by Wm. P. Wergin, courtesy of E. H. New-comb, University of Wisconsin–Madison)

(Chapter 10) is localized to the mitochondria. The cen-tral vacuole and its surrounding tonoplast membrane iscritical in the regulation of the osmotic properties of thecytoplasm.

1.6.2 OSMOSIS IN PLANT CELLS ISINDIRECTLY ENERGYDEPENDENT

Water, like any other substance, will only move down anenergy gradient—that is, when there is a difference inthe energy of water in two parts of a system. In the caseof the Figure 1.5, water initially moves from chamberB to chamber A because the energy of pure water inchamber B is greater than the energy of the water in thesolution in chamber A. Net movement of water stopswhen there is no longer an energy gradient across theselectively permeable membrane. Why is the energy ofpure water in chamber B of Figure 1.5 greater than theenergy of the water in the solution present in chamberA? The energy content of water, like any substance, ismost easily described in terms of its chemical potential.Chemical potential (μ) is defined as the free energy permole of that substance and is a measure of the capacity ofa substance to react or move. The rule is that osmosis occurs

only when there is a difference in the chemical potential (�μ)of water on two sides of a selectively permeable membrane. Inother words, osmosis occurs only when the molar freeenergy of water, that is, the chemical potential of water(μw) on one side of a selectively permeable membrane,exceeds the molar free energy or chemical potential ofwater on the other side of the same selectively permeablemembrane.

It appears that the dissolution of a solute in water insome way affects the chemical potential and hence thefree energy of the solvent water molecules. Why is thisso? This effect is due to the fact that increasing soluteconcentration in an aqueous solution decreases the molefraction of water in the solution. The mole fraction ofwater (Xw) in a solution can be represented as

Xw = w/w + s (1.2)where w represents the moles of H2O in a given volumeand s the moles of solute in the same volume of solution.Thus, as one increases s at a constant value of w, themole fraction of water, Xw, decreases. Remember thatthe concentration of pure water is 55.5 moles L−1.The chemical potential of water (μw) is related to themole fraction of water (Xw) according to the followingequation

μw = μw∗ + RT ln Xw (1.3)

where μw* is the chemical potential of pure water understandard temperature and pressure, R is the universalgas constant, and T is the absolute temperature. Thisequation tells us that as the mole fraction of waterdecreases, the chemical potential and hence the molarfree energy of water decreases. Therefore, the higherthe solute concentration of an aqueous solution, thelower the chemical potential of the solvent water. Sinceenergy flows ‘‘downhill’’ spontaneously, water flows ordiffuses spontaneously from chamber B to chamber Adue to the difference in the chemical potential of waterbetween the two chambers separated by a selectivelypermeable membrane (Figure 1.5).

Plant cells control the movement of water in andout of cells by altering the solute concentration of thecytosol relative to the solution external to the cell. Oneway to accomplish this is to regulate transport of ionsacross the cell membrane into or out of the cell. Forexample, root cells can take up nitrate ions NO−

3 fromthe soil by active transport to create a NO−

3 ion gradientacross the cell membrane (Figure 1.7) such that

(μnitrate)in > (μnitrate)out (1.4)

where μnitrate is the chemical potential of NO−3 . There-

fore, it follows that�μnitrate = (μnitrate)in − (μnitrate)out (1.5)

Since �μnitrate > 0, the uptake of NO−3 is an active

transport process that requires an input of energy. Con-comitantly, the increase in cytosolic NO−

3 decreases themole fraction of water in the cytosol relative to the mole

1.6 Osmosis is the Diffusion of Water Across a Selectively Permeable Membrane 9

H2OH2O

NO3–NO3

–

Out In

Cel

l mem

bran

e

Cel

l wal

l

Active

Passive

CYTOSOLSOIL

FIGURE 1.7 Water movement in plant cells requiressolute gradients. For example, soil nitrate (NO−

3 ) isactively transported across the selectively permeable cellmembrane into plant root cells. Thus, the establish-ment of a solute gradient such that chemical potentialof nitrate is higher in the cytosol than in the soil waterrequires an input of metabolic energy. The higher con-centration of nitrate in the cytosol than the soil wateralso establishes a gradient for the chemical potential ofwater such that the chemical potential of the soil wateris greater than the chemical potential of cytosolic water.As a consequence, water diffuses passively from the soilacross the cell membrane into the cell.

fraction of water in the soil water. Therefore, it followsthat

�μw = (μw)in − (μw)out (1.6)

Since �μw < 0, water will diffuse spontaneously intothe cell (Figure 1.7).

The chemical potential of water (μw) is particularlyuseful in the study of cellular and plant water relationsbecause it defines the amount of work that can be doneby water in one location, for example, a cell or vacuole,compared with pure water at atmospheric pressure andthe same temperature. More importantly, a differencein μw between two locations means that water is not inequilibrium and there will be a tendency for water toflow toward the location with the lower value. At thispoint, then, it can be said that the driving force for watermovement is a gradient in chemical potential. Clearly,the tendency for water to flow from a location where theμw is high to a location where the μw is lower will occurspontaneously and does not require energy because�μw < 0 (Figure 1.7). However, the only way plantcells can regulate the movement of water is throughthe establishment of solute concentration gradientsacross the selectively permeable cell membrane; thatis, the movement of water may be coupled to solutetransport (Figure 1.7). Although the flow of water ispassive (�μw < 0), the movement of solutes requires

active transport, which is energy dependent (�μnitrate> 0). Thus, osmosis is indirectly dependent on energythrough the requirement of solute concentration gradi-ents that alter the chemical potential of water. Since thecell membrane is permeable only to water (Figure 1.7),only the water diffuses freely across the cell membrane.The process of regulating the water status of a plantcell by the accumulation of solutes is known as osmoticadjustment, which is an energy-requiring process.

1.6.3 THE CHEMICAL POTENTIAL OFWATER HAS AN OSMOTIC ASWELL AS A PRESSURECOMPONENT

Osmosis can be easily demonstrated using a deviceknown as an osmometer, constructed by closing off theopen end of a thistle tube with a selectively permeablemembrane (Figure 1.8). If the tube is then filled with asugar solution and inverted in a volume of pure water,the volume of solution in the tube will increase overtime. The increase in volume is due to a net diffusion ofwater across the membrane into the solution.

The increase in the volume of the solution willcontinue until the hydrostatic pressure developed in the

h

Pressure

Membrane

A. B.

FIGURE 1.8 A demonstration of hydrostatic pressure. Aselectively permeable membrane is stretched across theend of a thistle tube containing a sucrose solution andthe tube is inverted in a container of pure water. Initially,water will diffuse across the membrane in response toa chemical potential gradient. Diffusion will continueuntil the force tending to drive water into the tube isbalanced by (A) the force generated by the hydrostatichead (h) in the tube or (B) the pressure applied by thepiston. When the two forces are balanced, the system hasachieved equilibrium and no further net movement ofwater will occur.

10 Chapter 1 / Plant Cells and Water

tube is sufficient to balance the force driving the waterinto the solution (Figure 1.8A). Alternatively, the tubecould be fitted with a piston that would allow us tomeasure the amount of force required to just preventany increase in the volume of solution (Figure 1.8B).This force, measured in units of pressure (force perunit area), is known as osmotic pressure (symbol =π; pi). The magnitude of the osmotic pressure thatdevelops is a function of solute concentration, at leastin dilute solutions. It is useful to note that an isolatedsolution cannot have an osmotic pressure. It has onlythe potential to manifest a pressure when placed in anosmometer. For this reason, we say that the solutionhas an osmotic potential (symbol = �S). It is conven-tion to define osmotic potential as the negative of theosmotic pressure, since they are equal but opposite forces(�S = − π).

Whereas simple diffusion is driven entirely by dif-ferences in mole fractions of the solvent, it is apparentfrom Figure 1.8 that pressure is also a factor in deter-mining both the direction and rate of water movementin an osmometer. Sufficient pressure applied to the pis-ton (Figure 1.8B) will prevent further net movementof water into the solution. If additional pressure wereapplied, we might expect the net movement of water toreverse its direction and instead flow out of the solution.Thus, osmosis is driven not only by the mole fraction ofdissolved solute but by pressure differences as well. Bothof these factors influence the overall chemical potentialof water, which is the ultimate driving force for watermovement in plants.

The effect of pressure on chemical potential isrepresented by the value VP. V is the partial molalvolume, or the volume occupied by one mole of thechemical species. Since one liter (= 10−3 m3) of watercontains 55.5 moles, Vw = (1000/55.5) or 18 ml per mole(= 18000 mm3 mol−1) of water. P is the pressure. Mea-surements in plant physiology are commonly made atatmospheric pressure but the presence of relatively rigidcell walls allows plant cells to develop significant hydro-static pressures. Hence it is convention to express Pas the difference between actual pressure and atmo-spheric pressure. The influence of pressure on chemicalpotential can now be added to equation 1.3:

μw = μw∗ + RT ln Xw + VwP (1.7)

The chemical potential of water may also be influencedby electrical potential and gravitational field. In spiteof its strong dipole nature, the net electrical charge forwater is zero and so the electrical term can be ignored.This is why we talk about the chemical potential ofwater rather than its electrochemical potential. While thegravitational term may be large and must be consideredwhere water movement in tall trees is concerned, itis not significant at the cellular level. Where water

movement involves heights of 5 to 10 meters or less, thegravitational term is commonly omitted.

Since μw = μw* under very dilute conditions, itfollows from Equation 1.7 that

PVw = −RT ln Xw (1.8)

Substituting π for P, we obtain the formal definition ofosmotic pressure (Equation 1.10):

πVw = −RT ln Xw (1.9)

or

π = −RT ln XwVw−1 (1.10)

As a matter of convenience, the value of Xw for purewater is arbitrarily designated as unity (= 1). Conse-quently, Xw for a solution is always less than 1. Thus,equation 1.10 tells us that addition of solute decreasesthe value of Xw, which in turn increases the osmoticpressure, π. By substituting equation 1.9 in equation 1.7,we can now arrive at an expression relating the chemi-cal potential of water, hydrostatic pressure and osmoticpressure:

μw = μw∗ + Vw(P − π) (1.11)

According to equation 1.11, the extent to which thechemical potential of water in a solution (μw) differsfrom that of pure water under conditions of standardtemperature and pressure (μ∗

w) is a function of an osmoticcomponent (π) and a pressure component (P).

Osmotic pressure or osmotic potential is one ofthe four colligative properties of a solution. Since themagnitude of the osmotic pressure or osmotic potentialexhibited by any solution is dependent on the number ofparticles present in solution, osmotic pressure (π) andtherefore osmotic potential (�S) is a colligative property.Thus, a solution of 0.5 M CaCl2 (Equation 1.12) willexhibit a greater osmotic pressure than a solution of0.5 M NaCl (Equation 1.13) because the former solutionwill exhibit twice as many Cl− ions present as the latterat the same concentration.

CaCl2 → Ca2+ + 2Cl− (1.12)NaCl → Na+ + Cl− (1.13)

The other three colligative properties of solutions arefreezing point depression, boiling point elevation, andthe lowering of the vapor pressure of the solvent.

Because π is directly proportional to the solutemolal concentration (moles of solute in 1 L of solvent),π can be used to calculate the molecular mass of thesolute molecule. According to the Ideal Gas Law,

PV = nRT (1.14)

where P is the pressure, V is the volume, n is the molesof the gaseous molecule, R is the universal gas constant,and T is the absolute temperature. Thus, for an ideal,dilute solution

πV = nRT (1.15)

1.8 Water Potential is the Sum of its Component Potentials 11

or

π = nRT V−1 (1.16)

where π is the osmotic pressure of the solution, V isthe volume, n is the moles of solute in the solution,R is the universal gas constant, and T is the absolutetemperature. Since n = mass (m)/molecular mass (M),it follows that

π = m M−1RT V−1 (1.17)

Thus, by knowing the mass of the solute (m) dissolvedin one liter (V−1), one can measure π experimentally forthis solution and subsequently calculate the molecularmass of the solute. Furthermore, if one knows themolecular mass of the solute, equation 1.17 allowsone to calculate the osmotic pressure (π) and hence theosmotic potential of a solution. Note that the solutemust be perfectly soluble in water in order to use thismethod to determine either the molecular mass of thesolute or the osmotic pressure of the solution.

1.7 HYDROSTATIC PRESSUREAND OSMOTIC PRESSUREARE TWO COMPONENTS OFWATER POTENTIAL

It is usually more convenient to measure relative valuesthan it is to measure absolute values. The absolutechemical potential of water in solutions is one of thosequantities that is not conveniently measured. However,equation 1.11 canbe rearranged as:

(μw − μw∗)V−1

w = P − π (1.18)

Although the value of (μw − μ∗w) is more easily mea-

sured, the task of plant physiologists was simplified evenfurther when, in 1960, R. O. Slatyer and S. A. Taylorintroduced the concept of water potential (symbol-ized by the Greek uppercase psi, �). Water potential isproportional to (μw − μ∗

w) and can be defined as:

� = (μw − μw∗)V−1

w = P − π (1.19)

or simply:

� = P − π (1.20)

where P is the hydrostatic pressure and π is the osmoticpressure.

The concept of water potential has been widelyaccepted by plant physiologists because it avoids thedifficulties inherent in measuring chemical potential.Instead, it enables experimenters to predict the behaviorof water on the basis of two easily measured quantities, Pand π. It also makes it possible to express water potentialin units of pressure (pascals), which is more relevantto soil-plant-atmosphere systems than units of energy(joules). This distinction is not trivial. In practice it is fareasier to measure pressure changes than it is to measurethe energy required to affect water movement. Finally,

we can restate the driving force for water movementas the water potential gradient; that is, water will movefrom a region of high water potential to a region oflower water potential. As we shall see, however, waterpotentials are usually negative. This means that watermoves from a region of less negative water potential toa region where the water potential is more negative.

In the same way that the chemical potential of waterin a solution is measured against that of pure water,water potentials of solutions are also measured againsta reference. For water potential, the reference state isarbitrarily taken as pure water at atmospheric pressure.Under these conditions there is neither hydrostatic pres-sure nor dissolved solutes; that is, both P and π are zero.According to equation 1.20, the value of � for purewater is therefore also zero.

1.8 WATER POTENTIAL IS THESUM OF ITS COMPONENTPOTENTIALS

Water potential may be also be defined as the sum of itscomponent potentials:3

� = �P + �S (1.21)

The symbol �P represents the pressure potential. It isidentical to P and represents the hydrostatic pressure inexcess of ambient atmospheric pressure. The term �Srepresents the osmotic potential. Note the change in sign(π = − �S). As pointed out earlier, osmotic potentialis equal to osmotic pressure but carries a negative sign.Osmotic potential is also called solute potential (hencethe designation �S) because it is the contribution due todissolved solute. The term osmotic (or solute) potential ispreferred over osmotic pressure because it is more properlya property of the solution.

We can see from equation 1.21 that hydrostaticpressure and osmotic potential are the principal factorscontributing to water potential. A third component,the matric potential (M), is often included in theequation for water potential. Matric potential is aresult of the adsorption of water to solid surfaces. Itis particularly important in the early stages of wateruptake by dry seeds (called imbibition) and whenconsidering water held in soils (Chapter 3). There isalso a matric component in cells, but its contribution towater potential is relatively small compared with solutecomponent. It is also difficult to distinguish the matriccomponent from osmotic potential. Consequently,

3Students reading further in the literature will find that avariety of conventions, names, and symbols have been used todescribe the components of water potential. Thesedifferences are for the most part superficial, but carefulreading is required to avoid confusion.

12 Chapter 1 / Plant Cells and Water

matric potential may be excluded for purposes of thepresent discussion. We will return to matric potentialwhen we discuss soil water in Chapter 3.

Returning to equation 1.21, wecan see that anincrease in hydrostatic pressure or osmotic potentialwill increase water potential while a decrease in thepressure or osmotic potential (more negative) lowers it.We can use these changes to explain what happenedearlier in our example of the osmometer (Figure 1.8).The dissolved sucrose generated an osmotic potentialin the thistle tube, thereby lowering the water potentialof the solution (� < 0) compared with the pure water(� = 0) on the other side of the membrane. Waterthus diffused across the membrane into the solution.As the volume of the solution increased, a hydrostaticpressure developed in the thistle tube. When the pos-itive hydrostatic pressure was sufficient to offset thenegative osmotic potential, the water potential of thesolution was reduced to zero. At that point � = 0 onboth sides of the membrane and there was no furthernet movement of water. It is also interesting to notethat where volume in the osmometer is permitted toincrease, the osmotic potential will decrease. This isbecause along with the volume change accompanyingdiffusion of water into the solution, the mole fraction ofwater would also increase. In effect, the solute concen-tration decreases due to dilution. At equilibrium, then,the osmotic potential of the solution would be higher(i.e., less negative) than at the beginning. In the end thepressure required to balance osmotic potential is lessthan what would have been required had the volumeincrease not occurred. Because the contribution of �Sto water potential is always negative, water will, at con-stant pressure, always move from the solution with thehigher (less negative) osmotic potential to the solutionwith the lower (more negative) osmotic potential.

We can now ask what contributes to the osmoticand pressure potentials, and thus water potential, inplant cells. The osmotic potential of most plant cellsis due primarily to the contents of the large centralvacuole. With the exception of meristematic and cer-tain other highly specialized cells, cell vacuoles containon the order of 50 to 80 percent of the cellular waterand a variety of dissolved solutes. These may includesugars, inorganic salts, organic acids, and anthocyaninpigments. Most of the remaining cellular water is locatedin the cell wall spaces, while the cytoplasm accounts foras little as 5 to 10 percent. Methods for determiningthe osmotic potential of cells and tissues do not gener-ally discriminate between the cytoplasmic and vacuolarcontributions—the result is an average of the two. Theosmotic potential of a parenchyma cell is typically in therange of −0.1 to −0.3 MPa, the largest part of which isdue to dissolved salts in the vacuole.

In a laboratory osmometer, pressure (�P) can beestimated as the difference between atmospheric pres-

sure (0.1 MPa) and the hydrostatic pressure generatedby the height of the water column. In cells, the pressurecomponent arises from the force exerted outwardlyagainst the cell walls by the expanding protoplast. Thisis known as turgor pressure. An equal but oppositeinward pressure, called wall pressure, is exerted bythe cell wall. A cell experiencing turgor pressure is saidto be turgid. A cell that experiences water loss to thepoint where turgor pressure is reduced to zero is saidto be flaccid. Instruments are available for measuringP directly in large algal cells, but in higher plantsit is usually calculated as the difference betweenwater potential and osmotic potential. In nonwoodyherbaceous plants, turgor pressure is almost solelyresponsible for maintaining an erect habit. Indeed, oneof the first outward signs of water deficit in plants is thewilting of leaves due to loss of turgor in the leaf cells.

1.9 DYNAMIC FLUX OF H2O ISASSOCIATED WITH CHANGESIN WATER POTENTIAL

The water status of plant cells is constantly changing asthe cells adjust to fluctuations in the water content of theenvironment or to changes in metabolic state. Incipientplasmolysis is the condition in which the protoplastjust fills the cell volume. At incipient plasmolysis, theprotoplast exerts no pressure against the wall but neitheris it withdrawn from the wall. Consequently, turgorpressure (�P) is zero and the water potential of thecell (�cell) is equal to its osmotic potential (�S). Whenthe cell is bathed by a hypotonic4 solution such aspure water (� = 0), water will enter the cell as itmoves down the water potential gradient. This causessimultaneously a small dilution of the vacuolar contents(with a corresponding increase in osmotic potential) andthe generation of a turgor pressure. Net movement ofwater into the cell will cease when the osmotic potentialof the cell is balanced by its turgor pressure and, byequation 1.21, the water potential of the cell is thereforealso zero. When the cell is bathed by a hypertonicsolution, which has a more negative osmotic potentialthan the cell, the water potential gradient favors loss ofwater from the cell. The protoplast then shrinks awayfrom the cell wall, a condition known as plasmolysis.Continued removal of water concentrates the vacuolarcontents, further lowering the osmotic potential. Turgor

4A solution with a lower solute content than a cell or anothersolution and, hence, less negative osmotic potential, isreferred to as hypotonic. A hypertonic solution has a highersolute content and more negative osmotic potential. Asolution with an equivalent osmotic potential is known asisotonic.

1.10 Aquaporins Facilitate the Cellular Movement of Water 13

pressure remains at zero and the water potential of thecell is determined solely by its osmotic potential. Ineither situation described above, the water potential ofthe cell is determined as the algebraic sum of the turgorpressure and osmotic potential (Equation 1.21).

In addition to water movement between cells andtheir environment, diffusion down a water potential gra-dient can also account for water movement between cells(Figure 1.9). Individual cells in a series may experiencedifferent values for �S and �P, depending on the spe-cific circumstances of each cell. Nonetheless, water willflow through the series of cells so long as a continuousgradient in water potential is maintained.

The phenomena of plasmolysis and wilting aresuperficially the same, but there are some importantdifferences. Plasmolysis can be studied in the labora-tory simply by subjecting tissues to hypertonic solutionsand observing protoplast volume changes under themicroscope. As plasmolysis progresses, protoplast vol-ume progressively decreases, and the protoplast pullsaway from the cell wall. The void between the outer pro-toplast surface (the plasma membrane) and the cell wallwill become filled with external solution, which read-ily penetrates the cell wall. For this reason, plasmolysisdoes not normally give rise to a significant negative pres-sure (or tension) on the protoplast. Plasmolysis remainsessentially a laboratory phenomenon and, with the pos-sible exception of conditions of extreme water stress orsaline environments, seldom occurs in nature. Wilting,on the other hand, is the typical response to dehydrationin air under natural conditions. Because of its extremesurface tension, water in the small pores of the cell wallresists the entry of air and the collapsing protoplastmaintains contact with the cell wall. This tends to pullthe wall inward and substantial negative pressures maydevelop. The water potential of wilted cells becomeseven more negative as it is the sum of the negativeosmotic potential plus the negative pressure potential.

1.10 AQUAPORINS FACILITATETHE CELLULAR MOVEMENTOF WATER

Porins are a class of membrane proteins that belongto a large family of proteins called major intrinsicproteins (MIPs) that are found in the cell membranesof all living organisms including plants, microorgan-isms, and animals. Porin-type channels are nonselectivecation channels which are characterized by a β-pleatedsheet protein structure. In plants, porins are gener-ally restricted to the outer membranes of mitochondriaand chloroplasts and account for their highly perme-able properties. In contrast, aquaporins are membraneprotein channels or pores controlling the selective move-ment of water primarily. This is indicated by the factthat the presence of aquaporins does not affect theelectrical conductance of a membrane which indicatesthat small ions such as H+ are not conducted by thesemembrane channels. The results of genome sequencingindicate that Arabidopsis thaliana exhibits 35 aquaporingene homologs and 33 homologues have been detectedin the rice genome. Thus, plants exhibit a multiplicity ofaquaporin isoforms which, in part, reflects the multiplic-ity of internal membrane types in which these channelsare localized within a plant cell (Figure 1.10A). Forexample, 13 homologues have been detected in plantplasma membranes of Arabidopsis and are designatedPIPs for plasma membrane intrinsic proteins whereasthe tonoplast membrane, which surrounds the innervacuole of plant cells, exhibits 10 homologues and aredesignated TIPs.

The structure of aquaporins is highly conservedbetween plants, microbes, and animals. Four subunits(tetramers) associate to form a single plant aquaporin.Each subunit has a molecular mass of 23 to 31 kDaand spans the membrane with six α-helices joined by

ΨS = −0.5ΨP = 0.4Ψ = −0.1

ΨS = −0.4ΨP = 0.2Ψ = −0.2

ΨS = −0.5ΨP = 0.2Ψ = −0.3

XylemVesselΨ = 0

Decreasing water potential gradient

Direction of water flow

FIGURE 1.9 Diagram illustrating the contributions of osmotic potential (�S), turgorpressure (�p), and water potential (�) to water movement between cells. The direc-tion of water movement is determined solely by the value of the water potential inadjacent cells.

14 Chapter 1 / Plant Cells and Water

N

C

Plasma Membrane

Cytoplasm

B.

21 3 4 5 6

Cell wall

Plasma Membrane

Cytosol

Vacuole

PIP

TIP

A.

Tonoplast

H2O H2O

H2O

H2O

Cytoplasm

C.

H2O

H2OH2O

FIGURE 1.10 (A) Plant aquaporins are found in the plasma membrane (PIPs) as wellas the tonoplast membrane (TIPs) of the vacuole to regulate cellular water flow.(B) Each of the four subunits of a plant aquaporin spans the membrane six times. Boththe N-terminus (N) and C-terminus (C) are located in the cytoplasm. (C) Aquaporinsenhance water flow (thick arrow) and because they are gated allow the cell to reg-ulate cellular flow compared to water flow through pure lipid bilayers (thin arrow).The interior of the aquaporin channel contains hydrophilic amino acids, which inter-act with water, whereas the exterior of the protein channel consists of hydrophobicamino acids, which interact with the lipid fatty acids of the membrane bilayer.

1.11 Two-Component Sensing/Signalling Systems are Involved in Osmoregulation 15

5 intervening loops (Figure 1.10B) in contrast to theβ-pleated sheet structure of porins. These protein sub-units fold within the membrane such that the hydropho-bic amino acids are on the outer side of the pore andinteract with the hydrophobic fatty acids of the lipidbilayer whereas the hydrophilic amino acids are in theinner side of the pore and interact with water moleculesas they move through the pore from one side of themembrane to the other. Aquaporins can be open orclosed to regulate the movement of water across themembrane. Gating is the term used to describe this reg-ulated opening and closing of these protein channels.Gating through PIPs can be controlled by cytoplasmicpH, the concentration of divalent cations such as Ca2+as well as by aquaporin protein phosphorylation.

As discussed above, lipid bilayers are quite perme-able to water. Thus, there are two possible pathwaysfor the movement of water across a membrane—onepathway through the lipid bilayer itself and the secondpathway through an aquaporin (Figure 1.10C). If thisis so, why does a cell require any aquaporins at all?The results of recent experiments clearly indicate thatthe rate of water movement through a lipid bilayer withaquaporins is faster than a membrane that contains lipidsonly. Thus, the presence of aquaporins provides a lowresistance pathway for the movement of water acrossa membrane. Furthermore, since aquaporins are gated,this provides greater control for the movement of waterintracellularly as well as intercellularly. For example,the water permeability of the tonoplast is two orders ofmagnitude (102 times) greater than that of the plasmamembrane. Thus, this allows the vacuole to replenish orbuffer the cytoplasm with water when the cell is exposedto hypertonic conditions. Thus, aquaporins are impor-tant in regulating the osmotic properties of plant cells.This process is called osmoregulation.

1.11 TWO-COMPONENTSENSING/SIGNALLINGSYSTEMS ARE INVOLVEDIN OSMOREGULATION

Plants, green algae, fungi, as well as prokaryoticorganisms (bacteria, cyanobacteria) have cell walls andthus are sensitive to turgor pressure. These organismsmust be initially able to sense and then subsequentlyrespond accordingly at the physiological, biochemical,and genetic level in order to ensure daily survival aswell as seasonal changes in their aqueous environment.These organisms have evolved sophisticated sensingand signaling strategies to respond to changes in theirabiotic environment. This strategy generally employs,first, a sensing mechanism, which can usually detectchanges in specific environmental parameters (e.g., tem-perature, pressure, light quality, irradiance). A change

in an environmental parameter will usually activate aspecific sensor that will generate a specific intracellularsignal. Second, the sensor is usually linked to or coupledto a transducing mechanism, which propagates thesignal generated by the sensor and subsequently elicitsa specific cellular response.Two-component systemsare examples of such sensory signaling mechanisms andwere first discovered in prokaryotes. All two-componentsystems consist of a transmembrane sensor proteinlocated in the cell membrane and a cytoplasmic proteincalled the response regulator (Figure 1.11). Thesensor protein detects an environmental signal andtransmits this signal to the response regulator locatedin the cytoplasm. The response regulator mediatesthe cellular response to the external signal by DNAbinding or other regulatory functions that providetranscriptional control over one or more target genes.Thus, response regulators are typically DNA-bindingproteins or transcriptional factors.

An osmosensor is a device that is able either todetect changes in the chemical potential of extracellu-lar water (�μH2O) directly or to detect the mechanicalperturbations to the cell membrane as a consequenceof changes in turgor pressure due to the differences inosmotic potential (�S) between the external aqueousenvironment and the cytosol. In the heterotrophic bac-terium, Escherichia coli, a two-component system madeup of a sensor, EnvZ, and a response regulator, OmpR,enable this bacterium to sense changes in external osmo-larity. EnvZ and OmpR form an osmosensing/signaltransduction pathway that responds to (�μH2O) by reg-ulating the relative transcription of genes encoding twopore proteins, OmpF and OmpC. Both proteins formpores in the outer membrane that control cell membranepermeability. Although the total amount of both OmpFand OmpC remains relatively constant in response to�μH2O, the relative proportions of OmpF : OmpCchanges with changes in �μH2O. Under conditions oflow external osmolarity the expression of the largerOmpF pores are favored, whereas under high externalosmolarity the expression of the smaller OmpC poresare favored. As a consequence, the permeability of thebacterial cell membrane is altered.

Several two-component sensing/signal transduc-tion pathways have been reported in both plants andfungi. In the yeast, Saccharomyces cerevisiae, the histidinekinase SLN1 acts as a transmembrane osmosensor.The involvement of a two-component, histidine kinaseosmosensing system (ATHK1) has been describedrecently in Arabidopsis thaliana. Transforming yeastcells, in which the SLN1 gene was inactivated, with theplant ATHK1 gene restored the ability of the yeast cellsto sense and transduce a signal of changes in externalosmolarity (�μH2O). The accumulation of ATHK1mRNA in Arabidopsis is tissue specific with highestlevels observed in root tissue. Furthermore, the high

16 Chapter 1 / Plant Cells and Water

Cytoplasm

MEMBRANESENSOR

RESPONSEREGULATOR

Pi

Out

ExtracellularSignal

In

HIS

ATPADP

ASP ASP ASP

HIS

DNA-Binding

Gene ExpressionPhosphatase

HIS

P P

A. B. C.

Kinase

FIGURE 1.11 A general model illustrating two-component sensory signaling. (A)Two-component systems contain a transmembrane-bound protein sensor thatexhibits an extracellular and intracellular domain. The intracellular domain exhibitsa histidine kinase enzyme activity, which phosphorylates a specific histidine residue(HIS) within the intracellular domain of the sensor. The intracellular domain of thesensor is capable of interacting with a cytoplasmic response regulator protein, whichmay be a transcriptional activator and thus has DNA-binding activity. (B) The stimu-lation of the transmembrane sensor by an extracellular signal causes a conformationalchange in the extracellular domain of the membrane sensor, which activates the histi-dine kinase activity to autophosphorylate the intracellular domain of the sensor usingATP. (C) Subsequently, the sensory information flows from a membrane-bound sen-sor to a response regulator located in the cytoplasm. The phosphate group on thesensor histidine is transferred to an aspartyl group (ASP) on the response regula-tor. This relaxes the conformational of the transmembrane sensor but activates theDNA-binding activity of the response regulator. The binding of the phosphorylatedresponse regulator to DNA leads to an alteration in the expression of a specific geneor gene families in response to the extracellular signal. The response regulator alsoexhibits protein phosphatase activity which dephosphorylates the ASP group of theresponse regulator and inhibits its DNA-binding activity.

Further Reading 17

external salt concentrations stimulate the expression ofATHK1 in the roots of Arabidopsis. The high level ofexpression of ATHK1 in root tissue combined with itssensitivity to external salt concentrations suggest thatthe histidine kinase ATHK1, is part of a two-componentsignal transduction system efficient in osmosensing.

SUMMARY

Water has numerous chemical and physical propertiesthat make it particularly suitable as a medium in whichlife can occur. Most of these properties are the resultof the tendency of water molecules to form hydro-gen bonds. At the cellular level, water moves primarilyby osmosis, in response to a chemical potential gra-dient across a selectively permeable membrane. Themovement of water can be predicted on the basis ofwater potential. Water potential is a particularly usefulconcept because it can be calculated from two readilymeasured quantities: pressure and osmotic potential.Plants derive mechanical support from the turgidity ofcells, due at least in part to the high structural strengthof cell walls. Aquaporins are gated, membrane proteinchannels that regulate the permeability of cell mem-branes to water. Two-component sensing/signallingmechanisms play an important role in the ability ofplant cells to osmoregulate in reponse to changes inexternal water potential.

CHAPTER REVIEW

1. What is a hydrogen bond and how does ithelp to explain many of the unique phys-ical and chemical properties of water?

2. Describe osmosis as a special case of diffusion.Distinguish between osmotic pressure and osmoticpotential.

3. Understand the concept of water poten-tial and how it is related to the chemicalpotential of water. In what way does theconcept of water potential help the plantphysiologist explain water movement?

4. Explain why osmosis is, indirectly, anenergy-dependent process in plants.

5. Show how one can use osmotic potential to deter-mine molecular mass.

6. Can you suggest an important role for turgor inthe plant?

7. Is it osmotic potential or turgor pressurethat has the more significant role in reg-ulating water potential of plant cells?

8. Estimate the values of �cell, �S, and �P for atissue that neither gains nor loses weight whenequilibrated with a 0.4 molal mannitol solutionand in which, when placed in a 0.6 molal mannitolsolution, 50 percent of the cells are plasmolyzed.

9. What is an aquaporin? What role do theyplay in cell membrane permeability to water?

10. What is an osmosensor? Describe how atwo-component sensing/signalling system con-tributes to osmosensing.

FURTHER READING

Borstlap, A. C. 2002. Early diversification of plant aquapor-ins. Trends in Plant Science 7: 529–530.

Buchanan, B. B., W. Gruissem, R. L. Jones. 2000. Biochem-istry and Molecular Biology of Plants. American Society ofPlant Physiologists Rockville, Maryland.

Evert, R. F. 2006. Esau’s Plant Anatomy. Hoboken, NJ: JohnWiley & Sons, Inc.

Hoch, J. A., T. J. Silhavy. 1995. Two-Component Signal Trans-duction. Washington, D. C. ASM Press.

Maurel, C., L. Verdoucq, D.-T. Luu, V. Santoni. 2008. Plantaquaporins: membrane channels with multiple integratedfunctions. Annual Review of Plant Biology 59: 595–624.

Nobel, P. S. 2005. Physicochemical and Environmental PlantPhysiology. Elsevier Science & Technology Burlington,MA.

Pollack, G. H. 2001. Cells, Gels and the Engines of Life. Seattle:Ebner and Sons Publishers.

Urao, T., K. Yamaguchi-Shinozaki, K. Shinozaki. 2000.Two-component systems in plant signal transduction.Trends in Plant Science 5: 67–74.

Wood, J. M. 1999. Osmosensing by bacteria: Signals andmembrane-based sensors. Microbiology and Molecular Biol-ogy Reviews 63: 230–262.

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Bulk air

Unstirred boundary layer

Cuticle

Intercellularspaces

Stoma

2Whole Plant Water Relations

The dominant process in water relations of the wholeplant is the absorption of large quantities of water fromthe soil, its translocation through the plant, and itseventual loss to the surrounding atmosphere as watervapor. Of all the water absorbed by plants, less than5 percent is actually retained for growth and even lessis used biochemically. The balance passes through theplant to be lost as water vapor, a phenomenon known astranspiration. Nowhere is transpiration more evidentthan in crop plants, where several hundred kilograms ofwater may be required to produce each kilogram of drymatter and excessive transpiration can lead to significantreductions in productivity.

The quantitative importance of transpiration hasbeen indicated by a variety of studies over the years.In his classic 1938 physiology textbook, E. C. Millerreported that a single maize plant might transpire asmuch as 200 liters of water over its lifetime— approxi-mately 100 times its own body weight. Extrapolated toa field of maize plants, this volume of water is sufficientto cover the field to a depth of 38 cm over the courseof a growing season. A single, 14.5 m open-grown sil-ver maple tree may lose as much as 225 liters of waterper hour. In a deciduous forest, such as that foundin the southern Appalachians of the United States,one-third of the annual precipitation will be absorbedby plants, only to be returned to the atmosphere asvapor.

Whether there is any positive advantage to be gainedby transpiration is a point for discussion, but the poten-tial for such massive amounts of water loss clearly hasprofound implications for the growth, productivity, andeven survival of plants. Were it not for transpiration, forexample, a single rainfall might well provide sufficientwater to grow a crop. As it is, the failure of plants togrow because of water deficits produced by transpirationis a principal cause of economic loss and crop failureacross the world. Thus, on both theoretical and practi-cal grounds, transpiration is without doubt a process ofconsiderable importance.

This chapter will examine the phenomena of tran-spiration and water movement through plants. Theprincipal topics to be addressed include

• the process of transpiration and the role of vaporpressure differences in directing the exchange ofwater between leaves and the atmosphere,

• the role of environmental factors, in particular tem-perature and humidity, in regulating the rate oftranspirational water loss,

• the anatomy of the water-conducting system inplants, and how plants are able to maintain standingcolumns of water to the height of the tallest trees,and

• water in the soil and how water is taken up by rootsto meet the demands of water loss at the other end.

19

20 Chapter 2 / Whole Plant Water Relations

2.1 TRANSPIRATION IS DRIVENBY DIFFERENCES IN VAPORPRESSURE

Transpiration is defined as the loss of water from theplant in the form of water vapor. Although a smallamount of water vapor may be lost through small open-ings (called lenticels) in the bark of young twigs andbranches, the largest proportion by far (more than 90%)escapes from leaves. Indeed, the process of transpira-tion is strongly tied to leaf anatomy (Figure 2.1). Theouter surfaces of a typical vascular plant leaf are cov-ered with a multilayered waxy deposit called the cuticle.The principal component of the cuticle is cutin, aheterogeneous polymer of long-chain—typically 16 or18 carbons—hydroxylated fatty acids. Ester formationbetween the hydroxyl and carboxyl groups of neigh-boring fatty acids forms cross-links, establishing anextensive polymeric network.

The cutin network is embedded in a matrix of cutic-ular waxes, which are complex mixtures of long-chain(up to 37 carbon atoms) saturated hydrocarbons, alco-hols, aldehydes, and ketones. Because cuticular waxes arevery hydrophobic, they offer extremely high resistanceto diffusion of both liquid water and water vapor fromthe underlying cells. The cuticle thus serves to restrictevaporation of water directly from the outer surfacesof leaf epidermal cells and protects both the epidermaland underlying mesophyll cells from potentially lethaldesiccation.

The integrity of the epidermis and the overlyingcuticle is occasionally interrupted by small pores calledstomata (sing. stoma). Each pore is surrounded bya pair of specialized cells, called guard cells. Theseguard cells function as hydraulically operated valves thatcontrol the size of the pore (Chapter 8). The interiorof the leaf is comprised of photosynthetic mesophyllcells. The somewhat loose arrangement of mesophyll

Stoma Air space

CuticleUpper epidermis

Palisade

Spongy

Vein (vascular bundle)

Lower epidermis

Cuticle

Mesophyll

FIGURE 2.1 Diagrammatic representation of a typicalmesomorphic leaf (Acer sp.) shown in cross-section. Noteespecially the presence of a cuticle covering the outersurfaces of both the upper and lower epidermis. Notealso the extensive intercellular spaces with access to theambient air through the open stomata.

cells in most leaves creates an interconnected system ofintercellular air spaces. This system of air spaces maybe quite extensive, accounting for up to 70 percent ofthe total leaf volume in some cases. Stomata are locatedsuch that, when open, they provide a route for theexchange of gases (principally carbon dioxide, oxygen,and water vapor) between the internal air space and thebulk atmosphere surrounding the leaf. Because of thisrelationship, this space is referred to as substomatalspace. The cuticle is generally impermeable to waterand open stomata provide the primary route for escapeof water vapor from the plant.

Transpiration may be considered a two-stage pro-cess: (1) the evaporation of water from the moist cellwalls into the substomatal air space and (2) the diffu-sion of water vapor from the substomatal space into theatmosphere. It is commonly assumed that evaporationoccurs primarily at the surfaces of those mesophyll cellsthat border the substomatal air spaces. However, sev-eral investigators have proposed a more restricted view,suggesting instead that most of the water evaporatesfrom the inner surfaces of epidermal cells in the imme-diate vicinity of the stomata. Known as peristomalevaporation, this view is based on numerous reportsindicating the presence of cuticle layers on mesophyll cellwalls. In addition, mathematical modeling of diffusionin substomatal cavities has predicted that as much as75 percent of all evaporation occurs in the immediatevicinity of the stomata. The importance of peristomalevaporation versus evaporation from mesophyll surfacesgenerally remains to be established by direct experi-ment. Whether the evaporation occurs principally atthe mesophyll or epidermal cell surfaces is an interest-ing problem, reminding us that physiological processesare often not as straightforward as they may first appear.

The diffusion of water vapor from the substomatalspace into the atmosphere is relatively straightforward.Once the water vapor has left the cell surfaces, it dif-fuses through the substomatal space and exits the leafthrough the stomatal pore. Diffusion of water vaporthrough the stomatal pores, known as stomatal tran-spiration, accounts for 90 to 95 percent of the water lossfrom leaves. The remaining 5 to 10 percent is accountedfor by cuticular transpiration. Although the cuticle iscomposed of waxes and other hydrophobic substancesand is generally impermeable to water, small quantitiesof water vapor can pass through. The contribution ofcuticular transpiration to leaf water loss varies consider-ably between species. It is to some extent dependent onthe thickness of the cuticle. Thicker cuticles are charac-teristic of plants growing in full sun or dry habitats, whilecuticles are generally thinner on the leaves of plantsgrowing in shaded or moist habitats. Cuticular transpira-tion may become more significant, particularly for leaveswith thin cuticles, under dry conditions when stomataltranspiration is prevented by closure of the stomata.

2.2 The Driving Force of Transpiration is Differences in Vapor Pressure 21

2.2 THE DRIVING FORCEOF TRANSPIRATION ISDIFFERENCES IN VAPORPRESSURE

In Chapter 1, it was shown that water movement isdetermined by differences in water potential. It then canbe assumed that the driving force for transpiration is thedifference in water potential between the substomatal airspace and the external atmosphere. However, becausethe problem is now concerned with the diffusion ofwater vapor rather than liquid water, it will be moreconvenient to think in terms of vapor systems. Considerwhat happens, for example, when a volume of pure wateris introduced into a closed chamber (Figure 2.2). Initiallythe more energetic water molecules will escape into theair space, filling that space with water vapor. Some ofthose water molecules will then begin to condense intothe liquid phase. Eventually water in the chamber willreach a dynamic equilibrium; the rate of evaporationwill be balanced by the rate of condensation. The airspace will then contain the maximum amount of watervapor that it can hold at that temperature. In otherwords, at equilibrium the gas phase will be saturated withwater vapor. The concentration of water molecules in avapor phase may be expressed as the vapor mass per unitvolume (g m−3), called vapor density. Alternatively, theconcentration may be expressed in terms of the pressureexerted by the water vapor molecules against the fluid

surface and walls of the chamber. This is called vaporpressure (symbol = e). With an appropriate equation,vapor density and vapor pressure are interconvertible.However, because we are now accustomed to dealingwith the components of water potential in pressure units,it will be more consistent for us to use vapor pressure(expressed as kilopascals, kPa) in our discussion. We canthen say that when a gas phase has reached equilibriumand is saturated with water vapor, the system will haveachieved its saturation vapor pressure.

The vapor pressure over a solution at atmosphericpressure is influenced by both solute concentration andtemperature. As was previously discussed with respect towater potential (Chapter 1), the effect of solute concen-tration on vapor pressure may be expressed in terms ofthe mole fraction of water molecules. This relationshipis given by a form of Raoult’s law, which states:

e = Xi eo (2.1)

where e is vapor pressure of the solution, Xi is the molefraction of water ( = number of water molecules/numberof water molecules + number of solute molecules), andeo is the saturation vapor pressure over pure solvent.

The actual reduction in vapor pressure due to soluteturns out to be quite small. This is because even inrelatively concentrated solutions the mole fraction ofsolvent remains large. Consider, for example, a 0.5 molalsolution, which is approximately the concentration ofvacuolar sap in a typical plant cell. A 0.5 molal solution

b. c.A.

FIGURE 2.2 Vapor pressure in a closed container. Initially (A), more moleculesescape from the water surface than condense, filling the air space with water vapormolecules. The vaporous water molecules exert pressure—vapor pressure—againstthe walls of the chamber and the water surface. At equilibrium (B) the rate of conden-sation equals evaporation and the air is saturated with water vapor. The vapor pres-sure when the air is saturated is known as the saturation vapor pressure. At highertemperature (C), a higher proportion of water molecules have sufficient energy toescape. Both the concentration of water molecules in the vapor phase and the satura-tion vapor pressure are correspondingly higher.

22 Chapter 2 / Whole Plant Water Relations

contains 1/2 mole of solute dissolved in 1,000 grams(55.5 mol) of water. The mole fraction of water in a 0.5molal solution is therefore 55.5/(55.5 + 0.5) = 0.991.According to equation 2.1, the saturation vapor pressureof a half-molal solution would be reduced by less than 1percent compared with pure water.

Temperature, on the other hand, has a significanteffect on vapor pressure. This is due to the effect oftemperature on the average kinetic energy of the watermolecules. As the temperature of a volume of wateror an aqueous solution increases, the proportion ofmolecules with sufficient energy to escape the fluidsurface also increases. This in turn will increase theconcentration of water molecules in the vapor phaseand, consequently, the equilibrium vapor pressure. Anincrease in temperature of about 12◦C will nearly doublethe saturation vapor pressure.

According to Fick’s law of diffusion (Chapter 1),molecules will diffuse from a region of high concen-tration to a region of low concentration, or, down aconcentration gradient. Because vapor pressure is pro-portional to vapor concentration, water vapor will alsodiffuse down a vapor pressure gradient, that is, froma region of high vapor pressure to a region of lowervapor pressure. In principle, we can assume that thesubstomatal air space of a leaf is normally saturated orvery nearly saturated with water vapor. This is becausethe mesophyll cells that border the air space present alarge, exposed surface area for evaporation of water. Onthe other hand, the atmosphere that surrounds the leaf isusually unsaturated and may often have a very low watercontent. These circumstances create a gradient betweenthe high water vapor pressure in the interior of the leafand the lower water vapor pressure of the external atmo-sphere. This difference in water vapor pressure betweenthe internal air spaces of the leaf and the surroundingair is the driving force for transpiration.

2.3 THE RATE OFTRANSPIRATION ISINFLUENCED BYENVIRONMENTAL FACTORS

The rate of transpiration will naturally be influencedby factors such as humidity and temperature, and windspeed, which influence the rate of water vapor diffusionbetween the substomatal air chamber and the ambientatmosphere. Fick’s law of diffusion tells us that therate of diffusion is proportional to the difference inconcentration of the diffusing substance. It thereforefollows that the rate of transpiration will be governedin large measure by the magnitude of the vapor pressuredifference between the leaf (eleaf) and the surroundingair (eair). In other words,

T α eleaf − eair (2.2)

where T is the rate of transpiration. At the same time,the escape of water vapor from the leaf is controlled toa considerable extent by resistances encountered by thediffusing water molecules both within the leaf and inthe surrounding atmosphere (Figure 2.3). Resistance isencountered by the vapor molecules as they pass throughthe intercellular spaces, which are already saturated withwater vapor, and the stomatal pores. Note that the sym-bol for stomatal resistance indicates that it is variable,to account for the fact that stomata may at any time befully open, partially open, or closed. Additional resis-tance is encountered by the boundary layer, a layer ofundisturbed air on the surface of the leaf. The boundarylayer and its effect on tranpiration are described morefully later in this chapter. The transpiration equation(2.2) now requires additional terms to account for theseresistances.

T = eleaf − eair

rair + rleaf(2.3)

FIGURE 2.3 A schematic representation of theprincipal resistances encountered by water vapordiffusing out of a leaf. Symbols for electricalresistance are used because resistance to waterdiffusion is analogous to resistance in an elec-trical circuit. Note the symbol for stomatal resis-tance indicates it is variable—taking into accountthe capacity of the stomata to open and close.

Bulk air

Unstirred boundary layer

Cuticle

Intercellularspaces

Stoma

2.3 The Rate of Transpiration is Influenced by Environmental Factors 23

where rair and rleaf are resistances due to the air and theleaf respectively. Equation 2.3 tells us that the rate oftranspiration is proportional to the difference in vaporpressure between the leaf and the atmosphere divided bythe sum of resistances encountered in the air and the leaf.

2.3.1 WHAT ARE THE EFFECTSOF HUMIDITY?

Humidity is the actual water content of air, which, asnoted earlier, may be expressed either as vapor density(g m−3) or vapor pressure (kPa). In practice, however, itis more useful to express water content as the relativehumidity (RH). Relative humidity is the ratio of theactual water content of air to the maximum amountof water that can be held by air at that temperature.Expressed another way, relative humidity is the ratioof the actual vapor pressure to the saturation vaporpressure. Relative humidity is most commonly expressedas RH × 100, or percent relative humidity. The effectsof humidity and temperature on the vapor pressure ofair are illustrated in Table 2.1. Air at 50 percent RHby definition contains one-half the amount of waterpossible at saturation. Its vapor pressure is thereforeone-half the saturation vapor pressure. Note also that a10◦C rise in temperature nearly doubles the saturationvapor pressure. Relative humidity and temperature alsohave a significant effect on the water potential of air(Table 2.2).

As indicated earlier, the vapor pressure of the sub-stomatal leaf spaces is probably close to saturation mostof the time. Even in a rapidly transpiring leaf the rela-tive humidity would probably be greater than 95 percentand the resulting water potential would be close to zero(Table 2.2). Under these conditions, the vapor pressurein the substomatal space will be the saturation vaporpressure at the leaf temperature. The vapor pressure ofatmospheric air, on the other hand, depends on boththe relative humidity of the air and its temperature.Humidity and temperature thus have the potential tomodify the magnitude of the vapor pressure gradient(eleaf –eair), which, in turn, will influence the rate oftranspiration.

TABLE 2.1 Water Vapor Pressure (kPa) in air asa function of temperature and varying degrees ofsaturation. Air is saturated with water vapor at100% relative humidity (RH)

Relative Humidity

Temperature (˚C) 100% 80% 50% 20% 10%

30 4.24 3.40 2.12 0.85 0.4220 2.34 1.87 1.17 0.47 0.2310 1.23 0.98 0.61 0.24 0.12

TABLE 2.2 Some values for water potential (�)as a function of relative humidity (RH) at 20◦C

RH(%) � (MPa)

100 095 −6.990 −14.250 −93.520 −217.1

Water potential is calculated from the following relationship:� = 1.06 T log (RH/100)

2.3.2 WHAT IS THE EFFECTSOF TEMPERATURE?

Temperature modulates transpiration rate through itseffect on vapor pressure, which in turn affects the vaporpressure gradient as illustrated by the three examplesin Table 2.3. In the first example (A), assuming anambient temperature of 10◦C and a relative humidityof 50 percent, the leaf-to-air vapor pressure gradientis 0.61 kPa. This might be a typical situation inthe early morning hours. As the sun comes up, theair temperature will increase. A 10◦C increase intemperature (Table 2.3B), assuming the water contentof the atmosphere remains constant, will increase theleaf-to-air vapor pressure gradient and, consequently,the potential for transpiration, by a factor of almost3. Note that in this example it is assumed that leaftemperature is in equilibrium with the atmosphere.This is not always the case. A leaf exposed to full sunmay actually reach temperatures 5◦C to 10◦C higherthan that of the ambient air. Under these circumstances,the vapor pressure gradient may increase as much as

TABLE 2.3 The effect of temperature andrelative humidity on leaf-to-air vapor pressuregradient. In this example it is assumed that thewater content of the atmosphere remains constant

Leaf Atmosphere eleaf − eair

(A)T = 10˚C T = 10˚C

e = 1.23 kPa e = 0.61 kPa 0.61 kPaRH = 100% RH = 50%

(B)T = 20˚C T = 20˚C

e = 2.34 kPa e = 0.61 kPa 1.73 kPaRH = 100% RH = 26%

(C)T = 30˚C T = 20˚C

e = 4.24 kPa e = 0.61 kPa 3.63 kPaRH = 100% RH = 26%

24 Chapter 2 / Whole Plant Water Relations

sixfold (Table 2.3C). As long as the stomata remainopen and a vapor pressure gradient exists between theleaf and the atmosphere, water vapor will diffuse outof the leaf. This means transpiration may occur evenwhen the relative humidity of the atmosphere is 100percent. This is often the case in tropical jungles whereleaf temperature and, consequently, saturation vaporpressure is higher than the surrounding atmosphere.Because the atmosphere is already saturated, the watervapor condenses upon exiting the leaf, thereby givingsubstance to the popular image of the steaming jungle.

2.3.3 WHAT IS THE EFFECT OF WIND?

Wind speed has a marked effect on transpiration becauseit modifies the effective length of the diffusion path forexiting water molecules. This is due to the existenceof the boundary layer introduced earlier (Figure 2.3).Before reaching the bulk air, water vapor moleculesexiting the leaf must diffuse not only through the thick-ness of the epidermal layer (i.e., the guard cells), butalso through the boundary layer. The thickness of theboundary layer thus adds to the length of the diffusionpath. According to Fick’s law, this added length willdecrease the rate of diffusion and, hence, the rate oftranspiration.

The thickness of the boundary layer is primarily afunction of leaf size and shape, the presence of leaf hairs(trichomes), and wind speed. The calculated thicknessof the boundary layer as a function of wind speed overa typical small leaf is illustrated in Figure 2.4. Withincreasing wind speed, the thickness of the boundarylayer and, consequently, the length of the diffusion pathdecreases. In accordance with Fick’s law, the vapor pres-sure gradient steepens and, all other factors being equal,the rate of transpiration increases. This relationshipholds truest at lower wind speeds, however. As windspeed increases it tends to cool the leaf and may causesufficient desiccation to close the stomata. Either oneof these factors tends to lower the rate of transpiration.

1.0

2.0

3.0

Bou

ndar

y la

yer

thic

knes

s (m

m)

0.2 0.4 0.6 0.8 1.0 7.0 8.0 9.0 10.0

Wind speed (m s-1)

FIGURE 2.4 The impact of wind speed on calculatedboundary layer thickness for leaves 1.0 cm (triangles) or5.0 cm (circles) wide. A wind speed of 0.28 m s−1 = 1 kmhr−1. (Plotted from the data of Nobel, 1991.)

High wind speeds will therefore have less of an effecton transpiration rate than expected on the basis of theireffect on boundary layer thickness alone.

Boundary layer thickness can also be influenced by avariety of plant factors. Boundary layers are thicker overlarger leaves and leaf shape may influence the wind pat-tern. Leaf pubescence, or surface hairs, helps to maintainthe boundary layer, and thus reduce transpiration, bybreaking up the air movement over the leaf.

Given our discussion of transpiration, it is clearthat plant–water relations reflects the acquisition ofwater from the soil through the constant loss of waterthrough the leaves. This apparent conundrum may leadone to question whether transpiration offers any positiveadvantage to plants. This question is addressed in Box 2.1

2.4 WATER CONDUCTIONOCCURS VIA TRACHEARYELEMENTS

The distinguishing feature of vascular plants is thepresence of vascular tissues, the xylem and phloem,which conduct water and nutrients between the vari-ous organs. Vascular tissues begin differentiating a fewmillimeters from the root and shoot apical meristemsand extend as a continuous system into other organssuch as branches, leaves, flowers, and fruits. In organssuch as leaves, the larger veins subdivide into smallerand smaller veins such that no photosynthetic leaf cellis more than a few cells removed from a small veinending. Xylem tissue is responsible for the transportof water, dissolved minerals, and, on occasion, smallorganic molecules upward through the plant from theroot through the stem to the aerial organs. Phloem,on the other hand, is responsible primarily for thetranslocation of organic materials from sites of synthesisto storage sites or sites of metabolic demand (Chapter 9).

Xylem consists of fibers, parenchyma cells, andtracheary elements (Figure 2.5). Fibers are veryelongated cells with thickened secondary walls. Theirprincipal function is to provide structural support forthe plant. Parenchyma cells provide for storage as wellas the lateral translocation of solutes. The trachearyelements include both tracheids and vessel elements(Figure 2.5). Tracheary elements are the most highlyspecialized of the xylem cells and are the principalwater-conducting cells. Tracheids and vessels are bothelongated cells with heavy, often sculptured, secondarycell walls. Their most distinctive feature, however, isthat when mature and functioning, both tracheids andvessels form an interconnected network of nonlivingcells, devoid of all protoplasm. The hollow, tubularnature of these cells together with their extensive inter-connections facilitates the rapid and efficient transportof large volumes of water throughout the plant.

2.4 Water Conduction Occurs via Tracheary Elements 25

BOX 2.1WHYTRANSPIRATION?

Our discussion of transpiration in this chapter hasfocused on the mechanism of water loss and therole of transpiration in the ascent of sap—a sort ofoperational approach to the problem. We cannot fail tobe impressed by the amount of water that must be madeavailable to a plant in order to support transpirationand the possible consequences of such water loss toplant survival. Transpiration often results in waterdeficits and desiccation injury, especially when hightemperature and low humidity favor transpirationbut the soil is deficient of water. This raises aninteresting and often controversial question: Is thereany positive advantage in transpiration to be gainedby the plant? It has been argued that transpirationis required to bring about the ascent of sap, thatit increases nutrient absorption, and that it assistsin the cooling of leaves. It has also been arguedthat transpiration is little more than a ‘‘necessaryevil.’’

Transpiration does speed up the movement of xylemsap, but it seems unlikely that this is an essential require-ment. The growth of cells alone would cause a slowascent of xylem sap, even in the absence of transpi-ration. Transpiration serves only to increase the rateand quantity of water moved and there is no evi-dence that the higher rates are beneficial. Anotherargument is that, because mineral nutrients absorbedby the roots move largely in the xylem sap, tran-spiration may benefit nutrient distribution. It is truethat minerals in the xylem sap will be carried alongwith a rapidly moving transpiration stream, but therate-limiting step in nutrient supply is more likely tobe the rate at which the nutrients are absorbed by theroots and delivered to the xylem. Moreover, experi-ments with radioactive tracers have shown that mineralscontinue to circulate within the plant in the absence oftranspiration.

Because transpiration involves the evaporation ofwater, it can assume a significant role in the cool-ing of leaves. This is illustrated by the energy budgetfor a typical mesophyte leaf shown in the accompany-ing table. Because leaves are heavily pigmented, theyabsorb large amounts of direct solar radiation. Someof this absorbed solar radiation will not be utilized inphotochemical reactions, such as photosynthesis, butwill instead account for a significant heat gain by theleaves. Leaves also exchange infrared energy with theirsurroundings, both absorbing and radiating infrared.

Overall a leaf will radiate more infrared energy thanit gains, leaving a negative net infrared exchange. Thisleaves a net radiation gain in this example of 370 Wm−2, which must be dissipated by other means. Oneway of dissipating the heat load is by evaporation ofwater from the leaf surface, or transpiration. The latentheat of vaporization of water is 44 kJ mol−1 and a typi-cal mesophyte leaf might transpire at the rate of about4 mmol of water m−2 per second. The heat energy con-sumed by transpiration may be calculated as: (4 × 10−3

mol m−2 s−1) (44 × 10−3 J mol−1) = 176 J m−2 s−1

= 176 W m−2. In this example, transpiration can thusaccount for dissipation of approximately one-half of thenet radiation balance. Dissipation of the remaining heatis probably accounted for by convection from the leaf tothe surrounding air.

The energy budget for a typical mesophyte leaf.

Energy gain W m−2

a. Absorbed solar radiation +605b. Net infrared exchange −235c. Net radiation balance (a + b) +370Energy lossd. Loss by transpiration −176e. Loss by convection −194Net = −370

Data from Nobel, 1991.

One argument raised against a significant role for tran-spiration is that there is seldom any clear correlationbetween transpiration and plant growth. While someplants may develop more slowly at high humidity, manyare able to complete their life cycle without apparentharm under conditions such as 100 percent relativehumidity, where transpiration is minimal. Under suchcircumstances, the supply of water and nutrients isclearly adequate. If the leaf were not cooled by evapora-tive water loss, other processes such as convection mightremove sufficient energy to prevent the leaf reachinglethal temperatures.

In Chapter 8, it is argued that the evolutionaryfunction of stomata is to ensure an adequate supply ofcarbon dioxide for photosynthesis. It has been suggestedthat transpiration is simply an unfortunate consequenceof this function; that is, a structure that is efficient for thediffusive uptake of carbon dioxide is equally efficient forthe outward diffusion of water vapor. According to thisview, leaf structure represents a compromise betweenthe need to restrict desiccation of leaf cells while at thesame time maintaining access to atmospheric carbondioxide.

26 Chapter 2 / Whole Plant Water Relations

Borderedpits

A. B. C. D.

FIGURE 2.5 Tracheids from (A) spring wood of whitepine (Pinus) and (B) oak (Quercus). Vessel elements from(C) Magnolia and (D) basswood (Tilia). Only short tipsections are shown. (From T. E. Weier et al., Botany,6th ed. New York, Wiley, 1982, Figure 7.18. Used bypermission of the authors.)

Tracheids are single cells with diameters in therange of 10 to 50 μm. They are typically less than1 cm in length, although in some species they mayreach lengths up to 3 cm. Tracheids also have thickenedsecondary walls composed mainly of cellulose, hemicel-lulose, and lignin. Because of the high lignin content,secondary walls are less permeable to water than are theprimary walls of growing cells. On the other hand, theadditional strength of secondary walls helps to preventthe cells from collapsing under the extreme negativepressure that may develop in the actively transpiringplants. Although their principal function is to conductwater, the thickened secondary walls of tracheids alsocontribute to the structural support of the plant.

The movement of water between tracheids is facil-itated by interruptions, known as pit pairs, in thesecondary wall (Figure 2.6). During the developmentof tracheids, regions that are to become pit pairs avoidthe deposition of secondary wall material. This leavesonly the middle lamella and primary walls to separatethe hollow core, or lumen, of one cell from that of theadjacent cell. The combined middle lamella and primarywall is known as the pit membrane. Pit membranes arenot solid but have openings (about 0.3 μm diameter) thatpermit the relatively free passage of water and solutes.The origin of these openings is not clear, but they arethought to represent regions where cytoplasmic strands(plasmodesmata) once penetrated the cell walls whenthe cells were still alive. Bordered pit pairs have sec-ondary wall projections over the pit area and a swollen

A.

B.

C.

Aperture

Pit membraneTorus

Pit cavity

Border

FIGURE 2.6 Diagram of bordered pit pairs. (A) Twobordered pit pairs in the wall between two xylem tra-cheids, in side view. (B) Surface view. (C) A pressuredifferential—lower to the right—pushes the torusagainst the border, thereby sealing the pit and preventingwater movement. (From K. Esau, Anatomy of Seed Plants,New York, Wiley, 1977. Reprinted by permission.)

central region of the pit membrane called the torus.When pressure is unequal in adjacent vascular elements,such as when one contains an air bubble, the torusis drawn toward the element with the lower pressure.Pressure of the torus against the borders seals off the pitas shown in Figure 2.6.

Successive tracheids commonly overlap at theirtapered ends. As a result, tracheids line up in filesrunning longitudinally through the plant. Water movesbetween adjacent tracheids (either vertically or laterally)through the pit pairs in those regions where overlapoccurs. The movement of water is no doubt facilitatedby the openings in the pit membranes.

Vessels are very long tracheary elements made up ofindividual units, known as vessel members, which arearranged end-to-end in longitudinal series. At maturity,the end walls of the vessel members have dissolved away,leaving openings called perforation plates. As the cellthat is to become a vessel member develops, the futureperforation area becomes thickened due to swelling ofthe middle lamella (Figure 2.7). The thickened areacontains little cellulose, consisting almost entirely ofnoncellulosic polysaccharides. As the cell matures andthe cytoplasm begins to break down, the unprotectedsite of the perforation is attacked by hydrolytic enzymesand dissolved away. The rest of the wall area has been

2.5 The Ascent of Xylem Sap is Explained by Combining Transpiration with the Cohesive Forces of Water 27

Swollen middle lamella Simple perforation

Secondary wall Rim

A. B. C. D.

FIGURE 2.7 The development of a vessel member. (A)Meristematic cell. (B) Swelling of the middle lamella inthe region of a future perforation plate. (C) Secondarywall deposition except over area of future perforation.(D) Mature vessel member. The primary wall and middlelamella have dissolved away and the protoplast has disap-peared. (From K. Esau, Anatomy of Seed Plants, 1st ed.,New York, Wiley, 1960. Reprinted by permission.)

covered with lignified secondary wall materials and isprotected from degradation. In some cases, the resultingperforation will encompass virtually the entire end wall,leaving only a ring of secondary wall to mark the junctionbetween two successive vessel members (Figure 2.7). Inother cases, the plate may be multiperforate. If the per-forations are elongate and parallel, the pattern is calledscalariform. An irregular, netlike pattern is called retic-ulate. The perforations generally allow for a relativelyfree flow of water between successive vessel members.There are no perforation plates at the ends of vessels (i.e.,the last vessel member in a sequence), but water is ableto move laterally from one vessel to the next due to thepresence of pit pairs similar to those found in tracheids.

The size of vessels is highly variable, although theyare generally larger than tracheids. In maples (Acer sps.),for example, vessels range from 40 μm to 60 μm in diam-eter, while in some species of oak (Quercus sps.) diametersmay range up to 300 μm to 500 μm. (The large-diametervessels account for the ring porus character of springwood in woody species.) The length of vessels in mapleis generally 4 cm or less, but some may reach lengths of30 cm. In oak, on the other hand, vessel lengths up to10 m have been recorded. However, because of extensivebranching of the vascular system and the large numberof lateral connections between overlapping trachearyelements, the xylem constitutes a single continuous,interconnected system of water-conducting conduitsbetween the extremes of the plant—from the tip ofthe longest root to the outermost margins of the highestleaf.

Vessels are considered evolutionarily moreadvanced than tracheids. For example, xylem tissue

in the gymnosperms, considered evolutionarily moreprimitive than the angiosperms, consists entirely oftracheids. Although tracheids do occur in angiosperms,the bulk of the water is conducted in vessels. Alsobecause of their larger size, vessels are considerablymore efficient than tracheids when it comes toconducting water. An empirical equation relatingflow rate to the size of conduits was developed inthe nineteenth century by the French scientist JeanL. M. Poiseuille. Poiseuille showed that when a fluid ispressure-driven, the volume flow rate ( Jv) is a functionof the viscosity of the liquid (η), the difference inpressure or pressure drop (�P), and the radius of theconduit:

Jv = �P πr4/8η (2.4)

Equation 2.4 applies to water movement in the xylemtracheary elements because, as will be shown below, itis driven by a difference in pressure between the soiland the leaves. The important point to note, then, isthat the volume flow rate is directly proportional to thefourth power of the radius. The impact of this relation-ship can be seen by comparing the relative volumeflow rates for a 40-μm-diameter (r = 20 μm) tracheidand a 200-μm-diameter (r = 100 μm) vessel. Althoughthe relative diameter of the vessel is 5 times that ofthe tracheid, its relative volume flow rate will be 625(i.e., 54) times that of the tracheid. The high rate offlow in the larger vessels occurs because the flow rate ofwater is not uniform across the conduit. The flow rateof molecules near the conduit wall is reduced by fric-tion, due to adhesive forces between the water and theconduit wall. As the diameter of the conduit increases,the proportion of molecules near the wall and conse-quently subject to these frictional forces will decrease.Put another way, the faster-moving molecules in thecenter of the conduit constitute a larger proportion ofthe population and the overall rate of flow increasesaccordingly.

2.5 THE ASCENT OF XYLEMSAP IS EXPLAINED BYCOMBININGTRANSPIRATION WITHTHE COHESIVE FORCESOF WATER

The tallest-standing trees are generally found growing inthe rainforests along the Pacific coast of the northwest-ern United States and southwestern British Columbia.The best known are the redwoods (Sequoia sempervirens)of northern California, some of which exceed 110 m inheight. Individual specimens of Douglas fir (Pseudotsugamenziesii) have been reported in excess of 100 m and

28 Chapter 2 / Whole Plant Water Relations

Pressure+ 101.3 k Pa

Vacuum

10.3 m

FIGURE 2.8 Atmospheric pressure can support a watercolumn to a maximum height of 10.3 m.

a Sitka spruce (Picea sitchensis) measuring 95 m has beenlocated in the Carmanah Valley of Vancouver Island.In Australia, there have been reports of Eucalyptus treesmeasuring more than 130 m in height.

The forces required to move water to such heightsare substantial. Were we able to devise a sufficientlylong tube closed at one end, fill it with water, andinvert it as shown in Figure 2.8, we would find thatatmospheric pressure (ca. 101 kPa at sea level) wouldsupport a column of water approximately 10.3 m inheight. To push the water column any higher wouldrequire a correspondingly greater pressure acting on theopen surface. Clearly, elevating water to the height ofthe tallest trees would require a force 10 to 15 timesgreater than atmospheric, or 1.0 to 1.5 MPa. This forcewould be equivalent to the pressure at the base of astanding column of water 100 m to 150 m high.

But even a force of this magnitude would not besufficient. In addition to the force of gravity, water mov-ing through the plant will encounter a certain amountof resistance inherent in the structure of the conductingtissues—irregular wall surfaces, perforation plates, andso forth. We can assume that a force at least equal tothat required to support the column would be neces-sary to overcome these resistances. In that case, a forceon the order of 2.0 to 3.0 MPa would be required tomove water from ground level to the top of the tallestknown trees. How can such a force be generated? Thisis a question that has long held the interest of plantphysiologists and over the years a number of theories

have been advanced. The three most prominent are rootpressure, capillarity, and the cohesion theory.

2.5.1 ROOT PRESSURE IS RELATED TOROOT STRUCTURE

If the stem of a well-watered herbaceous plant is cut offabove the soil line, xylem sap will exude from the cutsurface. Exudation of sap, which may persist for severalhours, indicates the presence of a positive pressure in thexylem. The magnitude of this pressure can be measuredby attaching a manometer to the cut surface (Figure 2.9).This pressure is known as root pressure because theforces that give rise to the exudation originate in theroot.

Root pressure has its basis in the structure of rootsand the active uptake of mineral salts from the soil. Thexylem vessels are located in the central core of a root,the region known as the stele. Surrounding the steleis a layer of cells known as the endodermis. In mostroots, the radial and transverse walls of the endoder-mal cells develop characteristic thickenings called theCasparian band (Figure 2.10). The Casparian band isprincipally composed of suberin, a complex mixture ofhydrophobic, long-chain fatty acids, and alcohols. Thesehydrophobic molecules impregnate the cell wall, fillingin the spaces between the cellulose microfibrils as wellas the intercellular spaces between the cells. Becauseit is both space-filling and hydrophobic, the Casparianband presents an effective barrier to the movement ofwater through the apoplastic space of the endodermis.The result is that water can move into or out of thestele only by first passing through the membranes of theendodermal cells and then through the plasmodesmatalconnections.

Mercury

Exuded xylem sap

FIGURE 2.9 A simple manometer for measuring rootpressure. Root pressure can be calculated from theheight of the mercury in the glass tube.

2.5 The Ascent of Xylem Sap is Explained by Combining Transpiration with the Cohesive Forces of Water 29

B.

Endodermis

10 μmPrimary phloem

Primaryxylem

Pericycle

Casparian band

A.

FIGURE 2.10 Suberin deposits (Casparian strip) in the walls of root endodermal cells.(A) Cross section. (B) Three-dimensional view. Suberin deposits in the radial wallsestablish a barrier to movement of water and salts in the apoplast of the endoder-mis. (From K. Esau, Anatomy of Seed Plants, New York, Wiley, 1977. Reprinted bypermission.)

As roots take up mineral ions from the soil, the ionsare transported into the stele where they are activelydeposited in the xylem vessels. The accumulation ofions in the xylem lowers the osmotic potential and,consequently, the water potential of the xylem sap. Inresponse to the lowered water potential, water follows,also passing from the cortical cells into the stele throughthe membranes of the endodermal cells. Since the Cas-parian band prevents the free return of water to thecortex, a positive hydrostatic pressure is established inthe xylem vessels. In a sense, the root may be thoughtof as a simple osmometer (refer to Figure 1.8) in whichthe endodermis constitutes the differentially permeablemembrane, the ions accumulated in the xylem representthe dissolved solute, and the xylem vessels are the ver-tical tube. So long as the root continues to accumulateions in the xylem, water will continue to rise in thevessels or exude from the surface when the xylem vesselsare severed.

The question to be answered at this point is whetherroot pressure can account for the rise of sap in a tree.The answer is probably no, for several reasons. Tobegin with, xylem sap is not as a rule very concentratedand measured root pressures are relatively low. Valuesin the range of 0.1 to 0.5 MPa are common, whichare no more than 16 percent of that required to movewater to the top of the tallest trees. In addition, rootpressure has not been detected in all species. Finally,it has been clearly established that during periods ofactive transpiration, when water movement through thexylem would be expected to be most rapid, the xylemis under tension (i.e., negative pressure). Root pressureclearly cannot serve as the mechanism for the ascentof sap in all cases. However, root pressure could serve

to fill vessels in small, herbaceous plants and in somewoody species in the spring when sap moves up to thedeveloping buds.

2.5.2 WATER RISE BY CAPILLARITY ISDUE TO ADHESION AND SURFACETENSION

If a glass capillary tube (i.e., a tube of small diameter)is inserted into a volume of water, water will rise in thetube to some level above the surface of the surround-ing bulk water. This phenomenon is called capillaryrise, or simply capillarity. Capillary rise is due to theinteraction of several forces. These include adhesionbetween water and polar groups along the capillarywall, surface tension (due to cohesive forces betweenwater molecules), and the force of gravity acting on thewater column. Adhesive forces attract water molecules topolar groups along the surface of the tube. When thesewater-to-wall forces are strong, as they are betweenwater and glass tubes or the inner surfaces of trachearyelements, the walls are said to be wettable. As water flowsupward along the wall, strong cohesive forces betweenthe water molecules act to pull the bulk water up thelumen of the tube. This will continue until these liftingforces are balanced by the downward force of gravityacting on the water column.

The following equation can be used to calculate therise of water in a capillary tube where h is the

h = 1.49 × 10−5m2/r m (2.5)

rise in meters and r is the radius of the capillary tubein meters (m). Clearly, the calculated rise of water ina capillary tube is inversely proportional to the radius

30 Chapter 2 / Whole Plant Water Relations

of the tube. In a large tracheid or small vessel, with adiameter of 50 μm (r = 25 μm), water will rise to aheight of about 0.6 m. For a large vessel (r = 200 μm),capillarity would account for a rise of only 0.08 m. Onthe basis of these numbers, capillarity in tracheids andsmall vessels might account for the rise of xylem sap insmall plants, say less than 0.75 m in height. However,to reach the height of a 100 m tree by capillarity, thediameter of the capillary would have to be about 0.15μm—much smaller than the smallest tracheids. Clearlycapillarity is inadequate as a general mechanism for theascent of xylem sap.

2.5.3 THE COHESION THEORY BESTEXPLAINS THE ASCENT OFXYLEM SAP

The most widely accepted theory for movement of waterthrough plants is known as the cohesion theory. Thistheory depends on there being a continuous columnof water from the tips of the roots through the stemand into the mesophyll cells of the leaf. The theory isgenerally credited to H. H. Dixon, who gave the firstdetailed account of it in 1914.

2.5.3.1 What is the driving force? According tothe cohesion-tension theory, the driving force for watermovement in the xylem is provided by evaporationof water from the leaf and the tension or negativepressure that results. Water covers the surfaces of themesophyll cells as a thin film, adhering to cellulose andother hydrophillic surfaces. As water evaporates fromthis film, the air–liquid interface retreats into the smallspaces between cellulose microfibrils and the angularjunctions between adjacent cells. This creates very smallcurved surfaces or microscopic menisci (Figure 2.11).As the radii of these menisci progressively decrease,surface tension at the air–water interface generates anincreasingly negative pressure, which in turn tends todraw more liquid water toward the surface. Because thewater column is continuous, this negative pressure, ortension, is transmitted through the column all the way tothe soil. As a result, water is literally pulled up throughthe plant from the roots to the surface of the mesophyllcells in the leaf.

The cohesion theory raises two very importantquestions: (1) Is the xylem sap of a rapidly transpir-ing plant under tension? (2) How is the integrity of verytall water columns maintained? In the absence of anydirect evidence, the answers to these questions providesubstantial indirect support for the theory.

Except in certain circumstances, such as when rootpressure is active, pressures in the xylem are rarelypositive. On the other hand, several lines of evidencesupport the conclusion that xylem water is instead undersignificant tension. First, if one listens carefully when

Cell wall

Vacuole

Intercellularspace

Cytoplasm

Cytoplasm

Vacuole

Cell wall

Water

FIGURE 2.11 Tension (negative pressure) in the watercolumn. Evaporation into the leaf spaces causes thewater–air interface (dashed lines) to retreat into thespaces between and at the junctions of leaf mesophyllcells. As the water retreats, the resulting surface tensionpulls water from the adjacent cells. Because the watercolumn is continuous, this tension is transmitted throughthe column, ultimately to the roots and soil water.

the xylem of a rapidly transpiring plant is severed, it issometimes possible to hear the sound of air being drawnrapidly into the wound. If severed beneath the surfaceof a dye solution, the dye will be very rapidly takenup into tracheary elements in the immediate vicin-ity of the wound. A second line of evidence involvessensitive measuring devices, called dendrographs, whichcan be used to measure small changes in the diameterof woody stems. Diameters decrease significantly dur-ing periods of active transpiration. This will happenbecause the stem is slightly elastic and the tension inthe water column pulls the tracheary walls inward. Inthe evening, when transpiration declines, the tensionis released and stem diameter recovers. Moreover, theshrinkage occurs first in the upper part of the tree, clos-est to the transpiring leaves, when transpiration beginsin the morning. Only later does it show up in the lowerpart. This observation has been confirmed by experi-ments in which a localized pulse of heat is generated inthe xylem and the flow of heat away from that point ismonitored. Flow rates indicate that tensions are greaternear the top of the tree. A variety of other experimentshave demonstrated that rapidly transpiring shoots areable to pull a column of mercury to heights greaterthan can be accounted for by atmospheric pressurealone.

2.5 The Ascent of Xylem Sap is Explained by Combining Transpiration with the Cohesive Forces of Water 31

Direct measurement of tension in xylem vesselswas made possible by the introduction of the pressurebomb technique by P. F. Scholander. If the xylemsolution is under tension, it will withdraw from thecut surface but can be forced back to the surface byincreasing the pressure in the chamber. With such adevice, Scholander and others have measured tensionson the order of −0.5 to −2.5 MPa in rapidly transpiringtemperate-zone trees.

Finally, it has been observed that water potentialsnear the bottom of the tree are less negative than waterpotentials higher in the crown. Such a pressure dropbetween the bottom and the top of the crown is con-sistent with a tension resulting from forces originatingat the top of the tree. The weight of evidence clearlysupports the hypothesis that the xylem water column isliterally pulled up the tree in response to transpiration.

2.5.3.2 How is the integrity of the water columnmaintained? The ability to resist breakage of thewater column is a function of the tensile strength ofthe water column. Tensile strength is a measure ofthe maximum tension a material can withstand beforebreaking. Tensile strength is expressed as force per unitarea, where the area for the purpose of our discussionis the cross-sectional area of the water column. Tensilestrength is yet another property of water attributableto the strong intermolecular cohesive forces, or hydro-gen bonding, between the water molecules. The tensilestrength of water (or any fluid, for that matter) is noteasily measured—a column of water does not lenditself to testing in the same way as a steel bar or acopper wire. The tensile strength of water will alsodepend on the diameter of the conduit, the propertiesof the conduit wall, and the presence of any dissolvedgases or solute. Still, a number of ingenious approacheshave been developed to measure the tensile strength ofwater with fairly consistent results. It is now generallyaccepted that pure water, free of dissolved gas, is ableto withstand tensions as low as −25 to −30 MPa at20◦C. This is approximately 10 percent of the tensilestrength of copper, and 10 times greater than the −2.5to −3.0 MPa required to pull an uninterrupted watercolumn to the top of the tallest trees. As noted above,tensions in the xylem are more typically in the rangeof −0.5 to −2.5 MPa for temperate deciduous treessuch as maple (Acer sps.), but may sometimes be as lowas–10 Mpa.

Because xylem water is under tension, it must re-main in the liquid state well below its vapor pressure—recall that the vapor pressure of water at 20◦C and100% RH is 2.3 kPa or 0.0023 MPa (Table 2.1). A watercolumn under tension is therefore physically unstable.Physicists call this condition a metastable state, a state inwhich change is ready to occur but does not occur in theabsence of an external stimulus. Stability can be achieved

in a water column under tension by introducing a vaporphase. Water molecules in the vapor phase have verylow cohesion, which allows the vapor to expand rapidly,thus causing the column to rupture and relieve the ten-sion. How might a vapor phase be introduced to thexylem column? Xylem water contains several dissolvedgases, including carbon dioxide, oxygen, and nitrogen.When the water column is under tension, there is atendency for these gases to come out of solution. Sub-microscopic bubbles first form at the interface betweenthe water and the walls of the tracheid or vessel, proba-bly in small, hydrophobic crevices or pores in the walls.These small bubbles may redissolve or they may coa-lesce and expand rapidly to fill the conduit. This processof rapid formation of bubbles in the xylem is calledcavitation (L. cavus, hollow). The resulting large gasbubble forms an obstruction, called an embolism (Gr.embolus, stopper), in the conduit. The implications ofembolisms with respect to the cohesion theory are quiteserious, because a conduit containing an embolism is nolonger available to conduct water. Indeed, the poten-tial for frequent cavitation in the xylem was raised as aprincipal objection to the cohesion theory when it wasinitially proposed. In order to satisfy these objections,it was necessary to determine just how vulnerable thexylem was to cavitation.

Early attempts to relate cavitation to tensions devel-oped in the xylem were largely inconclusive. Therewere no satisfactory methods for observing cavitationin the xylem itself and model systems, employing glasstubes, did not necessarily duplicate the interface con-ditions present in plant tissues. This all changed in1966 when J. A. Milburn and R. P. C. Johnson intro-duced an acoustic method for detecting cavitation inplants. In laboratory experiments with glass tubes, therapid relaxation of tension that follows cavitation pro-duces a shockwave that can be heard as an audibleclick. Milburn and Johnson found that similar clickscould be ‘‘heard’’ in plant tissue by using sensitivemicrophones and amplifiers. Each click is believed torepresent formation of an embolism in a single vesselelement.

Milburn and Johnson studied cavitation inwater-stressed leaves of castor bean (Ricinus communis).Water stress was introduced by detaching the leaffrom the plant and permitting it to wilt. As the leafwilted, the number of clicks occurring in the petiolewas recorded. A total of 3,000 clicks were detected,which is approximately equal to the number of vesselsthat might be expected in such a petiole. Cavitationcould be prevented by adding water to the severedend of the petiole. Various methods that eitherincreased or decreased transpiration from the leafresulted in a corresponding increase or decrease in the

32 Chapter 2 / Whole Plant Water Relations

number of clicks. These results indicate a reasonablystraightforward relationship between cavitation andtension in the xylem, which appears to support thecohesion theory. They further suggest that cavitationis readily induced by water stress, a condition thatherbaceous plants might be expected to encounter on adaily basis. A long-term study of cavitation in a standof sugar maple (Acer saccharum) has been conductedusing a method that measures changes in hydraulicconductance. In its simplest form, conductance isthe inverse of resistance. Hydraulic conductance istherefore a measure of the total capacity of the tissueto conduct water. The acoustic method is limited tocounting the number and frequency of cavitations.The hydraulic method, on the other hand, assesses theimpact of the resulting embolisms on the capacity ofthe tissue to transport water.

During the summer growing season, embolismsappeared to be confined to the main trunk and reducedhydraulic conductance by 31 percent. During the winter,loss of conductance in the main trunk increased to 60percent, while some twigs suffered a 100 percent loss! Adecline in conductance during the summer months is nodoubt attributable to water stress, as it is in herbaceousplants. The rise in embolisms during the winter isprobably related to freeze–thaw cycles. The solubilityof gases is very low in ice; when tissue freezes, gasis forced out of solution. During a thaw, these smallbubbles will expand and nucleate cavitation.

Problems related to cavitation are not limited tomature trees. Newly planted seedlings often experi-ence water stress due to poor root–soil contact andmay be vulnerable to cavitation. A recent study foundthat seedlings of western hemlock (Tsuga heterophylla)experienced water stress and declining xylem pressurepotential when planted out. The resulting cavitation andembolism formation in the tracheids caused a decline inhydraulic conductance in the seedling. If the decline inhydraulic conductance is severe enough, it can lead todefoliation or death of the seedling.

Clearly, the effect of cavitation and embolisms onlong-term survival of plants would be disastrous if therewere not means for their removal or for minimizingtheir effects. The principal mechanism for minimiz-ing the effect of embolisms is a structural one. Theembolism is simply contained within a single tracheidor vessel member. In those tracheary elements withbordered pit pairs, the embolism is contained by thestructure of the pit membrane (Figure 2.12A). A differ-ence in pressure between the vessel member containingthe embolism and the adjacent water-filled vessel causesthe torus to press against the pit border, thus pre-venting the bubble from being pulled through. Atthe same time, surface tension prevents the bubblefrom squeezing through the small openings in theperforation plates between successive vessel members

Air

A. B.

Air

FIGURE 2.12 Diagram to illustrate how water flowbypasses embolisms in tracheids and vessels. In tra-cheids (A), the pressure differential resulting from anembolism causes the torus to seal off bordered pits liningthe affected tracheary element. In vessels (B), the bubblemay expand through perforation plates, but will even-tually be stopped by an imperforate end wall. In bothtracheids and vessels, surface tension prevents the airbubbles from squeezing through small pits or capillarypores in the side walls. Water, however, continues tomove around the blockage by flowing laterally into adja-cent conducting elements.

(Figure 2.12B). Water, however, will continue to flowlaterally through available pits, thus detouring aroundthe blocked element by moving into adjacent conduits.In addition to bypassing embolisms, plants may alsoavoid long-term damage by repairing the embolism.This can happen at night, for example, when tran-spiration is low or absent. Reduced tension in thexylem water permits the gas to simply redissolve inthe xylem solution. An alternative explanation, partic-ularly in herbaceous species, is that air may be forcedback into solution on a nightly basis by positive rootpressure.

The repair of embolisms in taller, woody species isnot so easily explained. As noted above, sugar maples dorecover from freezing-induced embolisms in the spring.Maples and many other tree species also exhibit positivexylem pressures in the spring. These pressures accountfor the springtime sap flow and could also serve toreestablish the continuity of the water column. Spring-time recovery has also been documented for grapevines,although this is apparently due to relatively high rootpressures at this time of the year. It might be possible thatwoody plants in general develop higher-than-normal

2.6 Water Loss due to Transpiration must be Replenished 33

root pressures in the spring in order to overcome win-ter damage. Finally, most woody species produce newsecondary xylem each spring. This new xylem tissueis laid down before the buds break and may meet thehydraulic conductance needs of the plant, replacingolder, nonfunctional xylem.

It is clear, even on the basis of the relatively few stud-ies that have been completed, that xylem is particularlyvulnerable to cavitation and embolism. If herbaceousplants were unable to recover on a nightly basis orwoody species were unable to recover in the spring, theirgrowth and ultimate survival would be severely compro-mised. This might lead one to question why plantshave not evolved mechanisms to lessen their effects.Interestingly, M. H. Zimmermann turned this questionaround, proposing instead that cell walls might actuallybe designed to cavitate. This statement may appear tocontradict an earlier observation that surface tensionprevents the passage of embolisms through perforationsin xylem conduits, but it does not. Where a water-filledconduit under tension is separated from an air volume(at atmospheric pressure) by a porous wall, a concavemeniscus (the ‘‘lifting force’’) will form in the pore tobalance the negative pressure. As the pressure differen-tial increases, the radius of the meniscus will decrease.At a ‘‘suitable’’ tension in the xylem, the meniscus willreach a radius less than the diameter of the pore (a frac-tion of a μm in diameter) and will be pulled through thepore into the water-filled conduit. The resulting bub-ble nucleates a cavitation, which relieves the tension,reduces the pressure differential, and prevents the fur-ther entry of air. Results consistent with Zimmermann’shypothesis have been obtained through a comparisonof the pressure differences required to embolize sugarmaple stems with diameters of the pores in the pitmembranes.

Accordingly, Zimmermann’s designed leakagehypothesis represents a kind of safety valve. Plantsappear to be constructed in such a way as to allowcavitation when water potentials reach critically lowlevels, yet still allow the ‘‘damage’’ to be repaired whenconditions improve. Water stress and, consequently,cavitations would be expected to appear first in leavesand smaller branches. These localized cavitations wouldserve the additional advantage of cutting off peripheralstructures while preserving the integrity of the mainstem or trunk during extended dry periods.

Roots are even more vulnerable to cavitation thanshoots, which could benefit the whole plant duringperiods of drought. Complete cavitation of the xylemin the smallest roots, for example, would isolate thoseroots from drying soil, reduce hydraulic conductance,and ultimately reduce transpiration rates. This wouldhelp buffer the water status of the stem until the drought

eased and the cavitated conduits were refilled or newgrowth replaced the damaged roots.

2.6 WATER LOSS DUE TOTRANSPIRATION MUST BEREPLENISHED

Uptake of water from the soil by the roots replenishes thewater lost as a consequence of leaf transpiration. Thisestablishes an integrated flow of water from the soil,through the plant, and into the atmosphere, referred toas the soil-plant-atmosphere continuum. The con-cept of a soil-plant-atmosphere continuum reinforcesthe observation that plants do not exist in isolation, butare very interdependent with their environment.

2.6.1 SOIL IS A COMPLEX MEDIUM

In order to understand interactions between roots andsoil water, a review of the nature of soils would be help-ful. Soil is a very complex medium, consisting of a solidphase comprised of inorganic rock particles and organicmaterial, a soil solution containing dissolved solutes,and a gas phase generally in equilibrium with the atmo-sphere. The inorganic solid phase of soils is derived fromparent rock that is degraded by weathering processes toproduce particles of varying size (Table 2.4). In additionto the solid, liquid, and gas phases, soils also containorganic material in varying stages of decomposition aswell as algae, bacteria, fungi, earthworms, and variousother organisms.

The clay particles in a soil combine to form com-plex aggregates that, in combination with sand andsilt, determine the structure of a soil. Soil structure inturn affects the porosity of a soil and, ultimately, itswater retention and aeration. Porosity, or pore space,refers to the interconnected channels between irregu-larly shaped soil particles. Pore space typically occupiesapproximately 40 percent to 60 percent of a soil by vol-ume. Two major categories of pores— large pores and

TABLE 2.4 Classification of soil particles andsome of their properties. A mixture of 40 percentsand, 40 percent silt, and 20 percent clay is knownas a loam soil. A sandy soil contains less than 15percent silt and clay, while a clay soil containsmore than 40 percent clay particles

Particle (mm) WaterParticle Class Size Retention Aeration

Coarse sand 2.00–0.2 poor excellentSand 0.20–0.02Silt 0.02–0.002 good goodClay less than 0.002 excellent poor

34 Chapter 2 / Whole Plant Water Relations

capillary pores—are recognized. Although there is nosharp line of demarcation between large pores and cap-illary pores—the shape of the pore is also a determiningfactor—water is not readily held in pores larger than 10to 60 μm diameter. When a soil is freshly watered, suchas by rain or irrigation, the water will percolate downthrough the pore space until it has displaced most, ifnot all, of the air. The soil is then saturated with water.Water will drain freely from the large pore space dueto gravity. The water that remains after free (gravity)drainage is completed is held in the capillary pores. Atthis point, the water in the soil is said to be at fieldcapacity. Under natural conditions, it might requiretwo to three days for a loam soil to come to field capac-ity following a heavy rainfall. The relative proportionsof large and capillary pore space in a soil can be esti-mated by determining the water contents of the soilwhen freshly watered and at field capacity. Water con-tent, expressed as the weight of water per unit weightof dry soil, may be determined by drying the soil at105◦C.

It should not be surprising that a sandy soil, with itscoarse particles, will have a relatively high proportionof large pores. A sandy soil will therefore drain rapidly,has a relatively low field capacity, and is well-aerated(Table 2.4). The pore space of a clay soil, on the otherhand, consists largely of capillary pores. Clay soils holdcorrespondingly larger quantities of water and are poorlyaerated. A loam soil represents a compromise, balanc-ing water retention against aeration for optimal plantgrowth.

The water held by soil at or below field capacityis found in capillary channels and the interstitial spacesbetween contacting soil particles, much as it is in thecell walls and intercellular spaces of mesophyll cells ina leaf. Soil water is therefore also subject to the sameforces of surface tension found in the capillary spacesof mesophyll tissues, as discussed earlier. Consequently,soil water at or below field capacity will be under ten-sion and its water potential will be negative. As the watercontent of the soil decreases, either by evaporation fromthe soil surface or because it is taken up by the roots,the air–water interface will retreat into the capillaryspaces between the soil particles. Because water adheresstrongly to the soil particles, the radius of the meniscusdecreases and pressure becomes increasingly negative.In principle, water movement in the soil is primarilypressure-driven in the same manner that capillary ten-sion in the mesophyll cells of a leaf draws water fromthe xylem column. As water is removed from the soil bya root, tensions in the soil water will draw more bulkwater toward the root. If there is an abundance of waterin the soil, these pressure differences may draw waterfrom some distance.

Except in highly saline soils, the solute concentra-tion of soil water is relatively low—on the order of10−3 M—and soil water potential is determined prin-cipally by the negative pressure potential. As might beexpected, the uptake of water by roots occurs because ofa water potential gradient between the soil and the root.Thus as the soil dries and its water potential declines,plants may experience difficulty extracting water fromthe soil rapidly enough to balance losses by transpi-ration. Under such conditions, plants will lose turgorand wilt. If transpiration is reduced or prevented fora period of time (such as at night, or by covering theplant with a plastic bag), water uptake may catch up,turgor will be restored, and the plants will recover.Eventually, however, a point can be reached where thewater content of the soil is so low that, even should allwater loss by transpiration be prevented, the plant isunable to extract sufficient water from the soil and theloss of turgor is permanent. The soil water content atthis point, measured as a percentage of soil dry weight,is known as the permanent wilting percentage. Theactual value of the permanent wilting percentage variesbetween soil types; it is relatively low (in the range of1 to 2 percent) for sand and high (20 to 30 percent)for clay. Loam soils fall between these two extremes,depending on the relative proportion of sand and clay.Regardless of soil type, however, the water potential ofthe soil at the permanent wilting percentage is relativelyuniform at about −1.5 MPa. Although there are someexceptions to the rule, most plants are unable to extractsignificant amounts of water when the soil water poten-tial falls below −1.5 MPa. In a sense, field capacity maybe considered a property of the soil, while the perma-nent wilting percentage is a property of the plant. Thewater content of the soil between field capacity and thepermanent wilting percentage is considered availablewater, or water that is available for uptake by plants.The range of available water is relatively high in siltyloam soils, somewhat less in clay, and relatively low insand. Not all water in this range is uniformly available,however. In a drying soil, plants will begin to showsigns of water stress and reduced growth long beforethe soil water potential reaches the permanent wiltingpercentage.

2.7 ROOTS ABSORB ANDTRANSPORT WATER

Roots have four important functions. Roots (1) anchorthe plant in the soil; (2) provide a place for storage of car-bohydrates and other organic molecules; (3) are a site ofsynthesis for important molecules such as alkaloids and

2.8 The Permeability of Roots to Water Varies 35

some hormones; and (4) absorb and transport upwardto the stem virtually all the water and minerals taken upby plants.

The effectiveness of roots as absorbing organs isrelated to the extent of the root system. Over the years,a number of efforts have attempted to establish thetrue extent of root systems. Such studies involve carefuland often tedious procedures for excavating the plantand removing the soil without damaging the roots.More elaborate efforts involved construction of sub-terranean walkways, called rhizotrons, equipped withwindows through which the growth and developmentof roots could be observed. The root systems of severalprairie grasses have been examined in different types ofsoils. By carefully excavating the grasses and washingthe roots free of soil, they were able to show massivenetworks of roots penetrating to a depth of 1.5 m. Usinga radioactive tracer technique, it was later found theroots of a single 14-week-old corn plant had penetratedto a depth of more than 6 m and extended horizon-tally as much as 5 m in all directions. Furthermore, themeasurements of the roots of a single mature rye plantgrown in a box of soil measuring 30 cm × 30 cm × 56 cmdeep have been reported. The combined length of allroots was 623 km with an estimated total surface areaof 639 m2. Many species invest substantially more than50 percent of their body weight in roots. Roots clearlycomprise a large proportion of the plant body, althoughtheir importance to plants is less obvious to the casualobserver than the more visible shoot system.

2.8 THE PERMEABILITY OFROOTS TO WATER VARIES

A large number of anatomical and physiological studieshave established that the region of most active wateruptake lies near the root tip. Beyond this generalization,the permeability of roots to water varies widely withage, physiological condition, and the water status ofthe plant. Studies with young roots have shown thatthe region most active with respect to water uptakestarts about 0.5 cm from the tip and may extend downthe root as far as 10 cm (Figure 2.13). Little water isabsorbed in the meristematic zone itself, presumablybecause the protoplasm in this zone is dense and thereare no differentiated vascular elements to carry the wateraway. The region over which water appears to be takenup most rapidly corresponds generally with the zone ofcell maturation. This is the region where vascular tissue,in particular the xylem, has begun to differentiate. Alsoin this region the deposition of suberin and lignin in thewalls of endodermal cells is only beginning and has not

Suberization andlignificationreducepermeability

Suberization andlignification ofendodermisbeginning

Stele

Cortex

Xylemdifferentiating

Region of elongation

Meristematicregion

Root cap

Most rapid entranceof water and salt

Slow entrance ofwater and salt

Root hair zone

Relativelyimpermeableto water

Slow entrance of waterand salt because ofdecreasing permeability

FIGURE 2.13 Diagrammatic illustration of the relation-ship between differentiation of roottissues and water uptake.

yet reached the point of offering significant resistanceto water movement.

The region of most rapid water uptake also coin-cides with the region of active root hair development.Root hairs are thin-walled outgrowths of epidermal cellsthat increase the absorptive surface area and extendthe absorptive capacity into larger volumes of soil(Figure 2.14). Depending on the species and envi-ronmental conditions, root hairs may reach lengths of0.1 mm to 10 mm and an average diameter of 10 μm. Insome species such as peanut, pecan, and certain conifers,root hairs are rare or absent. More commonly a singleroot tip may contain as many as 2,500 hairs cm−2 andmay increase the absorbing surface of the root 1.5 totwentyfold. For two reasons, root hairs greatly increasethe contact of the root with soil water. First, their smalldiameter permits root hairs to penetrate capillary spacesnot accessible to the root itself (Figure 2.14A). Sec-ond, root hairs extend contact into a cylinder of soilwhose diameter is twice that of the length of the hair(Figure 2.14B).

36 Chapter 2 / Whole Plant Water Relations

800μm

Soilparticle

Roothair

Rootepidermis

200μm 800μm

1800μm

A.

b.

FIGURE 2.14 Root hairs and water uptake. (A) Roothairs enhance water uptake by their ability to penetratewater-containing capillary spaces between soil particles.(B) Root hairs increase by several times the volume of soilthat can be extracted of water by a root. (A from T. E.Weier et al., Botany, 6th ed., New York, Wiley, 1982,Figure 9.7A. Used by permission of the authors.)

2.9 RADIAL MOVEMENT OFWATER THROUGH THEROOT INVOLVES TWOPOSSIBLE PATHWAYS

Once water has been absorbed into the root hairs orepidermal cells, it must traverse the cortex in order toreach the xylem elements in the central stele. In princi-ple, the pathway of water through the cortex is relativelystraightforward. There appear to be two options: watermay flow either past the cells through the apoplast of thecortex or from cell to cell through the plasmodesmataof the symplast. In practice, however, the two pathwaysare not separate. Apoplastic water is in constant equi-libration with water in the symplast and cell vacuoles.This means that water is constantly being exchangedacross both the cell and vacuolar membranes. In effect,

then, water flow through the cortex involves both path-ways. The cortex consists of loosely packed cells withnumerous intercellular spaces. The apoplast would thusappear to offer the least resistance and probably accountsfor a larger proportion of the flow.

In the less mature region near the tip of the root,water will flow directly from the cortex into the develop-ing xylem elements, meeting relatively little resistancealong the way. Moving away from the tip toward themore mature regions, water will encounter the endo-dermis (see section 2.5.1 on root pressure). Suberizationof the endodermal cell walls imposes a permeabilitybarrier, forcing water taken up in these regions to passthrough the cell membranes. While the endodermisdoes increase resistance to water flow, it is far frombeing an absolute barrier. Indeed, under conditions ofrapid transpiration the region of most rapid water uptakewill shift toward the basal part of the root (Table 2.5).

The resistance offered by the root to water uptake isreflected in the absorption lag commonly observed whenwater loss by transpiration is compared with absorptionby the roots (Figure 2.15). That this lag is due to resis-tance in the root can be demonstrated experimentally.If the roots of an actively transpiring plant are cut off(under water, of course), there is an immediate increasein the rate of absorption into the xylem. In some species,absorption lag may cause a water deficit in the leavessufficient to stimulate a temporary closure of the stomata(Figure 2.15). This phenomenon is known as middayclosure. Closure of the stomata reduces transpiration,allowing absorption of water to catch up and the stomatato then reopen.

Finally, although the absorption of water by rootsis believed to be a passive, pressure-driven process,it is nonetheless dependent on respiration in the

TABLE 2.5 Relative water uptake for differentzones along a Vicia faba root as a function oftranspiration rate. The measured xylem tension atthe low transpiration rate was–0.13 MPa and–10.25 MPa at the high transpiration rate. Whentranspiration is low, most of the water uptakeoccurs near the root tip. When transpiration ishigh, the resulting increase in tension shifts theregion of uptake toward more basal regions

Transpiration RateWater Uptake Zone(cm from apex) Low High

0–2.5 100 1322.5–5.0 103 2165.0–7.5 54 2167.5–10.0 27 27010.5–12.5 19 283

From data of Brouwer, 1965.

Further Reading 37

Time of day (h)

0 2 4 6 8 10 12 14 16 18 20 22 24

Tran

spir

atio

n or

ste

m f

low

(kg

h-1

)

15

10

5

0

-.5

-1.0

-1.5Leaf

wat

erpo

tent

ial (

M P

a)TranspirationStem flow

FIGURE 2.15 The absorption lag in a Larix (larch) tree.Upper: Absorption of water, measured as the flow ofwater through the stem, lags about two hours behindtranspiration. A transient decline in transpiration ratenear midday may occur if the lag is sufficient to createa water deficit and stimulate temporary stomatal clo-sure. Lower: Rapid transpiration causes a decrease inthe leaf water potential, which slowly recovers as watermoves in to satisfy the deficit. (Adapted from E.-D.Schulze et al., Canopy transpiration and water fluxesin the xylem of Larix and Picea trees—A comparisonof xylem flow, porometer and cuvette measurements,Oecologia 66:475–483, 1985. Reprinted by permission ofSpringer-Verlag.)

root cells. Respiratory inhibitors (such as cyanide ordinitrophenol), high carbon dioxide levels, and lowoxygen (anaerobiosis) all stimulate a decrease in thehydraulic conductance of most roots. Anaerobiosis ismost commonly encountered by plants in water-loggedsoils and may lead to extreme wilting in species notspecifically adapted to such situations. The exact roleof respiration is not clear. The requirement is probablyindirect, such as maintaining the cellular integrityand continued elongation of roots. Active uptake ofnutrient ions (Chapter 3) may also be a factor. Onthe other hand, killing roots outright—for example,by immersion in boiling water—dramatically reducesresistance and allows water to be absorbed more rapidlythan when the roots were alive.

SUMMARY

Large amounts of water are lost by plants throughevaporation from leaf surfaces, a process known astranspiration. Transpiration is driven by differencesin water vapor pressure between internal leaf spacesand the ambient air. A variety of factors influence tran-spiration rate, including temperature, humidity, wind,

and leaf structure. Water is conducted upward throughthe plant primarily in the xylem, a tube-like systemof tracheary elements including tracheids and vessels.The principal driving force for water movement inthe xylem is transpiration and the resulting tension inthe water column. The water column is maintainedbecause of the high tensile strength of water. Waterlost by transpiration is replenished by the absorption ofwater from the soil through the root system.

CHAPTER REVIEW

1. Explain why transpiration rate tends to begreatest under conditions of low humidity,bright sunlight, and moderate winds.

2. Describe the anatomy of xylem tissue andexplain why it is an efficient system forthe transport of water through the plant.

3. Trace the path of water from the soil, through theroot, stem, and leaf of a plant, and into the atmo-sphere.

4. Explain how water can be moved to the top of a100 m tree, but a mechanical pump can lift waterno higher than about 10.3 m. What prevents thewater column in a tree from breaking? Underwhat conditions might the water column break,and, if it does break, how is it reestablished?

5. Many farmers have found that fertilizingtheir fields during excessively dry periodscan be counterproductive, as it may signif-icantly damage their crops. Based on yourknowledge of the water economy of plantsand soils, explain how this could happen.

6. Does transpiration serve any useful function in theplant?

7. Explain the relationships between field capacity,permanent wilting percentage, and available water.Even though permanent wilting percentage isbased on soil weight, it is often said to be a prop-erty of the plants. Explain why this might be so.

FURTHER READING

Waisel, Y., A. Eshel, U. Kafkafi. 1996. Plant Roots: The HiddenHalf . 2nd ed. New York: M. Dekker.

Buchanan, B. B., W. Gruissem, R. L. Jones. 2000. Biochem-istry and Molecular Biology of Plants. Rockville, MD.:American Society of Plant Physiologists.

Evert, R. F. 2006. Esau’s Plant Anatomy. Hoboken, NJ: JohnWiley & Sons, Inc.

Nobel, P. S. 2005. Physicochemical and Environmental PlantPhysiology. Burlington, MA: Elsevier Science & Technol-ogy.

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3Roots, Soils, and Nutrient Uptake

With the exception of carbon and oxygen, which aresupplied as carbon dioxide from the air, terrestrialplants generally take up nutrient elements from the soilsolution through the root system. The root systemsof most plants are surprisingly extensive. Through acombination of primary roots, secondary and tertiarybranches, and root hairs, root systems penetrate massivevolumes of soil in order to mine the soil for requirednutrients and water. Soil is a complex medium. It con-sists of a solid phase that includes mineral particlesderived from parent rock plus organic material in vari-ous stages of decomposition, a liquid phase that includeswater or the soil solution, gases in equilibrium with theatmosphere, and a variety of microorganisms. The solidphase, in particular the mineral particles, is the primarysource of nutrient elements. In the process of weather-ing, various elements are released into the soil solution,which then becomes the immediate source of nutrientsfor uptake by the plant.

The soil solution, however, is very dilute (totalmineral content is on the order of 10−3 M) and wouldquickly become depleted by the roots were it not con-tinually replenished by the release of elements from thesolid phase. In one study of phosphorous uptake, forexample, it was calculated that the phosphate content ofthe soil was renewed on the average of 10 times each

day. Availability of nutrient elements is not, however,limited to the properties of the soil itself. Access toelements is further enhanced by the continual growthof the very dynamic root system into new, nutrient-richregions of the soil.

In this chapter we will examine the availability ofnutrients in the soil and their uptake by roots. This willinclude

• the soil as a source of nutrient elements, the colloidalnature of soil, and ion exchange properties thatdetermine the availability of nutrient elements in aform that can be taken up by roots,

• mechanisms of solute transport across membranes,including simple and facilitated diffusion and activetransport, the function of membrane proteins as ionchannels and carriers, and the role of electrochemi-cal gradients,

• ion traffic into and through the root tissues and theconcept of apparent free space, and

• the beneficial role of microorganisms, especiallyfungi, with respect to nutrient uptake by roots.

The unique situation with respect to uptakeand metabolism of nitrogen will be addressed inChapter 11.

39

40 Chapter 3 / Roots, Soils, and Nutrient Uptake

3.1 THE SOIL AS A NUTRIENTRESERVOIR

Soils vary widely with respect to composition, struc-ture, and nutrient supply. Especially important from thenutritional perspective are inorganic and organic soilparticles called colloids. Soil colloids retain nutrientsfor release into the soil solution where they are availablefor uptake by roots. Thus, the soil colloids serve tomaintain a reservoir of soluble nutrients in the soil.

3.1.1 COLLOIDS ARE A SIGNIFICANTCOMPONENT OF MOST SOILS

The physical structure of soils was introducedpreviously in the discussion of water uptake by roots(Chapter 2). There it was noted that the mineralcomponent of soils consists predominantly of sand,silt, and clay, which are differentiated on the basisof particle size (see Chapter 2, Table 2.4). The threecomponents are easily demonstrated by stirring a smallquantity of soil into water. The larger particles of sandwill settle out almost immediately, leaving a turbidsuspension. Over the course of hours or perhaps days,if left undisturbed, the finer particles of silt will settleslowly to the bottom as well and the turbidity will inall likelihood disappear. The very small clay particles,however, remain in stable suspension and will notsettle out, at least not within a reasonable time frame.Clay particles in suspension are not normally visibleto the naked eye—they are simply too small. They are,however, small enough to remain suspended. These sus-pended clay particles can be detected by directing a beamof light through the suspension. The suspended clay par-ticles will scatter the light, causing the path traversed bythe light beam to become visible. Particles that are smallenough to remain in suspension but too large to go intotrue solution are called colloids and the light-scatteringphenomenon, known as the Tyndall effect, is a distin-guishing characteristic of colloidal suspensions. A truesolution, such as sodium chloride or sucrose in water, onthe other hand, will not scatter light. This is because, in atrue solution, the solute and solvent constitute a singlephase. A colloidal suspension, on the other hand, is a two-phase system. It consists of a solid phase, the colloidalmicelle, suspended in a liquid phase. Light scattered bythe solid phase is responsible for the Tyndall effect.

Clay is not the only soil component that formscolloidal particles. Many soils also contain a colloidalcarbonaceous residue, called humus. Humus is organicmaterial that has been slowly but incompletely degradedto a colloidal dimension through the action of weath-ering and microorganisms. In a good loam soil, thecolloidal humus content may be substantially greaterthan the colloidal clay content and make an even greatercontribution to the nutrient reservoir.

3.1.2 COLLOIDS PRESENT A LARGE,NEGATIVELY CHARGEDSURFACE AREA

The function of the colloidal soil fraction as a nutrientreservoir depends on two factors: (1) colloids presenta large specific surface area, and (2) the colloidalsurfaces carry a large number of charges. The chargedsurfaces in turn reversibly bind large numbers of ions,especially positively charged cations from the soilsolution. This ability to retain and exchange cations oncolloidal surfaces is the single most important propertyof soils, in so far as plant nutrition is concerned.Because of their small size, one of the distinguishingfeatures of colloids is a high surface area per unit mass,also known as specific surface area. Consider, for ex-ample, a cube with a mass of one gram that measures 10mm on a side. The specific surface area of this cube is600 mm2 g−1 (Figure 3.1). If the cube is then subdividedinto particles of colloidal dimensions, say 0.001 mm ona side, the specific surface area increases to 6,000,000mm2g−1, a 10,000-fold increase. On a mass basis,then, colloids provide an incredibly large surfacearea for interaction with mineral elements in the soilsolution.

A second important feature of soil colloids, in addi-tion to the large specific surface area, is the largenumber of charges on the colloidal surfaces. Colloidalclays consist primarily of aluminum silicates (the chemi-cal formula for kaolinite, one of the simplest clays, isAl2Si2O5(OH)4). The predominantly negative chargesarise by virtue of ionization of alumina and silica at the

10 mm

10 mm10 mm

FIGURE 3.1 Particles of colloidal dimension have a highsurface area per unit mass, or specific surface area.

3.1 The Soil as a Nutrient Reservoir 41

edges of the clay particle. Because colloidal carbon isderived largely from lignin and carbohydrates, it alsocarries negative charges arising from exposed carboxyland hydroxyl groups.

3.1.3 SOIL COLLOIDS REVERSIBLYADSORB CATIONS FROM THESOIL SOLUTION

The association of cations (positively charged ions)with negatively charged colloidal surfaces depends onelectrostatic interactions; hence, binding affinity variesaccording to the lyotropic series:

Al3+ > H+ > Ca2+ > Mg2+ > K+ = NH+4 > Na+

In this series, aluminum ions have the highest bindingaffinity and sodium ions the least, reflecting the generalrule that trivalent ions (3+) are retained more stronglythan divalent ions (2+) and divalent ions more stronglythan monovalent ions (1+). Ions, however, are alsohydrated, which means that they are surrounded byshells of water molecules, and electrostatic rules aremodulated by the relative hydrated size of the ion. Sinceions of smaller hydrated size can approach the colloidalsurfaces more closely, they tend to be more tightlybound. Also, cation adsorption is not an all-or-nonephenomenon. The degree of association and ion con-centration both decline in a continuous gradient withincreasing distance from the surface of the colloid.

Cation adsorption is also reversible. Consequently,any ion with a higher affinity (e.g., H+) is capableof displacing an ion lower in the series (e.g., Ca2+).Alternatively, an ion with a lower affinity can, if providedin sufficient quantity, displace an ion with higher affinityby mass action (Figure 3.2). This process of exchangebetween adsorbed ions and ions in solution is known asion exchange. The ease of removal, or exchangeabilityof an ion, is indicated by the reverse of the lyotropicseries shown above. Thus, sodium ions are the mostreadily exchanged in the series and aluminum ions theleast.

Although the immediate source of mineral nutri-ents for the plant are the ions in the soil solution, thecolloidal fraction with its adsorbed ions representsthe principal nutrient reservoir. It is important to viewthe soil as a very dynamic system, with cations in thesoil solution freely exchangeable with cations adsorbedto colloidal surfaces. As the soluble nutrients are takenup by the roots from the dilute soil solution, they arecontinually replaced by exchangeable ions held in thecolloidal reservoir. The reservoir is then replenishedby ions derived from the weathering of rock parti-cles. In this way, ion exchange at the colloidal surfaceplays a major role in providing a controlled release ofnutrients to the plant. It may not always work to theadvantage of the plant, however. One effect of acid

ClayMicelle

K+

K+

–

Ca2+–

K+

–

Ca2+

Acidify soil

–

K+

–

–

–

–

–

–

H+

–

NH4+

Mg2+

–

ClayMicelle

K+

K+

–

Ca2+

Ca2+

–

K+

–

Ca2+

–

K+K+

–

––––––

H+

H+

H+

H+

H+

H+

–

NH4+

Mg2+ Mg2+

–

FIGURE 3.2 Ion exchange in the soil. (A) Cations areadsorbed to the negatively charged soil particles by elec-trostatic attraction. (B) Acidifying the soil increases theconcentration of hydrogen ions in the soil. The addi-tional hydrogen ions have a stronger attraction for thecolloidal surface charges and so displace other cationsinto the soil solution.

rain, for example, appears to be the displacement ofcations from the colloidal reservoir due to the high con-centration of hydrogen ions. Those solubilized nutri-ent ions not immediately assimilated by the roots arereadily leached out of the soil by the rain and percolatingground water. Both the soil solution and the reservoirof nutrients are depleted more rapidly than they can bereplenished and the plants are deprived of an adequatenutrient supply. Under normal circumstances, roots alsosecrete hydrogen ions, which assist in the uptake ofnutrients.

3.1.4 THE ANION EXCHANGECAPACITY OF SOIL COLLOIDSIS RELATIVELY LOW

The soil colloids are predominantly negatively chargedand, consequently, they do not tend to attract negativelycharged anions. Although some of the clay minerals docontain cations such as Mg2+, the anion exchange capa-city of most soils is generally low. The result is thatanions are not held in the soil but tend to be read-ily leached out by percolating ground water. Thissituation has important consequences for agriculturalpractice. Nutrients supplied in the form of anions, inparticular nitrogen (NO−

3 ), must be provided in largequantity to ensure sufficient uptake by the plants. As arule, farmers sometimes find they must apply at leasttwice—sometimes more—the amount of nitrogen actu-ally required to produce a crop. Unfortunately, much ofthe excess nitrate is leached into the ground water andeventually finds its way into wells or into streams and

42 Chapter 3 / Roots, Soils, and Nutrient Uptake

lakes, where it contributes to problems of eutrophicationby stimulating the growth of algae.

3.2 NUTRIENT UPTAKE

3.2.1 NUTRIENT UPTAKE BY PLANTSREQUIRES TRANSPORT OF THENUTRIENT ACROSS ROOT CELLMEMBRANES

In order for mineral nutrients to be taken up by aplant, they must enter the root by crossing the plasmamembranes of root cells. From there they can be trans-ported through the symplast to the interior of the rootand eventually find their way into the rest of the plant.Nutrient uptake by roots is therefore fundamentally acellular problem, governed by the rules of membranetransport. Membrane transport is inherently an abstractsubject. That is to say, investigators measure the kineticsof solute movement across various natural and artificialmembranes under a variety of circumstances. Models arethen constructed that attempt to explain these kineticpatterns in terms of what is currently understood aboutthe composition and architecture of membranes. As ourunderstanding of membrane structure has changed overthe years, so have the models that attempt to interprethow solutes cross these membranes. There are, how-ever, three fundamental concepts—simple diffusion,facilitated diffusion, and active transport—thathave persevered, largely because they have provenparticularly useful in categorizing and interpretingexperimental observations. These three conceptsnow make up the basic language of transport acrossall membranes of all organisms. These three basic

modes of transport are interpreted schematicallyin Figure 3.3.

3.2.2 SIMPLE DIFFUSION IS A PURELYPHYSICAL PROCESS

According to Fick’s law (Chapter 1), the rate at whichmolecules in solution diffuse from one region to anotheris a function of their concentration difference. For amembrane-bound cell, Fick’s law may be restated as:

J = PA(Co − Ci) (3.1)

where J is the flux, or amount of solute crossing themembrane per unit time. A is the cross-sectional area ofthe diffusion path, which, in this case, is the area of thecell membrane (in cm2). P is the permeability coefficient.It measures the velocity (in cm−1) with which the solutecrosses that membrane and is specific for a particularmembrane-solute combination. Since the membranebarrier is primarily lipid in character, nonpolar solutemolecules tend to pass through more rapidly than polarmolecules. Membrane lipid bilayers are particularlyimpermeable to most ions. This is because their chargeand high degree of hydration renders ions insoluble inlipids and thus effectively prevents them from enteringthe hydrocarbon phase of membranes. Synthetic lipidbilayers, or artificial membranes, for example, are somenine orders of magnitude less permeable to smaller ionssuch as K+ or Na+ than to water. The permeabilitycoefficient in Fick’s equation thus generally reflectsthe lipid solubility of diffusing molecules. Few solutesof biological importance are nonpolar and only three(O2, CO2, NH3) appear to traverse membranes bysimple diffusion through the lipid bilayer. Water, inspite of its high polarity, also diffuses rapidly throughlipid bilayers; this is because water passes through

FIGURE 3.3 The exchange of ionsand solutes across membranesmay involve simple diffusion, facil-itated diffusion, or active trans-port.

CarrierProteins

ChannelProtein

ADP + PiATP

Facilitateddiffusion

Simplediffusion

Activetransport

Membrane

Passive transport

3.2 Nutrient Uptake 43

water-selective channels called aquaporins. Aquaporinswere discussed in Chapter 1.

Transport by diffusion is a passive process, mean-ing that the transport process does not require a directinput of metabolic energy. The energy for transportby diffusion comes from the concentration or electro-chemical gradient of the solute being transported. Asa consequence, transport by diffusion does not lead toan accumulation of solute against a concentration orelectrochemical gradient.

3.2.3 THE MOVEMENT OF MOSTSOLUTES ACROSS MEMBRANESREQUIRES THE PARTICIPATIONOF SPECIFIC TRANSPORTPROTEINS

In the 1930s, students of membrane transport recog-nized that certain ions entered cells far more rapidlythan would be expected on the basis of their diffu-sion through a lipid bilayer. We now know that thisis because natural membranes contain a large numberof proteins, many of which function as transport pro-teins. Some of these transport proteins facilitate thediffusion of solutes, especially charged solutes or ions,into the cell by effectively overcoming the solubilityproblem. The term facilitated diffusion was coined todescribe this rapid, assisted diffusion of solutes acrossthe membrane. In facilitated diffusion, as in simple dif-fusion, the direction of transport is still determined bythe concentration gradient (for uncharged solute) orelectrochemical gradient (for charged solutes and ions).Facilitated diffusion is also bidirectional and, like sim-ple diffusion, net movement ceases when the rate ofmovement across the membrane is the same in bothdirections. Two major classes of transport proteinsare known. Carrier proteins (also known as carriers,transporters, or simply, porters) bind the particu-lar solute to be transported, much along the lines ofan enzyme–substrate interaction. Binding of the solutenormally induces a conformational change in the carrierprotein, which delivers the solute to the other side of themembrane. Release of the solute at the other surface ofthe membrane completes the transport and the proteinthen reverts to its original conformation, ready to pickup another solute.

Channel proteins are commonly visualized as acharged-lined, water-filled channel that extends acrossthe membrane. Channels are normally identified by theion species that is able to permeate the channel, whichis in turn dependent on the size of the hydrated ion andits charge. Diffusion through a channel is dependent onthe hydrated size of the ion because the associated watermolecules must diffuse along with the ion. The numberof ion channels discovered in the membranes of plantcells is increasing. Currently there is solid evidence forK+, Cl−, and Ca2+ channels, while additional channels

for other inorganic and organic ions are strongly sug-gested. Channel proteins are frequently gated, whichmeans they may be open or closed (Box 3.1). Solutesof an appropriate size and charge may diffuse throughonly when the channel ‘‘gate’’ is open. Two types ofgates are known. An electrically gated channel opens inresponse to membrane potentials of a particular magni-tude. Other channels may open only in the presence ofthe ion that is to be transported and may be modulatedby light, hormones, or other stimuli. The precise mech-anism of gated channels is not known, although it ispresumed to involve a change in the three-dimensionalshape, or conformation, of the protein.

The importance of carriers lies in the selectivitythey impart with respect to which solutes are permittedto enter or exit the cell. Channels, on the other hand,appear to be involved wherever large quantities of solute,particularly charged solutes or ions, must cross themembrane rapidly. Whereas a carrier may transportbetween 104 and 105 solute molecules per second, achannel may pass on the order of 108 ions per second. Itshould also be stressed that large numbers of channelsare not required to satisfy the needs of most cells. Therate of efflux through guard cell K+ channels duringstomatal closure, for example, has been estimated at107 K+ ions sec−1 —a rate that conceivably could beaccommodated by a single channel. Many carrier andchannel proteins are inducible, which means that theyare synthesized by the cell only when there is soluteavailable to be taken up.

3.2.4 ACTIVE TRANSPORT REQUIRESTHE EXPENDITURE OFMETABOLIC ENERGY

Many transport processes, in addition to being rapid andspecific, will lead to an accumulation of solute inside thecell. In other words, the transport process will establishsignificant concentration or electrochemical gradientsand will continue to transport solute against those gra-dients. The transport process involved is known as activetransport. By definition, active transport is tightly cou-pled to a metabolic energy source—usually, althoughnot always, hydrolysis of adenosine triphosphate (ATP).In other words, active transport requires an input ofenergy and does not occur spontaneously. Unlike sim-ple and facilitated diffusion, active transport is alsounidirectional—either into or out of the cell—and isalways mediated by carrier proteins.

Active transport serves to accumulate solutes in thecell when solute concentration in the environment isvery low. When used to transport solute out of thecell, active transport serves to maintain a low internalsolute concentration. Because active transport systemsmove solutes against a concentration or electrochemicalgradient, they are frequently referred to as pumps.

44 Chapter 3 / Roots, Soils, and Nutrient Uptake

Gate

Channel protein

K+ K+

K+

K+

K+

K+

BOX 3.1ELECTROPHYSIO-LOGY—EXPLORINGION CHANNELS

The exchange of ions across cellular membranes isfacilitated by the presence of transmembrane proteinsreferred to as ion channels. Most ion channels arehighly specific for one or a limited number of ionspecies, which can diffuse through an open channel atrates as high as 108 s−1.

Channel proteins may exist in two different con-formations, referred to as open and closed. In the openconformation, the core of the protein forms a pathwayfor diffusion of ions through the membrane (Figure 3.4).A channel that can open and close is said to be gated—inthe open conformation, the ‘‘gate’’ is open and ions arefree to diffuse through the channel. When the gate isclosed, the channel is not available for ion diffusion. Anumber of stimuli, including voltage, light, hormones,and ions themselves, are known to influence the fre-quency or duration of channel opening. The channelprotein is believed to contain a sensor that responds tothe appropriate stimulus by changing the conformationof the protein and opening the gate.

Because ions are mobile and carry a charge,their movement across membranes establishes anelectrical current. These currents, typically on the

Gate

Channel protein

K+ K+

K+

K+

K+

K+

FIGURE 3.4 A gated membrane channel. Gated channelsmay be open, in which case ions are permitted to passthrough the channel, or closed to ion flow. Opening maybe stimulated by changes in membrane potential, thepresence of hormones, or the ion itself.

Electrode

Plasma membrane

Tonoplast

Cell wall

Ion channels

FIGURE 3.5 Changes in electrical properties of the cellrelated to ion flow can be measured by inserting a micro-electrode directly into the cell.

order of picoamperes (pA = 10−12 ampere), can be mea-sured using microelectrodes constructed from finelydrawn-out glass tubing. The first evidence for gatedchannels was based on experiments in which the elec-trode was inserted directly into the cell (Figure 3.5).This method has several limitations. It requires rela-tively large cells and the results reflect the activities ofmany different channels of various types at the sametime. Moreover, when applied to plant cells, the elec-trode usually penetrates the vacuolar membrane as wellas the plasma membrane, thus summing the behavior ofchannels in both membranes.

These problems were largely circumvented bydevelopment of the patch clamp method that permitsthe study of single ion channels in selected membranes.In this technique, the tip of the microelectrode is placedin contact with the membrane surrounding an isolatedprotoplast (a cell from which the cell wall has beenremoved) (Figure 3.6). A tight seal between the electrodeand the membrane is formed by applying a slight suction.The small region of membrane in contact with theelectrode is referred to as the ‘‘patch.’’ Measurementsmay be made in this configuration, with the wholecell attached, or, alternatively, the electrode can bepulled away from the cell. In that case the patchthen remains attached to the electrode tip andcan be bathed in solutions of known composition.Note that the exterior surface of the membrane isin contact with the microelectrode solution (calledthe ‘‘inside-out’’ configuration). Variations in thetechnique permit the orientation of the patch tobe reversed, with the internal surface facing theelectrode solution (the ‘‘outside-out’’ configuration).

3.2 Nutrient Uptake 45

Electrode

Protoplast

Ion channels

Pull electrodeaway fromprotoplast

Suction

Inside - out patch

FIGURE 3.6 Current flow through individual channels canbe measured by the patch-clamp technique. A small pieceof membrane containing a single channel can be isolatedat the tip of a microelectrode. Ions flowing through thechannel carry the current, which can be measured withsensitive amplifiers.

With a sufficiently small electrode tip (ca. 1.0 μm diam-eter), the patch may contain a single ion channel.

With an appropriate electrical circuit, the experi-menter can hold, or ‘‘clamp,’’ the potential difference(voltage) across the patch at some predetermined value.At the same time, the circuit will monitor any currentthat flows through the membrane patch. Typical exper-imental results are shown in Figure 3.7. Figure 3.7Aillustrates a patch-clamp recording for a single K+channel, using an inside-out patch of plasma membraneisolated from a stomatal guard cell protoplast. For thelower trace, the voltage was clamped at 0 mV. Note theappearance of one small, transient current pulse. Forthe upper trace, the voltage was stepped up to +90 mV.The current trace indicates that the channel openedimmediately, in response to the voltage stimulus,and remained open for about 50 milliseconds (ms).

A.

B.

Blue

1 min

40 ms

OmV

OmV (Control)

+90mV 5pA

5pA

FIGURE 3.7 (A) Current generated by a K+-selectivechannel in the plasma membrane of a stomatal guard cellprotoplast. Channel opening was stimulated by apply-ing a 90-mV pulse across the membrane. Note thatthe channel exhibited a transient closure and open-ing before the voltage pulse was terminated. A 0 mVcontrol is shown for comparison. (Reprinted with per-mission from J. I. Schroeder et al., Nature 312:361–362.Copyright 1984, Macmillan Magazines Ltd.) (B) Bluelight stimulates a current in whole-cell membranes,an indication that membrane ion pumps are acti-vated by blue light. (Reprinted with permission fromS. M. Assmann et al., Nature 318:285–287. Copyright1985, Macmillan Magazines Ltd.)

It then spontaneously closed and reopened again forabout 12 ms. This is an example of a voltage-gated ionchannel. Figure 3.7B is a trace obtained with guard cellprotoplasts, using a whole cell configuration. Note thedifference in time scale. In this case, a pulse of blue lightinitiates current flow by activating a blue-light-activatedproton pump.

The patch-clamp technique represents a majoradvance in electrophysiology that is providing insightsinto the mechanisms of plant membrane transport.

FURTHER READING

Molleman, A. 2003. An Introductory Guide to Patch ClampElectrophysiology. Hoboken, NJ: John Wiley & Sons, Inc.

Volkov, A.G. 2006. Plant Electrophysiology: Theory andMethods. Berlin: Springer-Verlag.

46 Chapter 3 / Roots, Soils, and Nutrient Uptake

3.3 SELECTIVE ACCUMULATIONOF IONS BY ROOTS

The selectivity of transport proteins and the ability ofcells to accumulate solutes by active transport togetherenables roots to selectively accumulate nutrient ionsfrom the soil solution. Selective accumulation of ionsby roots is illustrated by a typical set of data shown inTable 3.1. Accumulation refers to the observation thatthe concentrations of some ions inside the cell may reachlevels much higher than their concentration in the sur-rounding medium. This difference in ion concentrationis expressed quantitatively by the accumulation ratio,which can be defined as the ratio of the concentrationinside the cell (Ci) to the concentration outside the cell(Co). Note that in Table 3.1 the internal concentrationof K+ is more than 1,000 times greater than it is inthe bathing medium. In the past, an accumulation ratiogreater than 1 has been considered compelling evidencein favor of active transport, since that solute has evidentlymoved in against a concentration gradient. Conversely,an accumulation ratio less than 1 implies that the solutehas been actively excluded or extruded from the cell.As will be shown below, this is not always the case,especially where charged solutes are involved. Whenassessing solute uptake by cells, it is especially impor-tant to distinguish between uncharged and chargedsolutes.

Ion uptake is highly selective. Note there is virtu-ally no accumulation of Na+ by maize roots (Table 3.1)and accumulation ratios for K+ and NO−

3 are sub-stantially higher than for SO2−

4 . The low concentra-tions of Na+ in plant cells (unlike animal cells) mayresult from limited uptake of Na+ in the first place,but also because Na+ is actively expelled from mostplant cells.

TABLE 3.1 The uptake of selected ions by maizeroots.

Accumulation Ratio

Ion Co (m) Ci (m) [Ci/Co]

K+ 0.14 160 1142

Na+ 0.51 0.6 1.18

NO−3 0.13 38 292

SO2−4 0.61 14 23

Maize roots were bathed in nutrient solutions for four days. Co and Ci

are the ion concentrations of the medium and root tissue, respectively.Ci was measured as the concentration of ions in the sap expressedfrom the roots.From data of H. Marschner, 1986.

3.4 ELECTROCHEMICALGRADIENTS AND IONMOVEMENT

3.4.1 IONS MOVE IN RESPONSE TOELECTROCHEMICAL GRADIENTS

Accumulation ratios for uncharged solutes, such as sug-ars, are relatively straightforward. It can be assumed thatuptake is fundamentally dependent on the differencein concentration on the two sides of the membrane.In other words, for uncharged solutes it is the con-centration gradient alone that determines the gradientin chemical potential (see Chapter 1). The chemicalpotential gradient (�μ) can be expressed by the follow-ing equation, where μi is the chemical potential of theuncharged solute in the cytosol and μo is the chemicalpotential of uncharged solute outside of the cell.

�μ = μi − μo (3.2)

Thus, for the uncharged solute, U, its chemical potentialgradient (�μu) is represented by

�μu = RT ln[Ui]/[Uo] (3.3)

Equation 3.3 simplifies to

�μu = 59 log[Ui]/[Uo] (3.4)

where Ui is the concentration of U in the cytosol andUo is the concentration of U outside of the cell. It isa relatively simple matter to measure experimentallyinternal and external concentrations of the solute andthus calculate the accumulation ratio from which onecan calculate the chemical potential gradient. Fromequations 3.3 and 3.4, it is clear that when Ui > Uo,�μ is a positive value, which indicates that uptake ofU must occur by energy-dependent, active transport.On the other hand, when Ui < Uo, �μ is a negativevalue, which indicates that uptake of U will occur byenergy-independent, facilitated diffusion.

With charged solutes, or ions, the situation is morecomplex and the accumulation ratio is not always a validindication of passive or active transport. Because ionscarry an electrical charge, they will diffuse in responseto a gradient in electrical potential as well as chem-ical potential. Positively charged potassium ions, forexample, will naturally be attracted to a region with apreponderance of negative charges. Consequently, themovement of ions is determined by a gradient that hastwo components: one concentration and one electrical.In other words, ions will move in response to an elec-trochemical gradient, and the electrical properties ofthe cell, or its transmembrane potential, must be takeninto account.

3.4 Electrochemical Gradients and Ion Movement 47

A transmembrane potential (a voltage or potentialdifference across a membrane) develops because of anunequal distribution of anionic and cationic chargesacross the membrane. The cytosol, for example, containsa large number of fixed or nondiffusible charges suchas the carboxyl (—COO−) and amino (—NH+

4 ) groupsof proteins. At the same time, cells use energy to activelypump cations, in particular H+, Ca2+, and Na+, intothe exterior space. The resulting unequal distributionof cations establishes a potential difference, or voltage,across the membrane. The cytosol remains negativerelative to the cell wall space, which accumulates thepositively charged cations.

A simple example illustrates how a transmembranepotential can influence ion movement into and out ofcells (Figure 3.8). In this example, it is assumed that(1) the internal K+ concentration is high relative tothat outside the cell; (2) K+ can move freely across themembrane, perhaps through K+ channels; and (3) theinternal K+ concentration is balanced by a number oforganic anions restrained within the cell. Under theseconditions, it might be expected that K+ will diffuse outof the cell, driven by its concentration gradient, untilthe concentrations of K+ outside and inside the cell areequal. However, as K+ diffuses out of the cell it leavesbehind the nondiffusible anionic charges, thus creatinga charge imbalance and thus a voltage (potential)difference across the membrane. The potential that is

K+

K+

Conc. Gradient

Outside Cell

Low K+

Inside Cell

High K+

K+

K+

K+

K+

K+

K+

K+

K+

K+

-OOC

Protein

-OOC

-OOC

-OOC

-OOC

-OOC

FIGURE 3.8 An electrical potential gradient may drivean apparent accumulation of cations (e.g., K+) againsta concentration gradient. The combination of concen-tration differences and charge differences constitutes anelectrochemical gradient. Thus, potassium ions tend toaccumulate in cells in response to the large number offixed charges on proteins and other macromolecules.

generated by such a combination of nondiffusible anionsand mobile cations is referred to as a Donnan poten-tial. The negative charges tend to pull the positivelycharged potassium ions back onto the cell. As a result,equilibrium is achieved not when the concentrations ofK+ are equal on both sides of the membrane, but whenthe membrane potential difference reaches a value suchthat the force of the concentration gradient pulling K+out of the cell is balanced by the force of the electricalgradient pulling K+ back into the cell. Under thesecircumstances, the cell will maintain a high internalK+ concentration and the accumulation ratio will begreater than unity, yet the movement of potassiumion is solely by passive diffusion. Because the unequaldistribution of K+ at equilibrium results from a Donnanpotential, it is an example of Donnan equilibrium.

Anion distribution would also be influenced bythe membrane potential, but in the opposite direction.Anions would be repelled by the preponderance of inter-nal negative charges and attracted by the preponderanceof external positive charges, thus leading to an accumula-tion ratio less than unity. It is clear from these examplesthat an accumulation ratio other than unity does notnecessarily mean that active transport is involved.

Transmembrane potentials can be measured witha microelectrode made from finely drawn-out glasstubing. With the aid of a microscope, the electrode isinserted into the vacuole of a cell. A reference electrodeis placed in the medium surrounding the cell. Thedifference in potential between the two electrodes canbe measured with the aid of a sensitive voltmeter. It isby no means an easy technique, but many experimentershave become quite proficient with it. Potentialsmeasured in this way are commonly in the range of−100 to −130 mV for young roots and stems, althoughpotentials as high as −200 mV have been recorded forsome algal cells. The cytosol is always negative withrespect to the surrounding medium. Such potentialsdo not require a large charge imbalance. As few as oneunbalanced charge in a population of a million ions issufficient to generate a potential of 100 mV.

3.4.2 THE NERNST EQUATION HELPSTO PREDICT WHETHER AN IONIS EXCHANGED ACTIVELY ORPASSIVELY

To predict the distribution of a charged ion, C, across amembrane, we have to take into account the concentra-tion difference of the ion across the membrane as well asthe contribution of electrical potential difference (�Ec)across the membrane. Thus, equation 3.3 simplifies to

−z �Ec = 59 log[Ci]/[Co] (3.5)

48 Chapter 3 / Roots, Soils, and Nutrient Uptake

TABLE 3.2 The uptake of selected ions by roots of pea (Pisum sativum) and oat (Avena sativa). TheNernst equation was used to predict the internal concentration (Ci) assuming cell membrane potentials of−110 mV and −84 mV for pea and oat roots, respectively. The accumulation ratio was calculated on thebasis of the measured (or actual) Ci. The symbol E under probable uptake mechanism refers to activeexclusion from the root. The symbol U denotes active uptake by the roots.

ActualPredicted Actual Accumulation Probable Uptake

Ion Co Ci Ci Ratio Predicted Mechanism

Pea Root

K+ 1.0 74 75 75 1.01 DiffusionNa+ 1.0 74 8 8 0.108 ECa2+ 1.0 5400 1.0 1.0 0.00018 ENO−

3 2.0 0.027 28 14 1037 UH2PO−

4 1.0 0.014 21 21 1500 USO2−

4 0.25 0.000047 9.5 38 202,127 U

Oat Root

K+ 1.0 27 66 66 2.4 Diffusion (?)Na+ 1.0 27 3 3 0.11 ECa2+ 1.0 700 1.5 1.5 0.0021 ENO−

3 2.0 0.076 56 28 741 UH2PO−

4 1.0 0.038 17 17 447 USO2−

4 0.25 0.000036 2 8 5,555 U

From data of Higinbotham et al., 1967.

where z is the valency or absolute charge of the ion, C,and Ci and Co are the concentrations of this ion in thecytosol and outside the cell respectively. Equation 3.5,which shows relationship between transmembranepotential gradient and ion distribution across themembrane is called the Nernst equation where �Ecis the electrical potential difference (also known asthe Nernst potential). The value of z for a univalentcation, for example, would be 1 while for calcium ormagnesium it would be 2. For chloride or nitrate itwould be −1 and for sulphate it would be −2.

The Nernst equation is useful because it allowsus to make certain predictions about the equilibriumconcentrations for ions inside the cell (Ci) when iontransport is due to facilitated diffusion. In order to applythe equation, it is necessary to measure the transmem-brane potential and the concentrations of ions bothinside and outside the cell. If the actual concentrationsdeviate significantly from those predicted by the Nernstequation, it may be considered evidence that eitheractive uptake or active expulsion of the ions is involved.In other words, the Nernst equation can be used todetermine the probability of whether an ion is activelyor passively distributed across the membrane. If the mea-sured internal concentrations are approximately equalto the calculated Nernst value, it can be assumed thatthe ion has been distributed passively. If the measured

concentration is greater than predicted, active uptake isprobably involved, and, if lower, it is likely that the ionis being actively expelled from the cell.

Application of the predictive value of the Nernstequation is illustrated by the experiment of Higin-botham and colleagues shown in Table 3.2. At equilib-rium the measured membrane potentials were −110 mVfor pea roots and −84 mV for oat roots. Co, the concen-tration of ions in the external solution, was known andthe predicted value of Ci was calculated from the Nernstequation. The ratio of the predicted value to the actualvalue is a measure of how well the Nernst relationshipapplies to that particular ion. Note that the accumula-tion ratios for almost all of the ions are greater than1.0, indicating some degree of accumulation in the cell.Only in the case of K+ was the ratio of predicted concen-tration to actual concentration near 1.0. This indicatesthat K+ is near electrochemical equilibrium and wasprobably accumulated passively, at least in pea roots.There appears to be a possibility of some active accu-mulation of K+ by oat roots. Cellular concentrationsof both Na+ and Ca2+ are lower than predicted. Sinceother evidence supports the existence of Na+ and Ca2+pumps in the membrane, these ions probably enteredpassively down their electrochemical gradients but werethen actively expelled. Internal concentrations for allthree anions are much higher than predicted, indicating

3.5 Electrogenic Pumps are Critical for Cellular Active Transport 49

they are actively taken up by the cells. This is under-standable, since energy would be required to overcomethe normal transmembrane potential and move neg-atively charged ions into the predominantly negativeenvironment inside the cell.

Although these results are indicative of active andpassive transport, further tests would help to confirm theconclusion in each case. Since active transport requiresa direct input of metabolic energy, it is sensitive tooxygen and respiratory poisons. Thus reduced uptakeof a particular ion in the absence of oxygen or inthe presence of respiratory inhibitors such as cyanide or2,4-dinitrophenol would be evidence in support of activetransport. Even this evidence is not always compelling,however, since effects of inhibitors may indirectly influ-ence nutrient uptake. For example, even transport bydiffusion ultimately requires an expenditure of metabolicenergy, if only to establish and maintain the organizationof membranes and other properties of the cell that maketransport possible. It is not always a simple matter todistinguish between direct and indirect involvement ofenergy. The criteria for active transport usually requirethat the solute distribution not be in electrochemicalequilibrium and that there be a quantitative relation-ship between energy expended and the amount of solutetransported.

Finally, it should be noted that when ion con-centrations are known, the Nernst equation can also beused to estimate the transmembrane potential, or Nernstpotential, contributed by that ion. At steady state, however,calculation of the membrane potential is complicated bythe fact that many different ion species, each with adifferent permeability, are simultaneously crossing themembrane in both directions. The individual contribu-tions of all ion gradients must consequently be takeninto account and summed in order to arrive at the over-all potential for the cell. In practice, however, K+, Na+,and Cl− are the dominant ions. These ions have thehighest permeabilities and concentrations in plant cellsand a reasonable estimation of transmembrane potentialcan be based on these three ions alone.

3.5 ELECTROGENIC PUMPS ARECRITICAL FOR CELLULARACTIVE TRANSPORT

3.5.1 ACTIVE TRANSPORT IS DRIVENBY ATPASE-PROTON PUMPS

Energy to drive active transport comes chiefly fromthe hydrolysis of ATP. The energy-transducing mem-branes of chloroplasts and mitochondria contain large,multiprotein ATPase complexes (Chapters 7, 10). Thechloroplast and mitochondrial ATPases, known asF-type ATPases, utilize the energy associated with anelectrochemical proton gradient across a membrane to

ATP

Outside

1012

39

8

7

D

Inside

FIGURE 3.9 A model for the plasma membraneH+-ATPase. The enzyme is a single chain with 10hydrophobic, membrane-spanning domains. Only threeare shown here as helical coils, while the remaining sevenare schematically represented as cylinders. Linking adja-cent membrane-spanning domains are hydrophilic loopsthat project into the cytosol (inside) and the cell wallapoplast (outside). The ATP-binding site is an asparticacid residue (D) located on the hydrophilic loop betweenthe fourth and fifth membrane-spanning domains. TheH+-binding site is located in the hydrophobic domain.Hydrolysis of ATP at the binding site is thought tochange the conformation of the enzyme, thereby expos-ing the H+-binding site to the outside of the membranewhere the H+ is released. (Compare with Figure 5.9.)

drive ATP synthesis. ATPase complexes are also found inthe plasma and vacuolar membranes of cells and possiblyother membranes as well. These ATPases are calledATPase-proton pumps (or, H+-ATPase). Known asP-type ATPases, the plasma membrane ATPase-protonpumps are structurally distinct (Figure 3.9) and operatein reverse of the F-type. Rather than synthesizingATP, the plasma membrane ATPases hydrolyze ATPand use the negative free energy to ‘‘pump’’ protonsfrom one side of the membrane to the other againstan electrochemical gradient. An ATPase-proton pumpthus serves as a proton-translocating carrier proteinand the free energy of ATP hydrolysis is conserved inthe form of a proton gradient across the membrane.This proton gradient (�pH), together with the normalmembrane potential (��), contributes to a protonmotive force (pmf) that tends to move protons backacross the membrane (Equation 3.6).

pmf = �� − 59 �pH (3.6)

It is generally conceded that the proton motive forceestablished by pumping protons across membranes is theprimary source of energy for a variety of plant activities.We will revisit this equation and the concept of a protonmotive force in our discussion of ATP synthesis in thechloroplast and the mitochondrion (Chapter 5).

50 Chapter 3 / Roots, Soils, and Nutrient Uptake

H+ H+

H+H+

ADP

ATP

ADP

ATP

Plasma membrane

cVacuole

Cytosol

Mitochondrion

A−

S

H+

A−

H+

H+

C+

C+

C+

pH = 5.5 pH = 7.0, −120mV

-90mVpH 5.5

H+

b

e

d

a

FIGURE 3.10 Schematic diagram relating the activity ofa membrane ATPase-proton pump to solute exchange.The proton pump (a) uses the energy of ATP to estab-lish both a proton gradient and a potential difference(negative inside) across the membrane. The energy ofthe proton gradient may activate an ion channel (b), ordrive the removal of ions from the cell by an antiport car-rier (c), or drive the uptake of ions or uncharged soluteby a symport carrier (d, e). Similar pumps and carriersoperate across the vacuolar membrane. C+, cation; A−,anion; S, uncharged solute.

Included are activities such as active transportof solutes (cations, anions, amino acids, and sugars),regulation of cytoplasmic pH, stomatal opening andclosure, sucrose transport during phloem loading, andhormone-mediated cell elongation.

A schematic model relating ATPase-proton pumpsto solute exchange across membranes is shown inFigure 3.10. The ATP required to drive the pump isultimately derived from oxidative phosphorylation inthe mitochondria. The proton-translocating protein isshown extending across the plasma membrane, with itsATP binding site on the cytosolic side. Hydrolysis ofthe ATP results in the translocation of one or moreprotons from the cytosol to the surrounding apoplasticcell wall space.

There are several particularly interesting conse-quences of the ATPase-proton pump. First, a single ionspecies is translocated in one direction. This form oftransport is consequently known as a uniport system.Second, because the ion transported carries a charge,an electrochemical gradient is established across themembrane. In other words, the ATPase-proton pumpis electrogenic—it contributes directly to the negative

potential difference across the plasma membrane. Infact, the electrogenic proton pump is a major factorin the membrane potential of most plant cells. Fromequation 3.5, a tenfold difference in proton concentra-tion (one pH unit) at 25◦C contributes 59 mV to thepotential. Since the proton gradient across the plasmamembrane is normally on the order of 1.5 to 2 pH units,it can account for approximately 90 to 120 mV of thetotal membrane potential. Third, since the ions translo-cated are protons, the ATPase-proton pump establishesa proton gradient as well as an electrical gradient acrossthe membrane. Energy stored in the resulting electro-chemical proton gradient (or the proton motive force) canthen be coupled to cellular work in accordance withMitchell’s chemiosmotic hypothesis (see Chapter 5). Indeed,this is an excellent demonstration of how chemiosmoticcoupling is not restricted to ATP synthesis in chloro-plasts and mitochondria but can be used to performother kinds of work elsewhere in the cell.

Several ways of coupling the electrochemical protongradient to solute movement across the membrane areillustrated in Figure 3.10. In the first case, the electro-genic pump contributes to the charge-dependent uptakeof cations through ion-specific channels (Figure 3.10)(b)). Second, the return of protons to the cytosol can becoupled with the transport of other solute molecules atthe same time, or cotransport. Transport of both ions ismediated by the same carrier protein and the movementof the second solute is obligatorily coupled to the inwardflux of protons down their electrochemical gradient.If the second ion moves in opposite direction from theproton, the method of cotransport is referred to morespecifically as antiport. In Figure 3.10 (c), for example,proton flux into the cell is shown coupled to the effluxof other cations out of the cell. Here the energy of theproton electrochemical gradient is used to maintain lowinternal concentrations of specific cations. Any cationsthat do chance to ‘‘leak’’ into the cell, no doubt passivelythrough ion channels, are thus pumped out againsttheir electrochemical gradient. If the two solutes movein the same direction at the same time, the method ofcotransport is referred to as symport. Two examples areshown in Figure 3.10 (d, e). In the first example, protonflux into the cell is coupled with the uptake of anions(A−) against their electrochemical gradient. In thesecond example of symport, the proton gradient can beused to power the uptake of uncharged solutes (S), suchas sugars. All three examples of cotransport are formsof active transport mediated by specific carrier proteins.

3.5.2 THE ATPASE-PROTON PUMPSOF PLASMA MEMBRANES ANDVACUOLAR MEMBRANES AREDIFFERENT

Much of the pioneering experimental work on mem-brane ATPase has been conducted with small, spherical

3.5 Electrogenic Pumps are Critical for Cellular Active Transport 51

vesicles obtained from isolated cellular membranes.When membranes are disrupted, the pieces naturallyseal off to form vesicles because of their strong-ly hydrophobic nature. While the technique is relativelystraightforward, the preparation of vesicles from a singlemembrane source, and thus containing a single typeof ATPase, presents some difficulties. Contaminationby chloroplast ATPase can be avoided by isolatingthe membranes from dark-grown, etiolated tissue,while mitochondria can usually be separated fromother membranes by differential centrifugation. It ismore difficult, however, to separate plasma membranesfrom other cellular membranes such as the vacuolarmembrane and, consequently, many of the early studieswere characterized by inconsistent results from differentlaboratories. These inconsistencies were resolved whenit became clear that the membrane preparations oftencontained two types of electrogenic ATPase-protonpumps: one associated with the plasma membrane andone with the vacuolar membrane. Improved techniqueshave enabled at least partial separation of the twomembranes by density gradient centrifugation and it isnow possible to characterize their respective ATPases.The plasma membrane–type proton-pumping ATPaseis characteristically inhibited by vanadate ion (VO−

3 ) butis generally insensitive to other anions such as NO−

3 .Vanadate competes with phosphate for binding sites,indicating that ATP transfers a phosphate group to theATPase protein (Figure 3.9). The resulting energy-richphosphoenzyme then undergoes a conformationalchange that exposes the proton-binding site to theoutside. Evidence thus far indicates that a single protonis translocated for each ATP hydrolyzed.

The vacuolar-type ATPase-proton pump (orV-type) differs from the plasma membrane–type inseveral ways. It is, for example, insensitive to vanadatebut strongly inhibited by nitrate. In this respect it issimilar to mitochondrial, or F-type, ATPase, whichis also insensitive to vanadate. Should the preparationbe contaminated with any mitochondrial ATPase,however, its activity can be blocked by includingoligomycin or azide in the assay medium. Both inhibitmitochondrial ATPase without affecting the activityof the tonoplast-type. Structurally, the vacuolar-typeATPase is also more similar to the mitochondrialF-type ATPase than to the plasma membrane ATPase.Like the F-type, the V-type can be separated into acomplex of hydrophobic subunits embedded in themembrane (analogous to F0) and a complex of soluble,hydrophilic subunits (analogous to F1). Althoughthe soluble complex contains an ATP-binding site,insensitivity of the V-type ATPase to vanadate suggeststhat it does not form a phosphorylated intermediate.The vacuolar version also appears to differ in that ittransports two protons for each molecule of ATP thatis hydrolyzed. The function of the vacuolar ATPase is

to pump protons from the cytosol into the vacuole, thusaccounting for the fact that the potential of the vacuoleis more positive than the cytosol by some 20 to 30 mV(Figure 3.10). In extreme cases, large pH gradients canbe maintained across the vacuolar membrane and thevacuolar sap may become quite acidic. For example, thepH of lemon juice (which is predominantly vacuolarsap) is normally about 2.5. Like the plasma membranepump the vacuolar pump is also electrogenic, exceptthat the accumulated protons serve to reduce thepotential of the vacuole relative to the cytosol. Theresulting potential difference serves to drive anions(e.g., Cl− or malate) into the vacuole, which is lessnegative than the cytosol. The electrochemical protongradient can also be used to drive cations (e.g., K+ orCa2+) into the vacuole by an antiport carrier. Both ofthese activities make important contributions to theturgor changes that drive stomatal guard cell movementand the specialized motor cells that control nyctinasticresponses (Chapter 23).

In addition to the H+-ATPases, some membranes,such as the plasma membranes, chloroplast envelope, theendoplasmic reticulum, and vacuolar membrane, alsocontain calcium-pumping ATPases (Ca2 + − ATPases).Ca2+-ATPases couple the hydrolysis of ATP with thetranslocation of Ca2+ across the membrane. In the caseof the plasma membrane, the calcium is pumped outof the cytosol. This serves to keep the cytosolic Ca2+concentration low, which is necessary in order to avoidprecipitating phosphates and to keep Ca2+-depen-dent signaling pathways operating properly (Chapters 16and 17).

3.5.3 K+ EXCHANGE IS MEDIATED BYTWO CLASSES OF TRANSPORTPROTEINS

Since the 1950s, the uptake of K+ into cells—especiallyroot cells—has been studied more thoroughly than anyother ion species. Much of the early work was carriedout by Emanuel Epstein, who pioneered the use of86Rb+, a radioactive K+ analog, to follow K+ uptakein low-salt roots. Epstein was also the first to treat iontransporters as enzymes and to analyze their data usingthe methods of classical enzyme kinetics. An analysisof the initial rate of K+ absorption at different externalconcentrations showed that K+ absorption is biphasic.On the basis of these results, Epstein proposed thatthere were two types of K+ transport systems in plantcells: a high- affinity transport system (HAT) that isactive at low K+ concentrations (≤ 200 μM), and alow-affinity transport system (LAT) that is active at highK+ concentrations. Such transport systems have nowbeen identified for myriad macronutrient ions includingCa2 + , NO−

3 , SO2−4 and PO2−

4 . A consistent feature of

52 Chapter 3 / Roots, Soils, and Nutrient Uptake

all HATs is they mediate a slow rate of ion uptake whenthe external concentrations of the ion are low. Thismeans that HATs exhibit a low capacity for ion uptake.However, although their capacity for ion uptake is low,their efficiency for ion uptake is very high because oftheir high binding affinity for the specific ion. Thus, theuptake kinetics for HATs exhibit nonlinear, saturationkinetics. In contrast, although LATs exhibit low affinityfor ion uptake in a linear concentration dependence, thecapacity of LATs for uptake is high. The ion concentra-tion which typically induces a transition from LATs toHATs is about 1 mM for most macronutrient ions. Bio-physical and molecular genetic studies have confirmedthe existence of multiple transporters with differentsubstrate affinities. Mosts physiological, biochemical,and molecular studies of macronutrient ion transporthave focused on the characterization of HATs. How-ever, given that agricultural soil concentrations of K+,NO−

3 , and NH+4 may exceed 1 mM, LATs may also be

important in maintaining plant productivity under fieldconditions.

Experiments using microelectrodes that measurecytoplasmic K+ concentration and membrane poten-tial simultaneously in single root cells have confirmedthat when the external concentration is low, thereis a strong K+ electrochemical gradient across theplasma membrane. In order for K+ to move into thecell under those conditions, the high-affinity uptakesystem must be driven by an active transport mech-anism. This high-affinity uptake system is probablya H+-ATPase-linked K+-H+ symporter. Patch-clampstudies, on the other hand, have established that thelow-affinity uptake system involves channels that moveK+ either into (K+

in channels) or out (K+out channels)

of the cell. While the focus tends to be on K+in chan-

nels, channels that allow K+ to move out of the cellare also important in controlling osmotic adjustment,maintaining stomatal function, and driving nyctinasticmovements of leaves (Chapter 23).

K+ transporters are highly efficient and, conse-quently, their relative abundance in the cell membranesis very low. This makes it difficult to isolate andpurify transporters by traditional biochemical tech-niques. Some genes that encode K+ transporters andother plant transport proteins have been identified bycloning the genes in transport-deficient mutants of yeast(Saccharomyces cerevisiae). A yeast mutant deficient of ahigh-affinity K+ transporter, for example, will growon a medium with a high K+ concentration, but not ona low-concentration medium. However, transformationof the yeast with KAT1 (a gene encoding a high-affinityK+ transporter from the guard cells and vascular tissueof Arabidopsis), will restore the capacity of the yeast togrow on low K+. Experiments of this type have ledto the identification of at least a dozen putative K+

transporters as well as transporters for sugars, aminoacids, NH+

4 , and SO2−4 .

3.6 CELLULAR ION UPTAKEPROCESSES AREINTERACTIVE

Deprivation of the macronutrients nitrogen (N),phosphorus (P), potassium (K), and sulfur (S) can bea limiting factor for plant growth and survival undernatural conditions. How do plant root cells sensechanges in soil nutrient macronutrient availability andhow is this signal transduced by the root cell to elicit anappropriate response at the molecular, biochemical, andphysiological levels to adjust the rate of growth to matchthe macronutrient availability? Although the cellularnetworks involved in the sensing and signalling of lim-itations in macronutrient resource availability are notwell defined, it is clear that the deficiency in one nutrientcan affect the uptake of another nutrient. For example,a decrease in SO2−

4 availability can disrupt nitrogenmetabolism resulting in the accumulation of NO−

3 in theleaves. Similarly, NH+

4 availability is known at affect K+uptake. Numerous molecular studies indicate that K+limitation represses the expression of NO−

3 transportersbut stimulates the expression of NH+

4 transporters.PO2−

4 deprivation is one of the most studied phe-nomena associated with nutrient uptake. Recent genemicro-array studies indicate that phosphorus deficiencyin plants causes a plethora of transcriptional responses,including the up-regulation of SO2−

4 transporters as wellas iron transporters. PII proteins (Chapter 11) are keyregulators of cellular NO−

3 supply to ensure a balancebetween C- and N-metabolism. Thus, the mechanismscontrolling the uptake of these macronutrients nutrientsdo not occur necessarily independently of one anotherbut rather are interactive. The complex, interactiveeffects of macronutrient deprivation on gene transcrip-tion and cellular physiology indicate that nutrient uptakeis best described as a complex network of pathways.As a consequence, there must be molecular cross talkbetween the different ion uptake mechanisms to coun-teract potential imbalances caused by deficiencies innutrients.

3.7 ROOT ARCHITECTURE ISIMPORTANT TO MAXIMIZEION UPTAKE

Even though the uptake of ions by roots is essentially acellular problem, the organization of roots at the tissuelevel cannot be totally ignored. The organization and

3.7 Root Architecture is Important to Maximize Ion Uptake 53

architecture of roots are such that they can absorb somemineral salts without them ever entering a cell.

3.7.1 A FIRST STEP IN MINERALUPTAKE BY ROOTS IS DIFFUSIONINTO THE APPARENT FREE SPACE

Most nutrient uptake studies are carried out with‘‘intact’’ tissues, such as excised barley roots. A typicalpattern for the uptake of Ca2+ by low-salt barley rootsis illustrated by the kinetic diagram in Figure 3.11.Note that initially, usually within the first few minutes,uptake of Ca2+ is very rapid. It then settles into a slowbut steady accumulation over time. If at some pointthe roots are transferred to a large volume of solutionlacking calcium, Ca2+ will be lost from the root into thebathing solution as shown by the dashed lines. When thebathing solution is distilled water, the quantity of ionslost is usually less than the quantity taken up during theinitial rapid phase. If the roots are then transferred fromdistilled water to a bathing solution containing anothercation, say Mg2+, an additional quantity of Ca2+ will belost from the tissue. If volumes of the bathing solutionsare sufficiently large, the total quantity of Ca2+ lostfrom the tissue will approximately equal the quantitytaken up during the initial rapid phase.

The kinetics of calcium uptake and release in thisexperiment can be interpreted as follows. Assume that

SO4

2

Distilled water

Upt

ake

(−)

or lo

ss (

− −

−)

Time (hrs)

1 2 3 4 50

FIGURE 3.11 Typical kinetics for the uptake of Ca2+ intoroots. When low-salt roots are placed in a solution ofcalcium chloride, an initial rapid uptake is followed bya slower but steady accumulation of calcium ion. If theroots are then transferred to a large volume of distilledwater, some of the calcium diffuses out of the roots.Transfer to a strong magnesium solution releases addi-tional calcium into the medium. The total amount ofcalcium released is equivalent to the amount taken up byfree diffusion during the initial rapid phase.

there is a fraction of the root tissue volume, called appar-ent free space (AFS), that is not separated from theenvironment by a membrane or other diffusion barrier.Because there are no barriers, Ca2+ in the surroundingmedium would have access to the apparent free spaceby simple diffusion. When root tissue is immersed inthe calcium solution, Ca2+ will rapidly diffuse into theAFS until the Ca2+ concentration in the AFS reachesequilibrium with the bathing solution. This accounts forthe initial, rapid uptake of calcium. Thereafter, calciumions are more slowly but steadily transported across thecell membrane and accumulated by the tissue. Whenthe roots are transferred to distilled water, some of theCa2+ ions present in the free space are free to diffuseback into the surrounding solution—and they will do sountil equilibrium is again reached. The Ca2+ taken upby the cells, having already been transported across thecell membrane, is not free to diffuse back and remainsin the cells.

A further loss of calcium when the roots are trans-ferred to the solution containing magnesium ions istaken as evidence that the tissue behaves as a cationexchange material. That is to say, the AFS matrix, pri-marily cell wall components, is negatively charged andholds some cations by electrostatic attraction just as soilcolloids do. These adsorbed ions are not free to diffuseout of the tissue into distilled water, but can be displacedby other cations, such as magnesium in the examplegiven above. Thus, apparent free space describes thatportion of the root tissue that is accessible by free diffu-sion and includes ions restrained electrostatically due tocharges that line the space.

A variety of techniques have been developedto measure the root volume given over to AFS. Inprinciple, the volume of the AFS can be estimated by thefollowing experiment. A sample of roots weighing 1.0 gwas immersed in a solution containing 20 μmoles ml−1

potassium sulfate (K2SO4). The roots were thenremoved from the solution, blotted to remove excesssolution, and placed in a large volume of distilledwater. It was found that 4.5 μmoles of sulphate werereleased from the roots into the distilled water. If it isassumed that sulphate in the AFS was in equilibriumwith the external solution, that is, 20 μmoles ml−1,then the volume occupied by the sulphate in the tissuecan be calculated: 4.5 μmole/20 μmoles ml−1 = 0.22 ml.Thus, the volume of root tissue freely accessible bydiffusion is 0.22 ml. By further assuming that 1 g ofroot tissue occupies approximately 1 ml of volume, theproportion of tissue freely accessible by diffusion isapproximately 22 percent by volume. Estimates for thevolume occupied by AFS do vary, depending on thespecies, conditions under which the roots were grown,whether the measurements are corrected for surfacefilms, and so forth. Still, most measured values for AFStend to fall in the 10 to 25 percent range.

54 Chapter 3 / Roots, Soils, and Nutrient Uptake

3.7.2 APPARENT FREE SPACE ISEQUIVALENT TO THE APOPLASTOF THE ROOT EPIDERMAL ANDCORTICAL CELLS

Exactly what constitutes AFS in a root? If it is assumedthat AFS is the volume of the root that is accessibleby free diffusion, then it probably consists of the cellwalls and intercellular spaces (equivalent to the apoplas-tic space) of the epidermis and cortex. These are theregions of the root that can be entered without crossinga membrane. In most cases there is a strong correlationbetween the calculated volume of AFS and the calcu-lated volume of cell walls in the cortex of the root.Furthermore, the cation exchange capacity of the AFScan be traced to the carboxyl groups (—COO−) associ-ated with the galacturonic acid residues in the cell wallpectic compounds.

Almost certainly, the AFS stops at the endodermiswhere, in most roots, the radial and transverse wallsdevelop characteristic thickenings called the Casparianband (see Chapter 2, Figure 2.10). The Casparian bandis principally composed of a complex mixture of hydro-phobic, long-chain fatty acids and alcohols calledsuberin. These hydrophobic substances impregnatethe cell wall, filling in the spaces between the cellulosemicrofibrils. They are, in addition, strongly attachedto the plasma membrane of the endodermal cells. Thehydrophobic, space-filling nature of the Casparian bandalong with its attachment to the membrane greatly

reduces the possibility that ions or small hydrophilicmolecules can pass between the cortex and stele withoutfirst entering the symplast. This means, of course, thatthey must pass through the plasma membrane of thecortical or endodermal cells and are, consequently,subject to all of the control and selectivity normallyassociated with membranes.

3.8 THE RADIAL PATH OF IONMOVEMENT THROUGHROOTS

3.8.1 IONS ENTERING THE STELEMUST FIRST BE TRANSPORTEDFROM THE APPARENT FREESPACE INTO THE SYMPLAST

Rapid distribution of nutrient ions throughout the plantis accomplished in the xylem vessels. In order to reachthese conducting tissues, which are located in the cen-tral core, or stele, of the root, the ions must move ina radial path through the root. The path these ionsmust follow is diagrammed in Figure 3.12. For thesepurposes we may consider the root as consisting ofthree principal regions. The outermost region consistsof the root epidermis (often referred to as the rhizo-dermis) and the cortical cells. The innermost regionconsists of vascular tissues—the vessel elements andassociated parenchyma cells—which are of particular

Casparian band

Casparian band

Vesselelement

Epidermis/cortex

Soi

l sol

utio

n

Endodermis Stele

VacuoleVacuole

Vacuole

FIGURE 3.12 The radial paths of ion movement through a root. Arrows indicate thealternative paths that may be taken by nutrient ions as they move from the soilsolution into the vascular elements in the stele. Arrows with circles indicate activetransport of ions across plasma membranes.

3.8 The Radial Path of Ion Movement Through Roots 55

interest for our discussion. Separating the two is theendodermis with its suberized Casparian band.

Ion uptake begins with free diffusion into the appar-ent free space. As noted in the previous section, theapparent free space is equivalent to the apoplast out-side the endodermis and the Casparian band effectivelyprevents further apoplastic diffusion through the endo-dermis into the stele. Hence the only possible route forions to pass through the endodermis is to enter the sym-plast by some carrier- or channel-mediated transport atthe cell membrane. This may occur either on the outertangential wall of the endodermal cell itself or throughany of the epidermal or cortical cells. Regardless ofwhich cell takes up the ions, symplastic connections(i.e., plasmodesmata) facilitate their passive movementfrom cell to cell until they arrive at a xylem parenchymacell in the stele. At this point the ions may be unloadedinto the xylem vessels for long distance transport to theleaves and other organs.

3.8.2 IONS ARE ACTIVELY SECRETEDINTO THE XYLEM APOPLAST

With the exception of the very tip of the root where theyoung xylem vessel elements are still maturing, func-tional xylem is part of the apoplast. The interconnectedvessel elements are devoid of cytoplasm and consistonly of nonliving tubes filled with an aqueous solution.Release of ions into the xylem thus requires a transferfrom the symplast into the apoplast. At one time, it wasthought that this transfer was simply a passive leakage,but it is now clear that ions are actively secreted fromxylem parenchyma cells. Although there is some con-flicting evidence, ion concentration in the apoplast of thestele is generally much higher than in the surroundingcortex. This suggests that ions are being accumulatedin the xylem against a concentration gradient, presum-ably by an energy-dependent, carrier-mediated process.It is also interesting to speculate, in this regard, thatthe Casparian band also functions to prevent loss ofions from the stele by blocking their diffusion down aconcentration gradient.

In addition to working uphill against a con-centration gradient, delivery of ions into the xylemvessels is sensitive to metabolic inhibitors such ascarbonyl-cyanide-m-chlorophenylhydrazone (CCCP),which uncouples ATP formation. It is interestingthat ion transport into the xylem is also sensitive tocycloheximide, an inhibitor of protein synthesis, butuptake into the root, at least initially, is not affected.Two plant hormones (abscisic acid and cytokinin) havea similar effect. Whether inhibitors of protein synthesisand hormones are affecting symplastic transportthrough the endodermis or unloading of ions from theendodermis into the xylem is not certain, but theseresults at least raise the possibility that ion release into

the vessels is a different kind of process than ion uptakeby the roots.

3.8.3 EMERGING SECONDARY ROOTSMAY CONTRIBUTE TO THEUPTAKE OF SOME SOLUTES

The possibility remains that a limited portion ofion uptake may be accomplished entirely throughthe apoplast, at least in some roots. More basalendodermal cells—the distance from the tip is variable,but measured in centimeters—are characterized byadditional suberin deposits that cover the entire radialand inner tangential wall surfaces. This would seemto present an additional barrier to apoplastic flow.However, in some plants, a small number of endodermalcells, called passage cells, remain unsuberized. Passagecells might represent a major point of entry for solutesinto the stele.

Apoplastic continuity between the cortex and stelemay also be established at the point of lateral root forma-tion. One series of experiments, for example, followedthe path of fluorescent dyes into the vascular tissues andshoots of corn (Zea mays) and broad bean (Vicia faba)seedlings. These dyes were chosen because they cannotbe taken up by cells and thus are normally confinedto the apoplast, The point of dye entry was traced torecently emerged secondary roots. These branch rootsarise in the pericycle, a layer of cells immediately insidethe endodermis. The emergence of the root primor-dia through the endodermis disrupts the continuity ofthe Casparian band and establishes, at least temporar-ily, the apoplastic continuity required to allow diffusionof the dye into the vascular tissue. Continuity of theapoplast through passage cells and secondary roots hasbeen cited to explain increased calcium uptake in certainregions of corn roots. It may also help to account forthe fact that a plant appears to contain virtually everyelement that is found in its environment, even thosenot known to be essential or not accumulated by plantcells.

The uptake of ions is not uniform along the lengthof the root. As shown in Table 3.3, uptake of calcium ishighest in the apical 3 cm of the root while potassium istaken up in roughly equivalent amounts along the first15 cm. Moreover, most of what is taken up in the tip(almost two-thirds of the calcium and three-fourths ofthe potassium) remains in the root. The proportion ofions translocated to the shoot increases with increasingdistance from the tip. It is also interesting that whencalcium is taken up further along the root (12 to 15 cmfrom the tip), it is translocated to the shoot but notto the tip. Clearly, although substantial progress hasbeen made in several laboratories, the transport of ionsthrough roots and into the xylem remains a complex andchallenging field of study.

56 Chapter 3 / Roots, Soils, and Nutrient Uptake

TABLE 3.3 Uptake and translocation of potassium and calcium as a function of position along a corn root.

Percent Translocated to:Zone of Total Percent

application1 Ion Uptake2 Retained Root Tip Shoot

0–3 K+ 15.3 75 — 25Ca2+ 6.3 63 — 37

6–9 K+ 22.7 17 19 64Ca2+ 3.8 42 — 58

12–15 K+ 19.5 10 10 80Ca2+ 2.8 14 — 86

1Distance from root tip, cm.2Uptake expressed as microequivalents per 24 hours.Based on data of H. Marschner and C. Richter, 1973, Z. Pflenzenernaehr, Bodenkd, 135:1–15.

3.9 ROOT-MICROBEINTERACTIONS

The influence of living roots extends well beyond theimmediate root surface into a region of the soil defined asthe rhizosphere. A principal manifestation of this influ-ence is the numerous associations that develop betweenroots and soil microorganisms, especially bacteria andfungi. Root-microbe associations can at times be quitecomplex and may involve invasion of the host rootby the microorganism. Alternatively the microorganismmay remain free-living in the soil. In either case, theassociation may prove beneficial to the plant or it maybe pathogenic and cause injury.

3.9.1 BACTERIA OTHER THANNITROGEN FIXERS CONTRIBUTETO NUTRIENT UPTAKE BY ROOTS

Plant roots generally support large populations of bacte-ria, principally because of the large supply of energy-richnutrients provided by the growing root system. Theimmediate environment of the roots is so favorableto bacterial growth that the bacterial population inthe rhizosphere may exceed that in the surroundingbulk soil by as much as 50 percent. Nutrients pro-vided by the roots are comprised largely of aminoacids and soluble amides, reducing sugars, and otherlow-molecular-weight compounds. These compoundsmay either leak from the cells (a nonmetabolic pro-cess) or be actively secreted into the apoplastic spacefrom whence they readily diffuse into the surroundingrhizosphere.

Dominant among the root secretions are themucilages: polysaccharides secreted by Golgi vesiclesin cells near the growing tip. Secretion of mucilageappears to be restricted to cells such as root cap cells,young epidermal cells, and root hairs where sec-

ondary walls have yet to form. Secretion of mucilage inthe more basal regions of the root appears to be res-tricted by development of secondary walls. Themucilage is rapidly invaded by soil bacteria thatcontribute their own metabolic products, includingmucopolysaccharides of the bacterial capsule. Inaddition, mucilage also attracts colloidal mineral andorganic matter from the soil. The resulting mixture ofroot secretions, living and dead bacteria, and colloidalsoil particles is commonly referred to as mucigel.

There is no doubt that bacteria are intimatelyinvolved in the nitrogen nutrition of plants. Bothinvasive and free-living nitrogen-fixing bacteria, knownsince the late nineteenth century, are the primarysource of nitrogen for plants. In addition, other soilbacteria convert ammonium nitrogen to nitrate. But towhat extent do the bacteria influence other aspects ofplant nutrition? In the previous chapter we pointed outthat phosphorous is sparingly soluble in most soils and,in natural ecosystems, is often the limiting nutrient.There is some evidence that soil bacteria can assistin making phosphorous available by solubilizing thewater-insoluble forms. It is considered unlikely, how-ever, that this represents a major source of phosphorousfor plants, especially in light of the extensive fungalassociations described in the next section. Bacteriacan, however, enhance nutrient uptake other than bysimply making nutrients more available. One way is toinfluence the growth and morphology of roots. One ofthe more striking examples is the formation of proteoidroots. This is a phenomenon of localized, intense lateralroot production observed originally in the Proteaceae,a family of tropical trees and shrubs. (The Proteaceaeincludes the genus Macadamea, the source of thepopular macadamia nut.) Proteoid roots have now beenfound in several other families. Their induction hasbeen traced to localized aggregations of bacteria in themucigel. The larger number of lateral roots allows amore intensive mining of the soils for poorly mobile

3.9 Root-Microbe Interactions 57

nutrients, such as phosphorous. In addition, proteoidroots are generally found near the soil surface wherethey can take advantage of nutrients leached out ofthe litter. The mechanism for proteoid root inductionhas not been determined, but could be related to theproduction of a plant hormone (indoleacetic acid) by thebacteria.

3.9.2 MYCORRHIZAE ARE FUNGI THATINCREASE THE VOLUME OF THENUTRIENT DEPLETION ZONEAROUND ROOTS

Perhaps the most widespread—and from the nutritionalperspective, more significant—associations betweenplants and microorganisms are those formed betweenroots and a wide variety of soil fungi. A root infectedwith a fungus is called a mycorrhiza (literally, fungusroot). Mycorrhizae are a form of mutualism, anassociation in which both partners derive benefit.The significance of mycorrhizae is reflected in theobservation that more than 80 percent of plantsstudied, including virtually all plant species of economicimportance, form mycorrhizal associations. Two majorforms of mycorrhizae are known: ectotrophic andendotrophic. The ectotrophic form, also known asectomycorrhizae, is restricted to a few families consist-ing largely of temperate trees and shrubs, such as pines(Pinaceae) and beech (Fagaceae). Ectomycorrhizae aretypically short, highly branched, and ensheathed by atightly interwoven mantle of fungal hyphae. The fungusalso penetrates the intercellular or apoplastic space ofthe root cortex, forming an intercellular network calleda Hartig net. Endotrophic mycorrhizae, or endomy-corrhizae, are found in some species of virtually everyangiosperm family and most gymnosperms as well(except the Pinaceae). Unlike the ectomycorrhizae, thehyphae of endomycorrhizae develop extensively withincortical cells of the host roots.

The most common type of endomycorrhiza,found in the majority of the world’s vegetation, isthe vesicular-arbuscular mycorrhiza (VAM). Thehyphae of VAM grow between and into root corticalcells, where they form highly branched ‘‘treelike’’structures called arbuscules (meaning dwarf tree). Eachbranch of the arbuscule is surrounded by the plasmamembrane of the host cell. Thus, while the hyphae dopenetrate the host cell wall, they do not actually invadethe protoplast. The arbuscule serves to increase contactsurface area between the hypha and the cell by two tothree times. At the same time, it apparently influencesthe host cell, which may increase its cytoplasmicvolume by as much as 20 to 25 percent. Less frequently,VAMs form large ellipsoid vesicles either betweenor within the host cells. The presence of arbusculesand vesicles provides a large surface for the exchange

of nutrients between the host plant and the invadingfungus. Although VAMs do not form a well-definedsheath around the root, the hyphae, like those of theectomycorrhizae, do effectively extend the rhizosphereby growing outward into the surrounding soil.

Mycorrhizae were originally discovered by thenineteenth-century German botanist A. B. Frank, whoconcluded, on the basis of experiments conductedwith beech seedlings, that mycorrhizal inoculationstimulated seedling growth. Although not universallyaccepted in the beginning, these results have beenamply confirmed by more modern studies. Numerousstudies with pine and other tree seedlings in the UnitedStates, Australia, and the former Soviet Union havedemonstrated 30 to 150 percent increases in dry weightof tree seedlings infected with mycorrhizae when com-pared with noninfected controls. Similar results havebeen obtained in studies with agricultural plants such asmaize (Figure 3.13). In one experiment, for example, thedry weight of VAM-infected Lavendula plants increased8.5 times over noninfected controls. The primarycause of mycorrhizal-enhanced growth appears to beenhanced uptake of nutrients, especially phosphorous.In a classic experiment, Hatch demonstrated in 1937that infected pine seedlings absorbed two to three timesmore nitrogen, potassium, and phosphorous. Coupled

Nutrientdepletion zonewithoutmycorrhizae

Nutrientdepletion zone

with mycorrhizae

FIGURE 3.13 Infection of roots with mycorrhizal fungiextends the nutrient depletion zone for a plant. Thenutrient depletion zone is the zone from which nutrientsare drawn by the root system.

58 Chapter 3 / Roots, Soils, and Nutrient Uptake

with enhanced nutrient uptake is the observation thatmycorrhiza-induced growth responses are more pro-nounced in nutrient-deficient soils. VAM infection, forexample, can be effectively eliminated by supplying theplant with readily available phosphorous. With a surplusof phosphorous fertilizer, uninoculated plants will growas well as those inoculated with mycorrhizal fungi.

The beneficial role of mycorrhizae, particularly withrespect to the uptake of phosphorous, appears to berelated to the nutrient depletion zone that surroundsthe root. This zone defines the limits of the soil fromwhich the root is able to readily extract nutrient ele-ments. Additional nutrients can be made available onlyby extension of the root into new regions of the soilor by diffusion of nutrients from the bulk soil into thedepletion zone. The extent of the depletion zone variesfrom one nutrient element to another, depending onthe solubility and mobility of the element in the soilsolution. The depletion zone for nitrogen, for example,extends some distance from the root because nitrate isreadily soluble and highly mobile. Phosphorous, on theother hand, is less soluble and relatively immobile in soilsand, consequently, the depletion zone for phosphorousis correspondingly smaller. Mycorrhizal fungi assist inthe uptake of phosphorous by extending their myceliabeyond the phosphorous depletion zone (Figure 3.13).Apparently, mycorrhizal plants find it advantageous toexpend their carbon resources supporting mycorrhizalgrowth as opposed to more extensive growth of the rootsystem itself.

As we continue to learn about mycorrhizae, theirnutritional role becomes increasingly evident. Manymycorrhizae are host species–specific. Attempts toestablish a plant species in a new environment maybe unsuccessful if the appropriate mycorrhizal fungusis not present. Inoculation of fields with mycorrhizalfungi is now an additional factor taken into account bythe forest and agricultural industries when attemptingto resolve problems of soil infertility.

SUMMARY

The uptake of nutrient salts by plants involves acomplex interaction between plant roots and thesoil. The colloidal component of soil, consisting ofclay particles and humus, presents a highly specificsurface area carrying numerous, primarily negative,charges. Ions adsorbed to the charged colloidal surfacesrepresent the principal reservoir of nutrients for theplant. As ions are taken up from the dilute soil solutionby the roots, they are replaced by exchangeable ionsfrom the colloidal reservoir.

For nutrients to be taken up by a plant, they mustbe transported across the cell membrane into the root

cell—thus making nutrient uptake fundamentally a cel-lular problem. Solutes may cross a membrane by simplediffusion, facilitated diffusion, or active transport. Facil-itated diffusion and active transport are mediated bychannel and carrier proteins—proteins that span thelipid bilayer. Only active transport achieves accumula-tion of ions against an electrochemical gradient. Activetransport requires a source of metabolic energy, nor-mally in the form of ATP. The free movement of wateracross membranes has long been an enigma, although itnow appears to be a special case of facilitated diffusionthrough water-selective channels called aquaporins.The uptake of nutrients by most plants is enhanced byassociation of the roots with soil microorganisms, espe-cially fungi. Fungal-root associations (mycorrhizae)benefit the plant by significantly increasing the volumeof soil accessible to the roots.

CHAPTER REVIEW

1. Distinguish between simple diffusion, facilitateddiffusion, and active transport. Which of thesethree mechanisms would most probably accountfor:

(a) entry of a small lipid-soluble solute;(b) extrusion of sodium ions leaked into a cell;(c) rapid entry of a neutral hydrophilic sugar;(d) accumulation of potassium ions?

2. The Casparian band was encountered earlier withregard to root pressure and again in this chapterwith regard to ion uptake. What is the Casparianband and how does it produce these effects?

3. Trace the pathway taken by a potassium ion fromthe point where it enters the root to a leaf epider-mal cell.

4. What are channel proteins and what role do theyplay in nutrient uptake?

5. What is the Nernst equation and what does it tellus about ion transport?

6. How does the concept of molecular crosstalk pertain to plant cellular ion transport?

7. Describe the concept of apparent free space. Whatrole does apparent free space play in the uptake ofnutrient ions?

8. Why is an accumulation ratio greater than 1.0 notnecessarily an indication that active transport isinvolved?

9. How do mycorrhizae assist a plant in the uptake ofnutrient elements?

10. Describe the colloidal properties of soil. Howdo the properties of colloids help to ensure theavailability of nutrient elements in the soil?

Further Reading 59

FURTHER READING

Britto, D. T, H. J. Kronzucker. 2006. Futile cycling at theplasmamembrane: A hallmark of low affinity nutrienttransport. Trends in Plant Science 11: 529–534.

Buchanan, B. B., W. Gruissem, R. L. Jones. 2000.Biochemistry and Molecular Biology of Plants. Rockville,MD: American Society of Plant Physiologists.

Cooper, J. E., J. R. Rao. 2006. Molecular Approaches to Soil,Rhizosphere and Plant Micro-organism Analysis. Cam-bridge: CABI Publishing.

Demidchik, V., R. J. Davenport, M. Tester. 2002. Nonse-lective cation channels in plants. Annual Review of PlantBiology 53: 67–107.

Epstein, E., A. J. Bloom. 2005. Mineral Nutrition of Plants:Principles and Perspectives. Sunderland: Sinauer AssociatesInc.

Evert, R. F. 2006. Esau’s Plant Anatomy. Hoboken, NJ: JohnWiley & Sons, Inc.

Nobel, P. S. 2005. Physicochemical and Environmental PlantPhysiology. Burlington, MA: Elsevier Science & Technol-ogy.

Peterson, R. L. 2004. Mycorrhizas: Anatomy and Cell Biology.Ottawa: NRC Press.

Schachtman, D. P., R. Shin. 2007. Nutrient sensing: NPKS.Annual Review of Plant Biology 58: 47–69.

Waisel, Y., A. Eshel, U. Kafkafi. 1996. Plant Roots: The HiddenHalf . 2nd ed. New York: M. Dekker.

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4Plants and Inorganic Nutrients

Unlike heterotrophic organisms, which depend fortheir existence on energy-rich organic moleculespreviously synthesized by other organisms, plantsmust survive in an entirely inorganic environment. Asautotrophic organisms, plants must take in carbondioxide from the atmosphere and water and mineralnutrients from the soil, and from these simple, inorganiccomponents, make all of the complex molecules ofa living organism. Since plants stand at the bottomof the food chain, mineral nutrients assimilated byplants eventually find their way into the matter thatmakes up all animals, including humans. All organismsmust continually draw material substance from theirenvironment in order to maintain their metabolism,growth, and development. The means for making thesematerials available to the organism is the subject ofplant nutrition.

Plant nutrition is traditionally treated as two sep-arate topics: organic nutrition and inorganic nutrition.Organic nutrition focuses on the production of carboncompounds, specifically the incorporation of carbon,hydrogen, and oxygen via photosynthesis, while inor-ganic nutrition is concerned primarily with the acquisi-tion of mineral elements from the soil. Photosynthesisand the acquisition of mineral ions from the soil are so

interdependent, however, that this distinction betweenorganic and inorganic nutrition is more a matter of con-venience than real. Nevertheless, because the acquisitionand assimilation of carbon are addressed in Chapters 7and 8, this chapter will focus on the acquisition of min-eral elements and the role of those elements in plantmetabolism.

This chapter will examine the nutritional require-ments of plants that are satisfied by mineral elements.This will include

• methods employed in the study of mineral nutrition,• the concept of essential and beneficial elements

and the distinction between macronutrients andmicronutrients,

• a general discussion of the metabolic roles of the 14essential mineral elements, the concept of criticaland deficient concentration, and symptoms asso-ciated with deficiencies of the mineral elements,and

• a brief discussion of micronutrient toxicity.

The soil as a nutrient reservoir and mechanisms formineral uptake by roots was covered in Chapter 3.

61

62 Chapter 4 / Plants and Inorganic Nutrients

4.1 METHODS AND NUTRIENTSOLUTIONS

4.1.1 INTEREST IN PLANT NUTRITIONIS ROOTED IN THE STUDY OFAGRICULTURE AND CROPPRODUCTIVITY

Much of the groundwork for modern nutritional studieswas laid in Europe in the early to mid-nineteenth cen-tury, in response to a combination of political and socialfactors. The Napoleonic wars had devastated Europeand the industrial revolution was gaining momentum.Rising populations and massive migration to the citiescreated demands that could no longer be met by thetraditional agricultural economy, one that relied heav-ily on the use of organic manures. Greater efficiencyin agriculture was required and this was not possi-ble without a more thorough understanding of plantnutrition.

One of the first to make significant progress inthe study of plant nutrition was N. T. de Saussure(1767–1845), who studied both photosynthesis and theabsorption of nutrient elements with the same care-ful, quantitative methods. De Saussure conducted someof the first elemental analyses of plant material andintroduced the concept that some, but not necessarilyall, of the elements found might be indispensable, oressential, to plant growth. De Saussure’s ideas concern-ing the importance of elements derived from the soilgenerated considerable debate at the time, but receivedsupport from the work of C. S. Sprengel (1787–1859),working in Germany, and Jean-Baptiste Boussingault inFrance. Sprengel introduced the idea that soils mightbe unproductive if deficient in but one single elementnecessary for plant growth, and Boussingault stressedquantitative relationships between the effects of fertil-izer and nutrient uptake on crop yields. Boussingaultis also credited with providing the first evidence thatlegumes had the unique capacity to assimilate atmo-spheric nitrogen, a finding that was later confirmed bythe discovery of the nitrogen-fixing role of bacteria inroot nodules.

By the middle of the nineteenth century, manypieces of the nutritional puzzle were beginning to fallinto place. In 1860, Julius Sachs, a prominent Germanbotanist, demonstrated for the first time that plants couldbe grown to maturity in defined nutrient solutions in thecomplete absence of soil. J. B. Lawes and J. H. Gilbert,working at Rothamsted in England, had successfullyconverted insoluble rock phosphate to soluble phos-phate (called superphosphate), and by the end of thecentury the agricultural use of NPK (nitrogen, phospho-rous, and potassium) fertilizers was well established inEurope.

4.1.2 THE USE OF HYDROPONICCULTURE HELPED TO DEFINETHE MINERAL REQUIREMENTSOF PLANTS

In the mid-nineteenth century, J. Sachs was interestedin determining the minimal nutrient requirements ofplants. Recognizing that it would be difficult to pursuesuch studies in a medium as complex as soil, Sachsdevised an experimental system such that the rootsgrew not in soil but in an aqueous solution of mineralsalts. With this simplified system, Sachs was able todemonstrate the growth of plants to maturity on a rela-tively simple nutrient solution containing six inorganicsalts (Table 4.1). Variations on Sachs’s system, knownas solution or hydroponic culture (growing plants ina defined nutrient solution), have remained to this daythe principal experimental system for study of plantnutrient requirements. Hydroponic culture is also nowused extensively in North America for the year-roundcommercial production of vegetables such as lettuce,tomato, sweet peppers, and seedless cucumber.

The nutrient solution devised by Sachs contributeda total of nine mineral nutrients (K, N, P, Ca, S, Na, Cl,Fe, Mg). Carbon, hydrogen, and oxygen were excludedfrom this total because they were provided in the formof carbon dioxide and water and were not consideredmineral elements. It was at least another half centurybefore the need for additional mineral nutrients wasdemonstrated. There was no magic to the success ofSachs’s experiments. Many of the mineral nutrients usedby plants are required in very low amounts and Sachsunknowingly provided these nutrients as impurities inthe salts and water he used to make up his nutri-ent solution. Analytical techniques have now improvedto the point where it is possible to detect mineralcontents several orders of magnitude lower than waspossible in Sachs’s time. Most mineral elements arenow measured by either atomic absorption spectrome-try or atomic emission spectrometry. These techniquesinvolve vaporization of the elements at temperatures

TABLE 4.1 The composition of Sachs’s nutrientsolution (1860) used for solution culture of plants.

ApproximateConcentration

Salt Formula (mM)

Potassium nitrate KNO3 9.9Calcium phosphate Ca3(PO4)2 1.6Magnesium sulfate MgSO4z7H2O 2.0Calcium sulfate CaSO4 3.7Sodium chloride NaCl 4.3Iron sulfate FeSO4 trace

4.1 Methods and Nutrient Solutions 63

in excess of several thousand degrees. In the vaporousstate, the element will either absorb or emit light atvery narrow wavelength bands. The wavelength of lightabsorbed or emitted is characteristic of a particular ele-ment and the quantity of absorbed or emitted energyis proportional to the concentration of the elementin the sample. In this way, concentrations as low as10−8 g ml−1 for some elements can be measured insamples of plant tissue, soil, or nutrient solutions withina few minutes.

Aside from the commercial applications ofhydroponic plant culture, a great deal of plantphysiology and other botanical research is conductedwith plants grown under controlled environments. Thismay include relatively simple greenhouses or complexgrowth rooms in which temperature and lighting arecarefully regulated. Plant nutrient supply must also beregulated, and over the years a large number of nutrientsolutions have been formulated for this purpose. Mostmodern formulations are based on a solution originallydeveloped by D. R. Hoagland, a pioneer in the studyof plant mineral nutrition. Individual investigatorsmay introduce minor modifications to the composi-tion of the nutrient solution in order to accommo-date specific needs. Such formulations are commonlyreferred to as modified Hoagland’s solutions(Tables 4.2, 4.3).

The concentration of minerals in most nutrientsolutions is many times greater than that normally foundin soils. An excess is necessary in order to maintain acontinual supply of nutrients as they are taken up by theroots. The nutrient concentration of the soil solution,on the other hand, is relatively low but is continuallyreplenished by nutrients adsorbed on the soil particles(Chapter 3).

TABLE 4.2 The composition of a typicalone-half strength ‘‘modified’’ Hoagland’s nutrientsolution, showing the nutrient salts used and theirapproximate millimolar (mM) concentrations.

Concentration(mM)

Calcium nitrate Ca(NO)3 2.5Potassium phosphate KH2PO4 0.5Potassium nitrate KNO3 2.5Magnesium sulfate MgSO4 1.0Zinc sulfate ZnSO4 0.00039Manganous sulfate MnSO4 0.0046Copper sulfate CuSO4 0.00016Boric acid H3BO3 0.0234Molybdic acid MoO3 0.000051Iron sequestrene Fe 0.179

TABLE 4.3 The quantity of each nutrientelement in modified Hoagland’s nutrient solution.

Element Mg/L

Calcium 103Nitrogen 105Potassium 118Sulfur 33Magnesium 25Phosphorous 15Iron 10Boron 0.25Manganese 0.25Zinc 0.025Copper 0.01Molybdenum 0.0052

4.1.3 MODERN TECHNIQUESOVERCOME INHERENTDISADVANTAGES OF SIMPLESOLUTION CULTURE

In the simplest form of solution culture, a seedling issupported in the lid of a container, with its roots freeto grow in the nutrient solution (Figure 4.1). Note thatthe solution must be aerated in order to obtain optimalroot growth and nutrient uptake. A solution that is notaerated becomes depleted of oxygen, a condition knownas anoxia. Anoxia inhibits the respiration of root cellsand, because nutrient uptake requires energy, reducesnutrient uptake. The container in which the plantsare grown is usually painted black or wrapped with anopaque material in order to keep out light. The purpose

Funnel for addingwater and nutrients Aerating

tubeDacronwad

FIGURE 4.1 Diagram of a typical setup for nutrient solu-tion culture. (From Epstein, E. 1972. Mineral Nutritionof Plants: Principles and Perspectives. New York: Wiley.Reprinted by permission)

64 Chapter 4 / Plants and Inorganic Nutrients

of excluding light is to reduce the growth of algae thatwould compete with the plants for nutrients or possiblyproduce toxic byproducts.

There are some disadvantages in the use of a simplesolution culture to study the nutrient requirements ofplants. The major problems are a selective depletion ofions and associated changes in the pH of the solutionthat occur as the roots continue to absorb nutrients.Plants maintained in pure solution culture will con-tinue to grow vigorously only if the nutrient solutionis replenished on a regular basis. In order to avoidsuch problems, some investigators grow the plants in anonnutritive medium such as acid-washed quartz sand,perlite, or vermiculite.1 Plants can then be watered bydaily application of fresh nutrient solution from the topof the medium (a technique called slop culture) orby slowly dripping onto the culture from a reservoir(drip culture). Alternatively, the nutrient culture canbe subirrigated. In this case, the nutrient solution is

1Vermiculite is a silicate mineral of the mica family. Itexpands on heating to produce a lightweight product that hashigh water retention and is commonly used as a mulch in seedbeds. Perlite is a coarsely ground glassy volcanic rock. Bothvermiculite and perlite are effectively inert substances thatprovide no plant nutrients.

alternately pumped into the culture from below andthen allowed to drain out. This fill-and-empty pro-cess is repeated on a regular basis and serves bothto replenish the nutrient solution and to aerate theroots. Most commercial hydroponic operations nowutilize some variation of the nutrient film techniquein which the roots are continuously bathed with athin film of recirculating nutrient solution (Figure 4.2).The advantage of the nutrient film technique is that itnot only provides for good aeration of the roots andnutrient uptake; it also allows the pH and nutrient con-tent of the solution to be continuously monitored andadjusted.

These methods overcome some of the problemsinherent in pure solution culture, but may not besuitable for many laboratory experiments. This isbecause no medium is truly nonnutritive. Any medium,even the glass, plastic, or ceramic containers usedin solution culture, may provide some nutrients atvery low levels. For example, soft (sodium silicate)glass provides sodium, hard (borosilicate) glassesprovide boron, and plastics might provide chloride orfluoride, and so forth. Water used to prepare nutrientsolutions must be carefully distilled, avoiding, whereverpossible, metallic components in the distillationapparatus.

FIGURE 4.2 The nutrient film tech-nique for hydroponic plant produc-tion. Plants are grown in a tube ortrough placed on a slight incline. Apump (P) circulates nutrient solu-tion from a reservoir to the elevatedend of the tube. The solution thenflows down the tube by gravity,returning to the reservoir. Inset:the roots grow along the bottom ofthe tube, bathed continuously in athin film of aerated nutrient solu-tion. Arrows indicate the directionof nutrient flow.

P

4.2 The Essential Nutrient Elements 65

4.2 THE ESSENTIAL NUTRIENTELEMENTS

4.2.1 SEVENTEEN ELEMENTS AREDEEMED TO BE ESSENTIAL FORPLANT GROWTH ANDDEVELOPMENT

Most plants require a relatively small number of nutri-ent elements in order to successfully complete theirlife cycle. Those that are required are deemed to beessential nutrient elements. Essentiality is based pri-marily on two criteria formulated by E. Epstein in 1972.According to Epstein, an element is considered essentialif (1) in its absence the plant is unable to complete a normallife cycle, or (2) that element is part of some essential plantconstituent or metabolite. By the first criterion, if a plantis unable to produce viable seed when deprived of thatelement, then that element is deemed essential. By thesecond criterion, an element such as magnesium wouldbe considered essential because it is a constituent ofthe chlorophyll molecule and chlorophyll is essential forphotosynthesis. Similarly, chlorine is essential becauseit is a necessary factor in the photosynthetic oxidationof water. Most elements satisfy both criteria, althougheither one alone is usually considered sufficient.

Although the criteria for essentiality are quite clear,it is not always easy to demonstrate that an element is oris not essential. D. Arnon and P. Stout had earlier sug-gested a third criterion: they suggested that an essentialelement must act directly in the metabolism of the plantand not simply to correct an unfavorable microbial orchemical condition in the nutrient medium. The use ofsolution cultures, from which the element in questionhas been omitted, has largely circumvented the needto apply this third criterion. On the other hand, someplants may form viable seeds even though a particularelement has been excluded from the nutrient solutionand other symptoms of deficiency are evident. In suchcases there may be present in the seed, or contaminatingthe nutrient solution, a quantity of the element sufficientto moderate the deficiency and allow seed formation.It is assumed that in the complete absence of a nutri-ent, the deficiency symptoms would be severe enoughto kill the plant before viable seed could be formed.If required, this can be confirmed by careful purifi-cation of nutrient salts and exclusion of atmosphericcontaminants. Where a sufficient quantity of the ele-ment may be carried within the seed, essentiality canbe confirmed by growing several successive generationsfrom seed that was itself produced in the absence of thatelement. This is usually sufficient to reduce the con-centration of that element in the seed to the deficientrange.

It is generally agreed, based on these criteria, thatonly 17 elements are essential for the growth of allhigher plants (Table 4.4).

4.2.2 THE ESSENTIAL NUTRIENTSARE GENERALLY CLASSED ASEITHER MACRONUTRIENTSOR MICRONUTRIENTS

The essential elements are traditionally segregated intotwo categories: (1) the so-called macronutrients and (2)the trace elements or micronutrients. The distinctionbetween macro- and micronutrients simply reflects therelative concentrations found in tissue or required innutrient solutions (Table 4.4) and does not infer impor-tance relative to the nutritional needs of the plant. Thefirst nine elements in Table 4.4 are called macronutrientsbecause they are required in large amounts (in excessof 10 mmole kg−1 of dry weight). The macronutrientsare largely, but not exclusively, involved in the structureof molecules, which to some extent accounts for theneed for large quantities. The remaining eight essentialelements are considered micronutrients. Micronutri-ents are required in relatively small quantities (lessthan 10 mmole kg−1 of dry weight) and serve catalyticand regulatory roles such as enzyme activators. Somemacronutrients, calcium and magnesium for example,serve as regulators in addition to their structural role.

4.2.3 DETERMINING ESSENTIALITYOF MICRONUTRIENTS PRESENTSSPECIAL PROBLEMS

The essentiality of micronutrients is particularly difficultto establish because they are required in such smallquantities. Most micronutrient requirements are fullysatisfied by concentrations in the range of 0.1 to 1.0 μgL−1 —amounts that are readily obtained from impuritiesin water or macronutrient salts, the containers in whichthe plants are grown, and contamination by atmosphericdust. A micronutrient may be required at concentrationsbelow detectable limits, so it is far easier to establish thata micronutrient is essential than that it is not.

As micronutrients go, iron is usually supplied atrelatively high concentrations. This is necessary becauseavailability of iron is very sensitive to pH and other soilconditions. At a pH above 7, iron tends to form insol-uble iron hydroxides and calcium complexes. In acidicsolution, iron reacts with aluminum to form insolublecomplexes. In both cases, iron readily precipitates outof solution and, consequently, is frequently deficient innatural situations. For these reasons, the need for ironas an essential plant nutrient was established early in thestudy of plant nutrition. The need for other micronutri-ents, however, was not recognized until salts of sufficientpurity became available in the early part of the twentiethcentury. In some cases, what had been for some timerecognized as a plant disease turned out to be a nutri-ent deficiency. In 1922, for example, J. S. McHarguedemonstrated that the disorder known as gray speck of oatswas actually caused by a manganese deficiency, and the

66 Chapter 4 / Plants and Inorganic Nutrients

TABLE 4.4 The essential nutrient elements of higher plants and their concentrationsconsidered adequate for normal growth.

Chemical Available Concentration in DryElement Symbol Form Matter (mmol/kg)

MacronutrientsHydrogen H H2O 60,000Carbon C CO2 40,000Oxygen O O2,CO2 30,000Nitrogen N NO−

3 , NH+4 1,000

Potassium K K+ 250Calcium Ca Ca2+ 125Magnesium Mg Mg2+ 80Phosphorous P HPO−

4 , HPO2−4 60

Sulfur S SO2−4 30

MicronutrientsChlorine Cl Cl− 3.0Boron B BO3−

3 2.0Iron Fe Fe2+, Fe3+ 2.0Manganese Mn Mn2+ 1.0Zinc Zn Zn2+ 0.3Copper Cu Cu2+ 0.1Nickel Ni Ni2+ 0.05Molybdenum Mo Mo2−

4 0.001

following year Katherine Warington showed that boronwas required for several legume species. By 1939, theneed for zinc, copper, and molybdenum had also beenclearly established. In each case, the nutrient deficiencywas found to cause a well-known disorder previouslythought to be a disease. Chlorine was not added to thelist until 1954, although its essential nature was sug-gested nearly one hundred years earlier. The need forchlorine became evident in the course of experimentsto determine whether cobalt was required for tomato(Lycopersicum esculentum). T. C. Broyer and his cowork-ers had purified their nutrient salts by methods thatremoved not only cobalt but halides (including chlo-rine) as well. Plants grown in solutions prepared fromthese purified salts developed browning and necrosisof the leaves. The symptoms could be avoided by sup-plementing the nutrient solution with cobalt chloride.Subsequent investigation, however, established that itwas the deficiency of chloride rather than cobalt thatgave rise to the symptoms.

There is now mounting evidence that nickel shouldbe added to the list of essential elements. Nickel isan essential component of urease, an enzyme widelydistributed in plants, microorganisms, and some marineinvertebrates. Urease catalyzes the hydrolysis of ureainto NH3 and CO2 and is thought to play an importantrole in mobilization of nitrogenous compound in plants.In 1987, P. H. Brown and his colleagues showed that

nickel depletion led to the formation of nonviable seedin barley (Hordeum vulgare). The addition of nickelwould bring to 17 the total number of nutrient elementsessential for higher plants.

4.3 BENEFICIAL ELEMENTS

In addition to the 17 essential elements listed inTable 4.4, some plants appear to have additionalrequirements. However, because these have not beenshown to be requirements of higher plants generally,they are excluded from the list of essential elements.They are referred to instead as beneficial elements.If these elements are essential to all plants, they arerequired by most at concentrations well below what canbe reliably detected by present analytical techniques.The definition of beneficial currently applies primarilyto sodium, silicon, selenium, and cobalt. With time,and as experimental methods improve, one or more ofthese beneficial elements may be added to the list ofessential elements.

4.3.1 SODIUM IS AN ESSENTIALMICRONUTRIENT FOR C4 PLANTS

A sodium requirement was first demonstrated for thebladder salt-bush (Atriplex vesicaria), a perennial pasturespecies of arid inland areas of Australia. By carefully

4.4 Nutrient Functions and Deficiency Symptoms 67

purifying the water, recrystallizing the nutrient salts,and using sodium-free vessels, P. F. Brownell and C. J.Wood were able to reduce the sodium content of thefinal culture medium to less than 1.6 μg L−1. Plantsgrown in the depleted solution showed reduced growth,chlorosis (yellowing due to loss of chlorophyll), andnecrosis (dead tissue) of the leaves. Based on a surveyof 32 species of plants, it was concluded that sodium isgenerally essential as a micronutrient for plants utilizingspecifically the C4 photosynthetic pathway, but not formost C3 plants (see Chapter 8 and 15).

4.3.2 SILICON MAY BE BENEFICIALFOR A VARIETY OF SPECIES

Given the high content of silicon dioxide in normalsoils, it should not be surprising that many plants takeup appreciable quantities of silicon. Silicon may com-prise 1 to 2 percent of the dry matter of maize (Zeamays) and other grasses and as much as 16 percent of thescouring rush (Equisetum arvense), yet experiments havegenerally failed to demonstrate that silicon is essentialfor most other plants.2 The ubiquitous presence of sili-con in glass, nutrient salts, and atmospheric dust makesit especially difficult to exclude silicon from nutrientexperiments. However, there are numerous reports ofbeneficial effects of silicon in a variety of species. Siliconseems to be particularly beneficial to grasses, where itaccumulates in the cell walls, especially of epidermalcells, and may play a role in fending off fungal infectionsor preventing lodging, a condition in which stems arebent over by heavy winds or rain.

4.3.3 COBALT IS REQUIRED BYNITROGEN-FIXING BACTERIA

Cobalt is essential for the growth of legumes, which arehosts to symbiotic nitrogen-fixing bacteria (Chapter 11).In this case, the requirement can be traced to the needs ofthe nitrogen-fixing bacterium rather than the host plant.A similar cobalt requirement has been demonstrated forthe free-living nitrogen-fixing bacteria, including thecyanobacteria. In addition, when legumes are providedwith fixed nitrogen such as nitrate, a cobalt requirementcannot be demonstrated.

4.3.4 SOME PLANTS TOLERATE HIGHCONCENTRATIONS OF SELENIUM

Selenium salts tend to accumulate in poorly drained,arid regions of the western plains of North America.Although selenium is generally toxic to most plants, cer-

2Grazing animals appear to have adapted to the high siliconcontent of grasses. The teeth of grazing animals (such as cowsand horses) grow continuously, compensating for the wearcaused largely by silicon. On the other hand, the teeth ofbrowsing animals such as deer, whose diets contain littlegrass, do not continue to grow.

tain members of the legume genus Astragalus (milk-vetchor poison-vetch) are known to tolerate high concentra-tions of selenium (up to 0.5 percent dry weight) and arefound only on soils containing relatively high concen-trations of selenium. Such concentrations of seleniumwould be toxic to most other plants. At one time it wasthought that selenium might be essential to these ‘‘accu-mulator species,’’ but there is no definitive supportingevidence.

Selenium accumulators are of considerable impor-tance to ranchers, however, as they are among a diversegroup of plants known as ‘‘loco weeds.’’ The high sele-nium content in these plants causes a sickness known asalkali poisoning or ‘‘blind-staggers’’ in grazing animals.

4.4 NUTRIENT FUNCTIONS ANDDEFICIENCY SYMPTOMS

The essential elements are essential because they havespecific metabolic functions in plants. When they areabsent, plants will exhibit characteristic deficiency symp-toms that, in most cases, are related to one or more ofthose functions.

Some students of plant mineral nutrition prefer toclassify the macro- and micronutrients along functionallines. For example, elements such as carbon, hydrogen,and oxygen have a predominantly structural role—theyare the stuff of which molecules are made—while othersappear to be predominantly involved in regulatory roles,such as maintaining ion balance and activating enzymes.Other investigators have proposed more complicatedschemes with up to four categories of biochemicalfunction. Unfortunately, any attempt to categorize thenutrient elements in this way runs into difficulty becausethe same element often fills both structural and non-structural roles. Magnesium, for example, is an essentialcomponent of the chlorophyll molecule but also servesas a cofactor for many enzymes, including ATPases andothers involved in critical energy-transfer reactions. Cal-cium is an important constituent of cell walls where itsrole is largely structural, but there are also Ca-ATPasesand calcium is implicated as a second messenger in hor-mone responses and photomorphogenesis (Chapters 16,17, 22). Regardless of how they are classified, it is clearthat these elements are essential because they satisfy spe-cific metabolic requirements of the plant. When thoserequirements are not met or are only partially met,the plant will exhibit characteristic deficiency symptomsthat, if severe enough, result in death.

4.4.1 A PLANT’S REQUIREMENTFOR A PARTICULAR ELEMENTIS DEFINED IN TERMS OFCRITICAL CONCENTRATION

Typically, when the supply of an essential elementbecomes limiting, growth is reduced. The concentration

68 Chapter 4 / Plants and Inorganic NutrientsG

row

th

Criticalconcentration

Nutrient concentration

Adequate range

Deficientrange

FIGURE 4.3 Generalized plot of growth as a function ofnutrient concentration in tissue. The critical concentra-tion is that concentration giving a 10 percent reductionin growth. At concentrations less than critical, the nutri-ent is said to be deficient. Greater concentrations areconsidered adequate.

of that nutrient, measured in the tissue, just below thelevel that gives maximum growth, is defined as thecritical concentration. This concept is illustrated inFigure 4.3. At concentrations above the critical concen-tration, additional increments in nutrient content willhave no beneficial effect on growth and the nutrientcontent is said to be adequate. At concentrations belowthe critical concentration, the nutrient content becomesdeficient and growth falls off sharply. In other words,at tissue nutrient levels below the critical concentration,that nutrient becomes limiting to growth.

When nutrient levels exceed the critical concentra-tion, that nutrient is, with one qualification, no longerlimiting. The qualification is that at sufficiently hightissue levels, virtually all nutrients become toxic. Toxiclevels are seldom achieved with the macronutrients,but are common in the case of the micronutrients(Section 4.5). Normal concentrations of copper, forexample, are in the range of 4 to 15 μg g−1 of tissuedry weight (dwt). Deficiency occurs at concentrationsbelow 4 μg but most plants are severely damaged byconcentrations in excess of 20 μg g−1 dwt. Boron is toxicabove 75 μg g−1 dwt and zinc above 200 μg g−1 dwt.

Since each element has one or more specific struc-tural or functional roles in the plant, in the absenceof that element the plant will be expected to exhibitcertain morphological or biochemical symptoms of thatdeficiency. In some cases the deficiency symptoms willclearly reflect the functional role of that element. One

example is yellowing, or chlorosis, which is charac-teristic of several nutrient deficiencies. In the case ofmagnesium deficiency, for example, the plant turns yel-low because it is unable to synthesize the green pigmentchlorophyll. In other cases, the relationship betweendeficiency symptoms and the functional role of the ele-ment may not always be so straightforward. Moreover,deficiency symptoms for some elements are not alwaysconsistent between one plant and the next. Nonetheless,for each element there are certain generalizations thatcan be made with respect to deficiency symptoms.

Deficiency symptoms also depend in part on themobility of the element in the plant. Where elementsare mobilized within the plant and exported to youngdeveloping tissues, deficiency symptoms tend to appearfirst in older tissues. Other elements are relativelyimmobile—once located in a tissue they are not readilymobilized for use elsewhere. In this case, the deficiencysymptoms tend to appear first in the younger tissues.

In this section we will review the functional roles ofthe essential elements and, in general terms, morpho-logical and biochemical abnormalities that result fromtheir deficiencies. Carbon, hydrogen, and oxygen willbe excluded from this discussion as they are requiredfor the structural backbone of all organic molecules.A deficiency of carbon, consequently, leads quickly tostarvation of the plant, while a deficiency of water leadsto desiccation. Instead, our discussion will be limited todeficiencies of those essential elements taken in fromthe soil solution.

4.4.2 NITROGEN IS A CONSTITUENTOF MANY CRITICALMACROMOLECULES

Although the atmosphere is approximately 80 percentnitrogen, only certain prokaryote species—bacteriaand cyanobacteria—can utilize gaseous nitrogendirectly. The special problems of nitrogen availabilityand metabolism are addressed in Chapter 11. Tosummarize, most plants absorb nitrogen from the soilsolution primarily as inorganic nitrate ion (NO−

3 ) and,in a few cases, as ammonium (NH+

4 ) ion. Once in theplant, NO−

3 must be reduced to NH+4 before it can

be incorporated into amino acids, proteins, and othernitrogenous organic molecules. Nitrogen is most oftenlimiting in agricultural situations. Many plants, such asmaize (Zea mays), are known as ‘‘heavy feeders’’ andrequire heavy applications of nitrogen fertilizer. Themanufacture and distribution of nitrogen fertilizers foragriculture are, in both energy and financial terms, anextremely costly process.

Nitrogen is a constituent of many importantmolecules, including proteins, nucleic acids, certainhormones (e.g., indole-3-acetic acid; cytokinin), andchlorophyll. It should not be surprising, then, that the

4.4 Nutrient Functions and Deficiency Symptoms 69

most overt symptoms of nitrogen deficiency are a slow,stunted growth and a general chlorosis of the leaves.Nitrogen is very mobile in the plant. As the older leavesyellow and die, the nitrogen is mobilized, largely in theform of soluble amines and amides, and exported fromthe older leaves to the younger, more rapidly developingleaves. Thus the symptoms of nitrogen deficiency gen-erally appear first in the older leaves and do not occur inthe younger leaves until the deficiency becomes severe.At this point, the older leaves will turn completelyyellow or brown and fall off the plant. Conditions ofnitrogen stress will also lead to an accumulation ofanthocyanin pigments in many species, contributing apurplish color to the stems, petioles, and the undersideof leaves. The precise cause of anthocyanin accumula-tion in nitrogen-starved plants is not known. It may berelated to an overproduction of carbon structures that,in the absence of nitrogen, cannot be utilized to makeamino acids and other nitrogen-containing compounds.

Excess nitrogen normally stimulates abundantgrowth of the shoot system, favoring a high shoot/rootratio, and will often delay the onset of floweringin agricultural and horticultural crops. Similarly, adeficiency of nitrogen reduces shoot growth andstimulates early flowering.

4.4.3 PHOSPHOROUS IS PART OF THENUCLEIC ACID BACKBONE ANDHAS A CENTRAL FUNCTION ININTERMEDIARY METABOLISM

Phosphorous is available in the soil solution primarilyas forms of the polyprotic phosphoric acid (H3PO4).A polyprotic acid contains more than one proton, eachwith a different dissociation constant. Soil pH thusassumes a major role in the availability of phosphorous.At a soil pH less than 6.8, the predominant form ofphosphorous and the form most readily taken up byroots is the monovalent orthophosphate anion (H2PO−

4 ).Between pH 6.8 and pH 7.2, the predominant form isHPO2−

4 , which is less readily absorbed by the roots.In alkaline soils (pH greater than 7.2), the predomi-nant form is the trivalent PO3−

4 , which is essentially notavailable for uptake by plants. The actual concentra-tion of soluble phosphorous in most soils is relativelylow—on the order of 1 μM—because of several fac-tors. One factor is the propensity of phosphorous toform insoluble complexes. At neutral pH, for example,phosphorous tends to form insoluble complexes withaluminum and iron, while in basic soils calcium andmagnesium complexes will precipitate the phospho-rous. Because insoluble phosphates are only very slowlyreleased into the soil solution, phosphorous is alwayslimited in highly calcareous soils.

Substantial amounts of phosphorous may also bebound up in organic forms, which are not available for

uptake by plants. Organic phosphorous must first beconverted to an inorganic form by the action of soilmicroorganisms, or through the action of phosphataseenzymes released by the roots, before it is available foruptake. In addition, plants must compete with the soilmicroflora for the small amounts of phosphorous thatare available. For these sorts of reasons, phosphorous,rather than nitrogen, is most commonly the limitingelement in natural ecosystems. One of the more suc-cessful strategies developed by plants for increasing theuptake of phosphorous is the formation of intimate asso-ciations between roots and soil fungi, called mycorrhiza.Mycorrhizal associations were discussed in Chapter 3.

In the plant, phosphorous is found largely as phos-phate esters—including the sugar-phosphates, whichplay such an important role in photosynthesis and inter-mediary metabolism. Other important phosphate estersare the nucleotides that make up DNA and RNA as wellas the phospholipids present in membranes. Phospho-rous in the form of nucleotides such as ATP and ADP, aswell as inorganic phosphate (Pi), phosphorylated sugars,and phosphorylated organic acids also plays an integralrole in the energy metabolism of cells.

The most characteristic manifestation of phospho-rous deficiency is an intense green coloration of theleaves. In the extreme, the leaves may become mal-formed and exhibit necrotic spots. In some cases, theblue and purple anthocyanin pigments also accumulate,giving the leaves a dark greenish-purple color. Likenitrogen, phosphorous is readily mobilized and redis-tributed in the plant, leading to the rapid senescenceand death of the older leaves. The stems are usuallyshortened and slender and the yield of fruits and seedsis markedly reduced.

An excess of phosphorous has the opposite effect ofnitrogen in that it preferentially stimulates growth ofroots over shoots, thus reducing the shoot/root ratio.Fertilizers with a high phosphorous content, such asbone meal, are often applied when transplanting peren-nial plants in order to encourage establishment of astrong root system.

4.4.4 POTASSIUM ACTIVATESENZYMES AND FUNCTIONS INOSMOREGULATION

Potassium (K+) is the most abundant cellular cationand so is required in large amounts by most plants.In agricultural practice, potassium is usually providedas potash (potassium carbonate, K2CO3). Potassium isfrequently deficient in sandy soils because of its highsolubility and the ease with which K+ leaches out ofsandy soils. Potassium is an activator for a number ofenzymes, most notably those involved in photosynthesisand respiration. Starch and protein synthesis are alsoaffected by potassium deficiency. Potassium serves an

70 Chapter 4 / Plants and Inorganic Nutrients

important function in regulating the osmotic potentialof cells (Chapter 3). As an osmoregulator, potassiumis a principal factor in plant movements, such as theopening and closure of stomatal guard cells (Chapter 8)and the sleep movements, or daily changes in the ori-entation of leaves (Chapter 23). Because it is highlymobile, potassium also serves to balance the charge ofboth diffusible and nondiffusible anions.

Unlike other macronutrients, potassium is notstructurally bound in the plant, but like nitrogen andphosphorous is highly mobile. Deficiency symptomsfirst appear in older leaves, which characteristicallydevelop mottling or chlorosis, followed by necroticlesions (spots of dead tissue) at the leaf margins. Inmonocotyledonous plants, especially maize and othercereals, the necrotic lesions begin at the older tips of theleaves and gradually progress along the margins to theyounger cells near the leaf base. Stems are shortenedand weakened and susceptibility to root-rotting fungi isincreased. The result is that potassium-deficient plantsare easily lodged.

4.4.5 SULFUR IS AN IMPORTANTCONSTITUENT OF PROTEINS,COENZYMES, AND VITAMINS

Several forms of sulfur are found in most soils, includingiron sulfides and elemental sulfur. Sulfur is taken up byplants, however, as the divalent sulfate anion (SO2−

4 ).Sulfur deficiency is not a common problem becausethere are numerous microorganisms capable of oxidiz-ing sulfides or decomposing organic sulfur compounds.In addition, heavy consumption of fossil fuels in indus-try as well as natural phenomena such as geysers, hotsulfur springs, and volcanos together contribute largeamounts of sulfur oxides (SO2 and SO3) to the atmo-sphere. Indeed it is often difficult to demonstrate sulfurdeficiencies in greenhouses in industrial areas becauseof the high concentrations of airborne sulfur.

Sulfur is particularly important in the structure ofproteins where disulphide bonds (—S—S—) betweenneighboring cysteine and methionine residues con-tribute to the tertiary structure, or folding. Sulfur is alsoa constituent of the vitamins thiamine and biotin andof coenzyme A, an important component in respirationand fatty acid metabolism. In the form of iron-sulfurproteins, such as ferredoxin, it is important in elec-tron transfer reactions of photosynthesis and nitrogenfixation. The sulfur-containing thiocyanates and isoth-iocyanates (also known as mustard oils) are responsiblefor the pungent flavors of mustards, cabbages, turnips,horseradish, and other plants of the family Brassicaceae.Because of the presence of mustard oils, many speciesof the Brassicaceae prove fatal to livestock that grazeon them. Mustard oils also appear to serve as a defenseagainst insect herbivory.

Sulfur deficiency, like nitrogen, results in a general-ized chlorosis of the leaf, including the tissues surround-ing the vascular bundles. This is due to reduced proteinsynthesis rather than a direct impairment of chloro-phyll synthesis. However, chlorophyll is stabilized bybinding to protein in the chloroplast membranes. Withimpaired protein synthesis, the ability to form stablechlorophyll-protein complexes is also impaired. Unlikenitrogen, however, sulfur is not readily mobilized inmost species and the symptoms tend to occur initially inthe younger leaves.

4.4.6 CALCIUM IS IMPORTANT IN CELLDIVISION, CELL ADHESION, ANDAS A SECOND MESSENGER

Calcium is taken up as the divalent cation (Ca2+). Cal-cium is abundant in most soils and is seldom deficientunder natural conditions. Calcium is important to divid-ing cells for two reasons. It plays a role in the mitoticspindle during cell division and it forms calcium pec-tates in the middle lamella of the cell plate that formsbetween daughter cells. It is also required for the phys-ical integrity and normal functioning of membranesand, more recently, has been implicated as a secondmessenger in a variety of hormonal and environmentalresponses. As a second messenger involved in proteinphosphorylation, Ca2+ is an important factor in regu-lating the activities of a number of enzymes.

Because of its role in dividing cells, calcium defi-ciency symptoms characteristically appear in the meris-tematic regions where cell division is occurring and newcell walls are being laid down. Young leaves are typi-cally deformed and necrotic and, in extreme cases, deathof the meristem ensues. In solution cultures, calciumdeficiency results in poor root growth. The roots arediscolored and may feel ‘‘slippery’’ to the touch becauseof the deterioration of the middle lamella. Calcium isrelatively immobile and the symptoms typically appearin the youngest tissues first.

4.4.7 MAGNESIUM IS A CONSTITUENTOF THE CHLOROPHYLLMOLECULE AND AN IMPORTANTREGULATOR OF ENZYMEREACTION

Like calcium, magnesium is also taken up as the divalentcation (Mg2+). Magnesium is generally less abundantin soils than calcium but is required by plants in rela-tively large amounts. Magnesium deficiencies are mostlikely in strongly acid, sandy soils. Magnesium has sev-eral important functions in the plant. By far the largestproportion is found in the porphyrin moiety of thechlorophyll molecule, but it is also required to stabi-lize ribosome structure and is involved as an activator

4.4 Nutrient Functions and Deficiency Symptoms 71

for numerous critical enzymes. It is critical to reac-tions involving ATP, where it serves to link the ATPmolecule to the active site of the enzyme. Mg2+ is also anactivator for both ribulosebisphosphate carboxylase andphosphoenolpyruvate carboxylase, two critical enzymesin photosynthetic carbon fixation (Chapters 8 and 15).

The first and most pronounced symptom of mag-nesium deficiency is chlorosis due to a breakdown ofchlorophyll in the lamina of the leaf that lie betweenthe veins. Chloroplasts in the region of the veins are forsome reason less susceptible to magnesium deficiencyand retain their chlorophyll much longer. Magnesiumis also quite mobile. It is readily withdrawn from theolder leaves and transported to the younger leaves thatare more actively growing and synthesizing chlorophyll.Consequently, chlorosis due to Mg2+ deficiency is, atleast initially, most pronounced in the older leaves.

4.4.8 IRON IS REQUIRED FORCHLOROPHYLL SYNTHESISAND ELECTRON TRANSFERREACTIONS

Of all the micronutrients, iron is required by plants inthe largest amounts (it is considered a macronutrient bysome). Iron may be taken up as either the ferric (Fe3+) orferrous (Fe2+) ion, although the latter is more commondue to its greater solubility.3 The importance of ironis related to two important functions in the plant. It ispart of the catalytic group for many redox enzymesand it is required for the synthesis of chlorophyll.Important redox enzymes include the heme-containingcytochromes and non-heme iron-sulfur proteins (e.g.,Rieske proteins, ferredoxin, and photosystem I) involvedin photosynthesis (Chapter 7), respiration (Chapter 10)and nitrogen fixation (Chapter 11). During the courseof electron transfer the iron moiety is reversibly reduced

3In the scientific literature, particularly that body of literaturedealing with iron uptake by organisms, the ferric form of ironis also referred to as Fe(III) (iron-three). By the sameconvention, ferrous iron is referred to as FE(II) (iron-two).

from the ferric to the ferrous state. Iron is also a con-stituent of several oxidase enzymes, such as catalase andperoxidase.

Iron is not a constituent of the chlorophyll moleculeitself and its precise role in chlorophyll synthesis remainssomewhat of a mystery. There is, for example, nodefinitive evidence that any of the enzymes involvedin chlorophyll synthesis are iron-dependent. Instead,the iron requirement may be related to a more gen-eral need for iron in the synthesis of the chloroplastconstituents, especially the electron transport proteins.Iron deficiencies invariably lead to a simultaneous loss ofchlorophyll and degeneration of chloroplast structure.Chlorosis appears first in the interveinal regions of theyoungest leaves, because the mobility of iron in the plantis very low and it is not easily withdrawn from the olderleaves. Chlorosis may progress to the veins and, if thedeficiency is severe enough, the very small leaves mayactually turn white.

Iron deficiencies are common because of thepropensity of Fe3+ to form insoluble hydrous oxides(Fe2O3·3H2O) at biological pH. This problem isparticularly severe in neutral or alkaline calcareoussoils. On the other hand, iron is very soluble instrongly acidic soils and iron toxicity due to excessiron uptake can result. The problem of iron deficiencycan usually be overcome by providing chelated iron,either directly to the soil or as a foliar spray. A chelate(from the Greek, chele or claw) is a stable complexformed between a metal ion and an organic molecule,called a chelating agent or ligand. The ligand and themetal ion share electron pairs, forming a coordinatebond. Because chelating agents have a rather highaffinity for most metal ions, formation of the complexreduces the possibility for formation of insolubleprecipitates. At the same time, the metal can be easilywithdrawn from the chelate for uptake by the plant.One of the more common synthetic chelating agents isthe sodium salt of ethylenediaminetetraacetic acid(EDTA) (Figure 4.4), known commercially as versene orsequestrene. EDTA and similar commercially availablechelating agents, however, are not highly specific andwill bind a range of cations, including iron, copper,

FIGURE 4.4 Examples of organic acidsthat function as chelating agents.Ethylenediaminetetraacetic acid(EDTA) is a synthetic acid in commoncommercial use. Complexed with iron,it is sold under the trade name Versen-ate. Caffeic acid is one of several natu-rally occurring phenolic acids that maybe secreted by roots.

N CH2 CH2 N

OOC H2C CH2COO

OOC H2C CH2COO

Ethylenediamine tetraaceticacid (EDTA)

HO

OH

HC CCOO

H

Caffeic acid

–

–

–

–

–

72 Chapter 4 / Plants and Inorganic Nutrients

zinc, manganese, and calcium. Natural chelating agents,including porphyrins (as in hemoglobin, cytochromes,and chlorophyll) and a variety of organic and phenolicacids, are far more specific for iron.

The importance of iron in plant nutrition is high-lighted by the strategies plants have developed for uptakeunder conditions of iron stress. Iron deficiency inducesseveral morphological and biochemical changes in theroots of dicots and nongraminaceous monocots. These

ATP

ADP + Pi

Rhizosphere Membrane Cytosol

A.

B.

H+

FeIII

FeIII

FeIII

Soilcolloid

FeIII

FeIII

FeIII

FeII FeII

FeIII

Reductase

Ch

Ch

PS

PS PS

PS

FeIII

FeIII

FIGURE 4.5 Two strategies for the solubilization anduptake of sparingly soluble inorganic iron by higherplants. (A) ATPase proton pumps in the root corticalcells acidify the rhizosphere, which helps to solubilize asFe3+ (FeIII). The Fe3+ is then chelated by phenolic acids(Ch), also secreted into the rhizosphere by the roots. Thechelated iron is carried to the root surface where it isreduced by an FeIII reductase. The resulting Fe2+ (FeII) isimmediately transported across the plasma membrane byan FeII transporter. Both the FeIII reductase and the FeII

are induced by iron deficiency. (B) Fe3+ is solubilized byphytosiderophores (PS) secreted into the rhizosphere bythe root. The entire ferrisiderophore (siderophore-ironcomplex) is then taken into the root cell where the ironis subsequently released.

include the formation of specialized transfer cells in theroot epidermis, enhanced proton secretion into the soilsurrounding the roots, and the release of strong ligands,such as caffeic acid (Figure 4.4), by the roots. Simul-taneously, there is an induction of reducing enzymesin the plasma membrane of the root epidermal cells.Acidification of the rhizosphere encourages chelation ofthe Fe3+ with caffeic acid, which then moves to the rootsurface where the iron is reduced to Fe2+ at the plasmamembrane (Figure 4.5A). Reduction to Fe2+ causes theligand to release the iron, which is immediately takenup by the plant before it has the opportunity to forminsoluble precipitates.

A second strategy for iron uptake by organismsinvolves the synthesis and release by the organism oflow-molecular-weight, iron-binding ligands calledsiderophores (Gr. iron-bearers). Most of ourknowledge of siderophores comes from studies withaerobic microorganisms (bacteria, fungi, and algae),where they were first discovered and have been studiedmost extensively. More recently, however, it hasbeen discovered that siderophores are also releasedby the roots of higher plants (Figure 4.6). Known asphytosiderophores, to distinguish them from ligandsof microbial origin, these highly specific iron-bindingligands have thus far been found only in membersof the family Gramineae, including the cereal grains.Phytosiderophores are synthesized and released by theplant only under conditions of iron stress, have a highaffinity for Fe3+, and very effectively scavenge ironfrom the rhizosphere. The distinctive feature of thesiderophore system is that the entire iron-phytosiderophorecomplex, or ferrisiderophore, is then reabsorbed into theroots (Figure 4.5B). Once inside the root, the iron ispresumably reduced to Fe2+ and released for use bythe cell. The fate of the phytosiderophore is unknown.In microorganisms, siderophores may be chemicallydegraded and metabolized or, alternatively, the same

HO

COO COO COOH

OHNH2

+NH2

+

COOH

OH

OHH

COO

NH+ NH2

+

Avenic acid (AA)

Muginec acid (MA)

– –

–COO–

FIGURE 4.6 Phytosiderophores. The structures of twophytosiderophores released by the roots of higher plants.Ferric iron forms coordinate bonds with the nitrogen andcarboxyl groups.

4.4 Nutrient Functions and Deficiency Symptoms 73

molecule may again be secreted by the cell in order topick up more iron.

The study of phytosiderophores is a relatively youngfield and, although substantial progress has been madein recent years, there is still much to be learned. It is notyet known, for example, how widespread the use of phy-tosiderophores is and the nature of the ferrisiderophoretransport system has not been demonstrated in plants.One thing is clear: in those plants that use them, phy-tosiderophores are an important and effective strategyfor supplying iron to the plant under conditions of ironstress.

4.4.9 BORON APPEARS TO HAVEA ROLE IN CELL DIVISIONAND ELONGATION ANDCONTRIBUTES TO THESTRUCTURAL INTEGRITYOF THE CELL WALL

In aqueous solution, boron is present as boric acid, orH3BO3. At physiological pH (<8), it is found predomi-nantly in the undissociated form, which is preferred foruptake by roots. With respect to its biochemical andphysiological role, boron is perhaps the least under-stood of all the micronutrients. There is, for example,no solid evidence for involvement of boron with specificenzymes, either structurally or as an activator. Indeed,most of what we know about the role of boron is basedentirely on studies of what happens to plants when boronis withheld.

A substantial proportion of the total borate contentof cells is found in the cell wall. This is apparentlybecause borate has a propensity to form stable esterswith cell wall saccharides that have adjacent hydroxylgroups. This so-called cis-diol configuration is char-acteristic of some common cell wall polysaccharides,such as mannose and its derivatives. Glucose, fructose,and galactose, on the other hand, do not have thisconfiguration and so do not bind boron. The primarywalls of boron-deficient cells exhibit marked structuralabnormalities, suggesting that boron is required for thestructural integrity of the cell wall.

Other responses to boron deficiency point towarda role in cell division and elongation. One of the mostrapid responses to boron deficiency, for example, isan inhibition of both cell division and elongation inprimary and secondary roots. This gives the roots astubby and bushy appearance. Cell division in the shootapex and young leaves is also inhibited, followed bynecrosis of the meristem. In addition, boron is knownto stimulate pollen tube germination and elongation. Itis not known how boron is involved in cell growth, butboth hormone and nucleic acid metabolism have beenimplicated. Inhibition of cell division and elongation isaccompanied by an increased activity of enzymes that

oxidize the hormone indole-3-acetic acid and a decreasein RNA content (possibly through impaired synthesis ofuracil, an RNA precursor).

In addition to the effects on shoot meristems notedabove, common symptoms of boron deficiency includeshortened internodes, giving the plant a bushy or rosetteappearance, and enlarged stems, leading to the disorderknown as ‘‘stem crack’’ in celery. In storage roots such assugar beets, the disorder known as ‘‘heart rot’’ is due tothe death of dividing cells in the growing region becauseof boron deficiency.

4.4.10 COPPER IS A NECESSARYCOFACTOR FOR OXIDATIVEENZYMES

In well-aerated soils, copper is generally available tothe plant as the divalent cupric ion, Cu2+. Cu2+ readilyforms a chelate with humic acids in the organic fractionof the soil and may be involved in providing copper tothe surface of the root. In wet soils with little oxygen,Cu2+ is readily reduced to the cuprous form, Cu+, whichis unstable. As a plant nutrient, copper seems to functionprimarily as a cofactor for a variety of oxidative enzymes.These include the photosynthetic electron carrier plas-tocyanin; cytochrome oxidase, which is the final oxidaseenzyme in mitochondrial respiration; and ascorbic acidoxidase. The browning of freshly cut apple and potatosurfaces is due to the activity of copper-containingpolyphenoloxidases (or phenolase). Superoxide dis-mutase (SOD), which detoxifies superoxide radicals(O−

2 ), is another important copper enzyme.Common disorders due to copper deficiency are

generally stunted growth, distortion of young leavesand, particularly in citrus trees, a loss of young leavesreferred to as ‘‘summer dieback.’’

4.4.11 ZINC IS AN ACTIVATOROF NUMEROUS ENZYMES

Zinc is taken up by roots as the divalent cation Zn2+.Zinc is an activator of a large number of enzymes, includ-ing alcohol dehydrogenase (ADH), which catalyzesthe reduction of acetaldehyde to ethanol; carbonicanhydrase (CA), which catalyzes the hydration of car-bon dioxide to bicarbonate; and, copper SOD whichdetoxifies O2

−. However, there is general agreementthat disorders associated with zinc deficiency reflect dis-turbances in the metabolism of the auxin hormoneindole-3-acetic acid. Typically, zinc-deficient plantshave shortened internodes and smaller leaves (e.g., ‘‘lit-tle leaf ’’ disorder of fruit trees). The precise role of zincin auxin metabolism remains obscure, but auxin lev-els in zinc-deficient plants are known to decline beforethe overt symptoms of zinc deficiency appear. Further-more, restoration of the zinc supply is followed by a

74 Chapter 4 / Plants and Inorganic Nutrients

rapid increase in hormone level and then resumptionof growth. Available evidence supports the view thatzinc is required for synthesis of the hormone precursortryptophan.

4.4.12 MANGANESE IS AN ENZYMECOFACTOR AS WELL AS PARTOF THE OXYGEN-EVOLVINGCOMPLEX IN THECHLOROPLAST

Manganese is absorbed and transported within the plantmainly as the divalent cation Mn2+. Manganese isrequired as a cofactor for a number of enzymes, particu-larly decarboxylase and dehydrogenase enzymes, whichplay a critical role in the respiratory carbon cycle. Inter-estingly, manganese can often substitute for magnesiumin reactions involving, for example, ATP. However, thebest known and most studied function of manganese isin photosynthetic oxygen evolution (Chapter 7). In theform of a manganoprotein, manganese is part of theoxygen-evolving complex associated with photosystemII, where it accumulates charges during the oxidation ofwater.

Manganese deficiency can be widespread in someareas, depending on soil conditions, weather, and cropspecies. Deficiency is aggravated by low soil pH (<6)and high organic content. Manganese deficiency isresponsible for ‘‘gray speck’’ of cereal grains, a dis-order characterized by the appearance of greenish-gray,oval-shaped spots on the basal regions of young leaves.It may cause extreme chlorosis between the leaf veins aswell as discoloration and deformities in legume seeds.

4.4.13 MOLYBDENUM IS A KEYCOMPONENT OF NITROGENMETABOLISM

Although molybdenum is a metal, its properties moreclosely resemble those of the nonmetals. In aqueoussolution it occurs mainly as the molybdate ion MoO2−

4 .Molybdenum requirements are among the lowest ofall known micronutrients and appear to be primarilyrelated to its role in nitrogen metabolism. Among theseveral enzymes found to require molybdenum are dini-trogenase and nitrate reductase. The molybdenumrequirement of a plant thus depends to some extenton the mode of nitrogen supply (Chapter 11). Dini-trogenase is the enzyme used by prokaryotes, includingthose in symbiotic association with higher plants, toreduce atmospheric nitrogen. Nitrate reductase is foundin roots and leaves where it catalyzes the reduction ofnitrate to nitrite, a necessary first step in the incorpora-tion of nitrogen into amino acids and other metabolites.

In plants such as legumes, which depend on nitrogenfixation, molybdenum deficiency gives rise to symptoms

of nitrogen deficiency. When nitrogen supplies are ade-quate, a deficiency of molybdenum shows up as a classicdisorder known as ‘‘whiptail’’ in which the young leavesare twisted and deformed. The same plants may exhibitinterveinal chlorosis and necrosis along the veins ofolder leaves. Like many of the micronutrients, molyb-denum deficiency is highly species dependent—it isparticularly widespread for legumes, members of thefamily Brassicaceae, and for maize. Molybdenum defi-ciency is aggravated in acid soils with a high content ofiron precipitates, which strongly adsorb the molybdateion.

4.4.14 CHLORINE HAS A ROLE INPHOTOSYNTHETIC OXYGENEVOLUTION AND CHARGEBALANCE ACROSS CELLULARMEMBRANES

Chloride ion (Cl−) is ubiquitous in nature and highlysoluble. It is thus rarely, if ever, deficient. Deficienciesnormally can be shown only in very carefully controlledsolution culture experiments. Along with manganese,chloride is required for the oxygen-evolving reactionsof photosynthesis (Chapter 7). Cl− is a highly mobileanion with two principal functions: it is both a majorcounter-ion to diffusible cations, thus maintaining elec-trical neutrality across membranes, and one of theprincipal osmotically active solutes in the vacuole. Chlo-ride ion also appears to be required for cell division inboth leaves and shoots. Chloride is readily taken up andmost plants accumulate chloride ion far in excess of theirminimal requirements. Plants deprived of chloride tendto exhibit reduced growth, wilting of the leaf tips, and ageneral chlorosis.

4.4.15 THE ROLE OF NICKELIS NOT CLEAR

Nickel is a relatively recent addition to the list of essen-tial nutrient elements. Nickel is an abundant metallicelement and is readily absorbed by roots. It is ubiq-uitous in plant tissues, usually in the range of 0.05 to5.0 mg Kg−1 dry weight. One of the principal difficultiesencountered in attempting to establish a role for nickelis its extremely low requirement. It has been estimatedthat the quantity of nickel needed by a plant to completeone life cycle is approximately 200 ng, a requirementthat can be met by the initial nickel content of the seedin most cases. In order to establish a nickel deficiency,it is necessary to undertake extensive purification of thenutrient salts and then grow several successive genera-tions to seed in nickel-deficient solutions. The strongestevidence in favor of essential status for nickel is basedon studies with legumes and cereal grains. In one suchstudy of barley (Hordeum vulgare), the critical nickel

Summary 75

concentration for seed germination was found to be90 ng g−1 of seed dry weight. By growing plants forthree generations in the absence of nickel, the nickelcontent of the seed could be reduced to 7.0 ng g−1 dryweight. Germination of these seeds was less than 12 per-cent. When the plants were grown for the same numberof generations in nutrient solution supplemented with0.6 μM or 1.0 μM nickel, seed germination was 57 and95 percent, respectively. In other studies, nickel defi-ciencies have led to depressed seedling vigor, chlorosis,and necrotic lesions in leaves.

The basis for a nickel requirement by plants isnot clear, but it may be related to mobilization ofnitrogen during seed germination. Nickel is known to bea component of two enzymes; urease and hydrogenase.Urease catalyzes the hydrolysis of urea into NH3 andCO2 and is found widely through the plant kingdom.Urease from jack bean seeds (Canavalia ensiformis) wasin fact the first protein to be crystallized by J. B. Sumnerin 1926. One of the principal effects of nickel defi-ciency in soybean (Glycine max) is decreased ureaseactivity in the leaves, although the metabolic significanceof urease is not yet clear.

Free urea is rarely, if ever, detected in plant tissue,but it is formed by the action of the enzyme arginaseon arginine and its structural analog, canavanine(Chapter 11). Canavanine, a nonprotein amino acid, isabundant in the seeds of some plant groups, such as jackbean, but its concentration diminishes rapidly upongermination. Arginine is also abundant in seeds andboth amino acids could function as stored nitrogen thatis readily mobilized during seed germination. If thisview should be proven valid, then urease, and thus nickelas well, would play an important role in the mobilizationof nitrogen during germination and early seedlinggrowth.

A common form of mobile nitrogen in somelegumes is a family of urea-based compounds known asureides, such as allantoic acid or citrulline (Chapter 11).Ureides are formed in root nodules during nitrogenfixation and transported via the xylem throughoutthe host plant. Ureides are also formed in senescingleaves and transported out to the developing seedsfor storage. The breakdown of ureides produces urea,which accumulates to toxic levels in Ni-deficient plants.Furthermore, the metabolism of purine bases (adenineand guanine) in all plants also produces ureides. Itseems reasonable to assume that most, if not all, plantshave a requirement for urease and nickel.

Hydrogenase is another important enzyme in somenitrogen-fixing plants. Hydrogenase is responsible forrecovering hydrogen for use in the nitrogen-fixingprocess (Chapter 11). A deficiency of nickel leads todepressed levels of hydrogenase activity in the nodulesof soybean, which in turn would be expected to depressthe efficiency of nitrogen fixation.

4.5 TOXICITY OFMICRONUTRIENTS

As a group, the micronutrient elements are an excellentexample of the dangers of excess. Most have a rather nar-row adequate range and become toxic at relatively lowconcentrations. Critical toxicity levels, defined as thetissue concentration that gives a 10 percent reduction indry matter, vary widely between the several micronutri-ents as well as between plant species. As noted earlier,critical concentrations for copper, boron, and zinc areon the order of 20, 75, and 200 μg g−1 dry weight,respectively. On the other hand, critical toxicity lev-els for manganese vary from 200 μg g−1 dry weight forcorn, to 600 μg g−1 for soybean, and 5300 μg g−1 for sun-flower. Toxicity symptoms are often difficult to decipherbecause an excess of one nutrient may induce deficienciesof other nutrients. For example, the classic symptom ofmanganese toxicity, which often occurs in waterloggedsoils, is the appearance of brown spots due to deposi-tion of MnO2 surrounded by chlorotic veins. But excessmanganese may also induce deficiencies of iron, mag-nesium, and calcium. Manganese competes with bothiron and magnesium for uptake and with magnesium forbinding to enzymes. Manganese also inhibits calciumtranslocation into the shoot apex, causing a disorderknown as ‘‘crinkle leaf.’’ Thus the dominant symptomsof manganese toxicity may actually be the symptoms ofiron, magnesium, and/or calcium deficiency.

Excess micronutrients typically inhibit root growth,not because the roots are more sensitive than shoots butbecause roots are the first organ to accumulate the nutri-ent. This is particularly true of both copper and zinc.Copper toxicity is of increasing concern in vineyardsand orchards due to long-term use of copper-containingfungicides as well as urban and industrial pollution. Zinctoxicity can be a problem in acid soils or when sewagesludge is used to fertilize crops.

In spite of the apparent toxicity of micronutrients,many plant species have developed the capacity to tol-erate extraordinarily high concentrations. For example,most plants are severely injured by nickel concentrationsin excess of 5 μg g−1 dry weight, but species of the genusAlyssum can tolerate levels in excess of 10 000 μg g−1 dryweight.

SUMMARY

Plants are autotrophic organisms, taking their entirenutritional needs from the inorganic environment.Plants require carbon, hydrogen, and oxygen, plus14 other naturally occurring elements that are takenfrom the soil. These 17 elements are consideredessential because it has been demonstrated that in their

76 Chapter 4 / Plants and Inorganic Nutrients

absence all plants are unable to complete a normallife cycle. Essential elements may be consideredeither macronutrients or micronutrients, dependingon the quantity normally required. Micronutrientsare normally required in concentrations less than10 mmole/kg of dry weight.

Each essential element has a role to play in the bio-chemistry and physiology of the plant and its absenceis characterized by one or more deficiency symptoms,commonly related to that role. Additional elementsmay be considered beneficial because they satisfy spe-cial requirements for particular plants. Essential ele-ments, especially micronutrients, may be toxic whenpresent in excess amounts.

CHAPTER REVIEW

1. Explain the difference between autotrophic andheterotrophic nutrition.

2. What is meant by essentiality? What is thedifference between an essential element and abeneficial element? Describe the steps you wouldgo through in order to determine the essentialityor nonessentiality of an element for a higher plant.

3. List the 17 elements that are essential forthe growth of all higher plants. Be able toidentify one or more principal structural ormetabolic roles for each essential element.

4. Deficiencies of iron, magnesium, and nitrogenall cause chlorosis. Iron chlorosis develops onlybetween the veins of young leaves while chlorosisdue to both magnesium and nitrogen deficienciesdevelops more generally in older leaves. Explainthese differences. Why does each deficiency leadto chlorosis and why are the patterns different?

5. For what reasons might a soil rich in calciumsupply too little phosphorous for plant growth?

6. What is a chelating agent? Explain howchelating agents help to maintain ironavailability in nutrient cultures and soils.

7. What is meant by critical toxicity level? Whichelements are most likely to be both essential andtoxic to plants?

8. There are currently 17 elements known tobe essential for higher plants. Is it possi-ble that other elements might be added tothis list in the future? Explain your answer.

FURTHER READING

Blevins, D. G, K. M. Lukaszewski. 1999. Boron and plantstructure and function. Annual Review of Plant Biology49: 481–500.

Broadley, M. R., P. J. White. 2005. Plant NutritionalGenomics. Oxford, U.K.: Blackwell Publishing.

Buchanan, B. B., W. Gruissem, R. L. Jones. 2000.Biochemistry and Molecular Biology of Plants. Rockville,MD: American Society of Plant Physiologists.

Epstein, E., A. J. Bloom. 2005. Mineral Nutrition of Plants:Principles and Perspectives. Sunderland: Sinauer AssociatesInc.

Kochıan, L. V., O. A. Hoekenga, M. A. Pineros. 2004.How do crop plants tolerate acid soils? Mechanisms ofaluminum tolerance and phosphate efficiency. AnnualReview of Plant Biology 55: 459–493.

Salt, D. E. 2008. Ionomics and the study of the plant ionome.Annual Review of Plant Biology 59: 709–733.

Schachtman, D. P., R. Shin. 2007. Nutrient sensing: NPKS.Annual Review of Plant Biology 58: 47–69.

α α

αβ

β

β

γ

δ

ε

Lumen

Stroma

ADP + Pi

CF1

CF0

ATP

3H+

b

c

3H+

a

5Bioenergetics and ATP Synthesis

A unique characteristic of planet earth is the presenceof life. On a grand scale, one can consider the entireearth’s biosphere to be an enormous, exquisite, but com-plex energy-transforming system consisting of myriadcountless organisms. In addition to water (Chapters1–4), energy is an absolute requirement for the main-tenance and replication of life regardless of its form.Each organism plays a specific role in this teeming webof carbon-based life forms. Regardless of whether weexamine the scale of biology at the community, indi-vidual, cellular, or molecular level, organization is thevery essence of life and yet it is constantly under attack.At the cellular level, proteins, nucleic acids, and othermolecules that make up the cell are continually subjectto breakdown by hydrolysis. Membranes leak solutes tothe environment. Everything on earth, cells and envi-ronment alike, is subject to persistent oxidation. Still, allaround us we see biological organisms extracting mate-rials from their environment and using them to maintaintheir organization or to build new, complex structures.Energy to build and preserve order in the face of aconstantly deteriorating environment is a fundamentalneed of all organisms. Two strategies have evolved tosatisfy this need. One is photosynthesis—the pho-toautotropic lifestyle—which traps energy from the

sun to build complex structures out of simple inor-ganic substances. By contrast, organisms that live bythe alternative lifestyle, chemoheterotropic, require aconstant intake of organic substances from their envi-ronment, from which they can extract their necessaryenergy through respiration. But even many of thesesubstances trace their origins back to photosynthesis. Inthe end, most life on earth is powered by energy fromthe sun through photosynthesis. This chapter is con-cerned with the basic principles of bioenergetics—thestudy of energy transformations in living organisms.

The principal topics to be covered are

• thermodynamic laws and the concepts of free energyand entropy,

• free energy and its relationship with chemical equi-libria, illustrating how displacement of a reactionfrom equilibrium can be used to drive vital reactions,

• oxidation–reduction reactions, showing how theyalso are involved in biological energy transforma-tions, and

• the chemiosmotic model for synthesis of adenosinetriphosphate (ATP), a key mediator of biologicalenergy metabolism.

77

78 Chapter 5 / Bioenergetics and ATP Synthesis

5.1 BIOENERGETICS ANDENERGY TRANSFORMATIONSIN LIVING ORGANISMS

5.1.1 THE SUN IS A PRIMARY SOURCEOF ENERGY

Given the complex composition and organization ofour biosphere, it may seem surprising that the basicingredients required to sustain most life on earth arerather simple: water, visible light, and air. Light maybe considered the ultimate form of energy requiredto maintain most carbon-based life forms. The sourceof this light of course is the sun. How is this lightgenerated? The thermonuclear fusion reactions in theheart of this star convert four protons (4H+) to onehelium (He) atom, which has an atomic weight of 4.0026.However, since each H+ has an atomic weight of 1.0079,the expected atomic weight of He should be 4.0316.Clearly, we are missing 0.0290 gm-atoms of mass, whichrepresents a mere 0.72 percent of the total mass of4H+! Einstein showed us that there is a very importantrelationship between energy and mass:

E = mc2 (5.1)

where E is energy, m is mass, and c is the speed of light.Thus, the missing mass (m) of 0.0290 gm-atoms duringthe conversion of 4H+ to He is converted into energy(E) in the form of electromagnetic radiation. A smallportion of this electromagnetic energy is in the formof visible light (Chapter 6), which reaches the earth’ssurface after a trip of about 160 million km. Since thespeed of light is 300,000 km s−1, each photon of visiblelight generated by the sun requires 8.88s to reach theearth’s surface. This means that any image of the sunthat we detect on earth can never be an original image,but rather, an image of the sun that is 8.88s old!

Air provides the basic elements for all living organ-isms: C, N, and O. C in the air is in the form of CO2(about 0.035%) and the N in air is in the form of N2(about 80%). However, most living organisms cannotdirectly utilize either CO2 or N2 as a source of C andN, respectively. Water provides the solvent necessaryfor enzyme catalysis and formation of biological mem-branes (Chapters 1 and 2). In Chapters 7 to 11 we willaddress the role played by plants in utilizing light energyto transform C and N into forms required not only tosustain plant life but all forms of life. However, it isessential that we establish some understanding of thebasic principles that govern energy transformations inbiological systems, that is, bioenergetics.

5.1.2 WHAT IS BIOENERGETICS?

In 1944, the Nobel Laureate in physics, ErwinSchrodinger, published an intriguing little monograph

entitled What Is Life? In this book, this famous physicistattempts to unravel the basis of life on physical andchemical principles. Schrodinger simply asked whetherthe laws of physics and chemistry can account for thecomplex events that take place within ‘‘the spatialboundary of a living organism’’ through space and time.Schrodinger used a thermodynamic approach to addressthis question. The term thermodynamics and muchof its language and mathematics reflect an historicalinterest to discover the fundamental laws that governheat flow. Although the study of thermodynamics isnow concerned with energy flow in a more generalsense, the science of thermodynamics arose fromnineteenth-century interests in the workings of steamengines or why heat was evolved when boring cannonbarrels. Energy flow is governed by certain fundamentalthermodynamic rules. A general understanding ofthermodynamic principles is necessary because theseprinciples provide the quantitative framework forunderstanding energy transformations in biology. Inaddition to energy transformations, thermodynamicsalso helps to describe the capacity of a system to dowork. Work may be defined in several different ways.The physicist defines work as displacement againsta force: the sliding of an object against friction orrolling a boulder uphill, for example. The chemist, onthe other hand, views work in terms of pressure andvolume. For example, work must be done to overcomethe force of atmospheric pressure when the volume of agas increases. In biology the concept of work is appliedmore broadly, embracing a variety of work functionsagainst a wide spectrum of forces encountered in cellsand organisms. In addition to mechanical work such asmuscular activity, the biologist is concerned with suchdiverse activities as chemical syntheses, the movementof solute against electrochemical gradients, osmosis,and ecosystem dynamics. These and a host of otheressential activities of living things can all be describedin thermodynamic terms.

Bioenergetics is the application of thermodynamiclaws to the study of energy transformations in biologi-cal systems. The energetics of cellular processes can berelated to chemical equilibrium and oxidation–reductionpotentials of chemical reactions. Whether at the levelof molecules, cells, or ecosystems, the flow of energyis central to the maintenance of life. A basic under-standing of energy flow is therefore essential to graspingthe true beauty, significance, and complexity of biology.The field of study concerned with the flow of energythrough living organisms is called bioenergetics. It isimportant to note that although the laws of bioenerget-ics provide critical insight into the driving forces thatgovern cellular processes, bioenergetics does not provideany insights into the biochemical reaction mechanismsunderlying these processes. For the past several decades,a central focus of bioenergetics has been to unravel the

5.1 Bioenergetics and Energy Transformations in Living Organisms 79

complexities of energy transformations in photosynthe-sis and respiration and understand how that energy isused to drive energy-requiring reactions such as ATPsynthesis and accumulation of ions across membranes.

The purpose of this section is to facilitate an under-standing of the laws that govern biological energy trans-formations by assembling, in the simplest form possible,some basic thermodynamic principles. Thus, a com-plete understanding of cellular physiology requires anintegration of bioenergetics with biochemical reactionmechanisms which is a pervading theme of this textbook.The interested student will find a more comprehen-sive treatment of thermodynamics and bioenergetics inthe excellent monograph by D. G. Nicholls and S. J.Stuart (2002). This publication provided the basis forthe discussion that follows.

5.1.3 THE FIRST LAW OFTHERMODYNAMICS REFERSTO ENERGY CONSERVATION

Biological energy transformations are based on twothermodynamic laws. The first law, commonly knownas the law of conservation of energy, states that theenergy of the universe is constant. This is not a difficultconcept to comprehend—it means simply that there isa fixed amount of energy and, while it may be movedabout or changed in form, it can all be accounted forsomewhere. More to the point, energy is never ‘‘lost’’ ina reaction—an apparent decrease in one form of energywill be balanced by an increase in some other form ofenergy. In one of the examples mentioned earlier, someof the energy expended in displacing an object appearsas work while some appears as heat generated due tofriction. In the same way, some of the chemical energyreleased in the combustion of glucose will also be foundas heat in the environment, while some will be found asbond energy in the product molecules, CO2 and water.

5.1.4 THE SECOND LAW OFTHERMODYNAMICS REFERSTO ENTROPY AND DISORDER

As biologists we are concerned above all with howmuch work can be done. But, as suggested earlier, notall energy is available to do work. This brings us tothe second law of thermodynamics and the concept ofentropy. Because it involves the concept of entropy (S),the second law is a bit more difficult to comprehend.What is entropy? Entropy has been variously describedas a measure of randomness, disorder, or chaos.

However, since entropy is a thermodynamic con-cept, it is useful to describe it in terms of thermalenergy. Temperature is defined as the mean molecularkinetic energy of matter. Thus, any molecular systemnot at absolute zero (−273◦C, or 0◦K) contains a certain

amount of thermal energy—energy in the form of thevibration and rotation of its constituent molecules aswell as their translation through space. This quantityof thermal energy and temperature go hand-in-hand:as the quantity of energy increases or decreases, sodoes temperature. Because temperature cannot be heldconstant when this energy is given up, it is said to be‘‘isothermally unavailable’’ (Gr. isos, equal). Quantita-tively, isothermally unavailable energy is given by theterm TS, where T is the absolute temperature and S isentropy.

Since isothermally unavailable energy, and conse-quently, entropy, are related to the energy of molecularmotion, it follows that the more molecules are free tomove about, that is, the more random or less ordered orchaotic the system, the greater will be their entropy. Bythis same argument it also follows that at absolute zero,a state in which all molecular motion ceases, entropy isalso zero. Consider a familiar example: the combustionof glucose. The highly ordered structure of a glucosemolecule imposes certain constraints on the movementof the constituent carbon atoms. In the form of six indi-vidual carbon dioxide molecules, however, those sameatoms are far less constrained. They are individuallyfree to rotate and tumble through space. With respectto carbon, the glucose molecules and carbon dioxidemolecules each have entropy, but the product carbondioxide molecules are less ordered, their freedom ofmovement is greater, and so is their entropy. It is impor-tant to note, however, that for any given reaction, theentropy of all reactants and all products must be takeninto consideration. For a system not at rest, the naturaltendency is for entropy to increase, that is, for systemsto become increasingly chaotic. Equation 5.2 shows therelationship between entropy, S, and disorder, D, wherek is Boltzmann’s constant (1.3800662 × 10−23 J · ◦K−1).Thus, as disorder increases so does entropy.

S = k log D (5.2)

This tendency was summarized by R. J. Clausius: theentropy of the universe tends toward a maximum. Clausius’sdictum is one way of stating the second law of thermody-namics. The primary characteristic of all life is complexorder which appears to contradict the second law ofthermodynamics. However, this is not the case since allliving organisms exhibit a finite lifetime which can varyfrom hours to days to years to centuries depending onthe species. Since order (1/D) is the inverse of disorder(D), Equation 5.2 becomes

−S = k log(1/D) (5.3)

Thus, life is negative entropy (−S) !As physiologists, however, our concern with entropy

is primarily that it represents energy that is not availableto do work. In this context, the second law can thenbe restated as: the capacity of an isolated system to do work

80 Chapter 5 / Bioenergetics and ATP Synthesis

continually decreases. In other words, it is never possibleto utilize all of the energy of a system to do work.

5.1.5 THE ABILITY TO DO WORKIS DEPENDENT ON THEAVAILABILITY OF FREE ENERGY

From the above discussion, it is apparent that someenergy will be available under isothermal conditions andis, consequently, available to do work. This energy iscalled Gibbs free energy in honor of J. W. Gibbs, thenineteenth-century physical chemist who introducedthe concept. Free energy (G) is related to TS in thefollowing way:

H = G + TS (5.4)

H is the total heat energy (also called enthalpy), includ-ing any work that might be done. H is comprised ofisothermally unavailable TS plus G. Equation 5.4 thusidentifies two kinds of energy: free energy, which isavailable to do work, and entropy, which is not. Exceptin a limited number of situations, it is free energy, theenergy available to do work, that is of greatest interestto the biologist. Equation 5.4 also suggests a corollary ofthe second law: the free energy of the universe tends towarda minimum.

It is neither convenient nor relevant to measureabsolute energies (either G or S), but changes (designatedby the symbol �) in energy during the course of areaction can usually be measured with little difficulty as,for example, heat gain or loss, or work. Equation 5.4 canbe restated as follows:

�G = �H − T�S (5.5)

Changes in free energy can tell us much about a reaction.It can tell us, for example, the feasibility of a reactionactually taking place and the quantity of work that mightbe done if it does take place. Feasibility is indicatedby the sign of �G. If the sign of �G is negative (i.e.,�G < 0), the reaction is considered spontaneous, meaningthat it will proceed without an input of energy. Sincethe free energy of the products is less than the reactants,reactions with a negative �G are sometimes known asexergonic, or energy yielding. If, on the other hand,�G is positive, an input of energy is required for thereaction to occur. The oxidation of glucose is an exampleof a reaction with a negative �G. Once an activationbarrier (see Chapter 8 for details) is overcome, glucosewill spontaneously oxidize to form CO2 and water.Despite the existence of large quantities of CO2 andwater in the atmosphere, however, they are not knownto spontaneously recombine to form glucose! This isbecause the equilibrium constant favors the formationof CO2 and H2O and the �G for glucose formationis positive. Reactions with a positive �G (i.e., �G >

0) are known as endergonic, or energy consuming.Equation 5.5 also tells us that any change in free energy

is associated with a change in entropy. This means thatthere is always a price to pay in any reaction involvedin an energy transformation. The price paid is in theinevitable increase in entropy. Therefore, no energytransformation process can be 100 percent efficient withrespect to the retention of free energy.

The magnitude of free energy changes is very mucha function of the particular set of conditions for thatreaction. For that reason it is convenient to comparethe free energy changes of reactions under standardreaction conditions. In biochemistry the standard freeenergy change, �G◦′, defines the free energy changeof a reaction that occurs at physiological pH (pH = 7.0)under conditions where both reactants and products areat unit concentration (1 M).

5.1.6 FREE ENERGY IS RELATEDTO CHEMICAL EQUILIBRIA

Under appropriate conditions, all chemical reactionswill achieve a state of equilibrium, at which there will beno further net change in the concentrations of reactantsand products. There is a fairly straightforward rela-tionship between free energy and chemical equilibria.This relationship, which is central to an understand-ing of bioenergetics, is illustrated diagrammatically inFigure 5.1. In a reaction where the reactant A is con-verted to product B, K is the equilibrium mass-actionratio—the ratio of concentration of products to theconcentration of reactants when the reaction has cometo equilibrium. Thus

Keq = [B]eq/[A]eq (5.6)In Figure 5.1 the slope of the line represents the changein free energy (�G) when a small amount of the reactantA is converted to the product B. Several useful pointscan be drawn from this diagram.

1. At equilibrium, the slope of the line is zero. Con-sequently, when reactions are at equilibrium, �G = 0and no useful work can be accomplished.

2. The further the mass-action ratio is displacedfrom equilibrium (Keq), the greater the free energychange for conversion of the same small amount ofA to B. The free energy change for a reaction isa function of its displacement from equilibrium.Therefore, the further a reaction is poised away fromequilibrium, the more free energy is available as thereaction proceeds toward equilibrium.

3. As A approaches equilibrium, �G is negative andfree energy is available to do work. However, as thereaction proceeds past equilibrium toward B, �Gis becomes positive and energy must be supplied.A system can do work as it moves toward equilibrium.Note that if the reaction were initiated with pure B,the direction of the arrows would be reversed andwork could be done as B approached equilibrium.

5.2 Energy Transformations and Coupled Reactions 81

Reactant A B

0.001K 0.01K 0.1K K

Ratio[Product][Reactant]

10K 100K 1000K

Product

Free

ene

rgy

(G)

ΔG ΔG

ΔG

ΔG

ΔG

ΔG

FIGURE 5.1 The free energy of a reaction is a function of its displacement fromequilibrium. K is the mass-action ratio when the reaction is at equilibrium. Verti-cal arrows indicate the slope of the free energy curve, or change in free energy, asreactant is converted to product. Note that the free energy change at equilibrium iszero and that the magnitude of the free energy change, indicated by the length ofthe arrow, increases as the reaction moves away from equilibrium toward pure reac-tant or pure product. A downward arrow indicates a negative free energy change; anupward arrow indicates a positive free energy change. (Redrawn from D. G. Nichollsand S. J. Ferguson, Bioenergetics, New York: Academic Press, 2002. Reprinted bypermission.)

The relationship between the free energy change (�G)and the equilibrium constant (Keq) can be expressedquantitatively as:

�G = �G◦′ + RT ln Keq (5.7)

However, at equilibrium, �G = 0 and Equation 5.7 canbe rearranged to give

�G◦′ = −RT ln Keq = −2.3RT log Keq (5.8)

Furthermore, the actual free energy change (�G) of areaction not at equilibrium is given by:

�G = �G◦′ + 2.3RT log � (5.9)

where R is the universal gas constant, T is the absolutetemperature, and � equals the observed (i.e., nonequi-librium) mass-action ratio. Equation 5.8 can then besubstituted in Equation 5.9 and rearranged to give:

�G = −2.3RT log(Keq/�) (5.10)

Equation 5.10 reinforces the observation that the valueof �G is a function of the degree to which a reaction isdisplaced from equilibrium. When � = Keq, the reactionis at equilibrium and �G = 0 and no useful work can bedone. When � is less than Keq, �G < 0 and the reactionwill occur spontaneously with the release of energywhich can be used to perform useful work. However,when � is greater than Keq, �G > 0 and the reaction

will not occur spontaneously, but rather, requires aninput of energy to proceed.

5.2 ENERGY TRANSFORMATIONSAND COUPLED REACTIONS

5.2.1 FREE ENERGY OF ATP ISASSOCIATED WITH COUPLEDPHOSPHATE TRANSFERREACTIONS

Biological organisms overcome metabolic restrictionsimposed by thermodynamically unfavorable reactionsthat exhibit a positive �G by coupling them or linkingthem to reactions with a negative �G. Thus, the freeenergy released by the exergonic reaction provides thenecessary energy to ensure that the endergonic reactionproceeds. However, in order that this system of coupledreactions will occur spontaneously, the net free energychange must always be negative. Thus, the concept of suchcoupled reactions was critical to our understanding ofhow life, an overall endergonic process, is maintainedwithout defying the laws of thermodynamics. Thenotion of biological coupled reactions was unknownto Schrodinger in 1944 to explain the complex events

82 Chapter 5 / Bioenergetics and ATP Synthesis

that take place within ‘‘the spatial boundary of a livingorganism’’ through space and time. For example, in gly-colysis, the hydrolysis of adenosine triphosphate (ATP)(Equation 5.11), which is an exergonic reaction, is cou-pled to the phosphorylation of glucose (Equation 5.12),which is an endergonic reaction. The overall netreaction proceeds because the net reaction is exergonic(Equation 5.13).

ATP → ADP + Pi �G◦′ = −32.2 kJ/mol (5.11)

Glucose + Pi →Glucose-6-P�G◦′ = +13.8 kJ/mol (5.12)

ATP + Glucose →Glucose-6-P + ADP�G◦′ = −18.4 kJ/mol (5.13)

When ATP was first isolated from extracts of musclein 1929, F. Lipmann was one of the first to recognizeits significant role in the energy metabolism of the cell(Figure 5.2). He called it a high-energy moleculeand designated the terminal phosphate bonds by asquiggle symbol (∼). Actually, the designation of ATPas a high-energy molecule is somewhat misleadingbecause it implies that ATP is in some way a uniquemolecule. It is not. The two terminal phosphate bondsof the ATP molecule are normal, covalent anhydridebonds. Hydrolysis of an anhydride is accompanied by afavorable increase in entropy, largely due to resonancestabilization of the product molecules: the electronshave one additional bond through which they canresonate. In addition, both the ADP and Pi products are

acidic anions and the two negatively charged products,because of mutual charge repulsion, do not readilyrecombine. Consequently, the equilibrium constant(Keq) for ATP hydrolysis is rather large—on the orderof 105. Thus, for ATP hydrolysis

Keq = [ADP][Pi]/[ATP] ≈ 105 (5.14)

Such a large equilibrium constant helps to explainwhy ATP is so important in cellular metabolism.Photosynthesis and respiration serve to maintain alarge pool of ATP such that the observed mass-actionratio (�) can be maintained as low as 10−4. Thisis nine orders of magnitude away from equilibrium(Equation 5.14). By substituting these values forKeq and A in Equation 5.10, we can see that acell generates considerable free energy (�G ≈−56 kJ/mol) simply through its capacity to maintainhigh concentrations of ATP. Thus, it is the cell’scapacity to maintain the mass-action ratio so far fromequilibrium that enables ATP to function as an energystore.

It is important to emphasize that it is the extentof displacement of G from equilibrium that definesthe capacity of a reactant to do work, rather than anyintrinsic property of the molecule itself. In the words ofD. G. Nicholls, if the glucose-6-phosphate reac-tion (�G◦′ = 13.8 kJ/mol) ‘‘were maintained tenorders of magnitude away from equilibrium, thenglucose-6-phosphate would be just as capable of doingwork in the cell as is ATP. Conversely, the Pacific

O OP P PO OHN

N CC

C

N

NH2

CHHC

N N

H

OH

H H

OH

C

C

H

C

C

H O O O

H

C

OOH OH OH

O OP P PO O–C

H O O O

OHH OH OH

OC P PO O–

H O O

H OH

HO P O–

O

OHOH

Phosphate [�3]

Ribose

Adenine

+

HO–

A.

B.

FIGURE 5.2 Adenosine triphosphate (ATP).(A) The ATP molecule consists of ade-nine (a nitrogenous base), ribose (a sugar),and three terminal phosphate groups. (B)Hydrolysis of ATP yields ADP (adenosinediphosphate) plus an inorganic phosphatemolecule.

5.2 Energy Transformations and Coupled Reactions 83

Ocean could be filled with an equilibrium mixture ofATP, ADP and Pi but the ATP would have no capacityto do work’’ (Nicholls and Ferguson, 2002). Of course,it goes without saying that the biochemistry of the cell isstructured so as to use ATP, not glucose-6-phosphate,in this capacity.

The relationship between free energy and the capac-ity to do work is not restricted to chemical reactions.Any system not at equilibrium has a capacity to do work,such as an unequal distribution of solute moleculesacross a membrane (see Chapter 3). The maintenanceof such a steady-state but nonequilibrium condition iscalled homeostasis. Indeed, homeostasis that reflectsthe capacity to avoid equilibrium in spite of changingenvironmental conditions is an essential characteristicof all living organisms. When �G = 0, no useful workcan be done and life ceases to exist.

5.2.2 FREE ENERGY CHANGES AREASSOCIATED WITH COUPLEDOXIDATION–REDUCTIONREACTIONS

Photosynthesis and respiration, which we will discussin more detail in Chapters 7 and 10, are electro-chemical phenomena. Each operates as a sequence ofoxidation–reduction reactions in which electrons aretransferred from one component to another. Thus, theoxidation of one component is linked or coupled to thereduction of the next component. Such coupled electrontransfer reactions are known as reduction–oxidation orredox reactions. This can be illustrated in the followinggeneral way. Any compound, A, in its reduced form(Ared), becomes oxidized (Aox) when it gives up an elec-tron (e−) (Equation 5.15). Similarly, any compound, B,in its oxidized form (Box), becomes reduced (Bred) whenit accepts an electron (Equation 5.16). When thesetwo reactions are coupled, the net effect is the trans-fer of an electron from compound A to compound B(Equation 5.17) with Ared being the reductant and Boxbeing the oxidant.

Ared → Aox + e− (5.15)

Box + e− → Bred (5.16)

Ared + Box → Aox + Bred (5.17)

As a specific biological example, consider the netreduction of three-phosphoglyceric acid (PGA) toglyceraldehyde-3-P (GAP) by the reduced formof nicotinamide adenine dinucleotide phosphate(NADPH):

PGA + NADPH + H+ � GAP + NADP+ (5.18)

Redox reactions may be conveniently dissected into twohalf-reactions involving the donation and acceptance ofelectrons. Thus the reduction of PGA to GAP may be

considered as the two half-reactions:NADPH � NADP+ + H+ + 2e− (5.19)

PGA + 2e− + H+ � GAP (5.20)Thus, the oxidation of NADPH to NADP+ is coupled tothe reduction of PGA to GAP. A reduced/oxidized pairsuch as NADPH/NADP+ is known as a redox couple.Note that oxidation–reduction reactions often involvethe transfer of protons. The positively charged protonsbalance the negative charge of the acquired electrons,which maintains electroneutrality. The involvement ofprotons indicates that the redox reaction is pH sensi-tive! The structures of some principal redox compoundsinvolved in biological electron transport are shown inFigure 5.3.

Since each of the half-reactions described above, aswell as the net reaction, is reversible, their free energiescould be described on the basis of chemical equilibria.However, it is not clear how to treat the electrons,which have no independent existence. Moreover, ourinterest in redox couples is more in their tendency toaccept electrons from or donate electrons to anothercouple, a tendency known as redox potential. Redoxpotentials allow the feasibility and direction of electrontransfers between components in a complex system tobe predicted. Indeed, in order to understand electronflow in photosynthesis and respiration it is necessary tohave a working understanding of redox potential andhow it is applied.

The direction of electron transfer between redoxcouples can be predicted by comparing their midpointpotentials (Em). Thermodynamically spontaneous elec-tron transfer will proceed from couples with the morenegative (less positive) redox potential to those withthe less negative (more positive) redox potential. Theenergy-transducing membranes of bacteria, mitochon-dria, and chloroplasts all contain electron-transportsystems involving a number of electron carriers withdifferent midpoint redox potentials (Table 5.1).

TABLE 5.1 Midpoint redox potentials for aselection of redox couples involved inphotosynthesis and respiration.

Reductant/Oxidant Em (mV)

Ferredoxin red/ox −430H2/2H+ −420NADH + H+/NAD+ −320NADPH + H+/NADP+ −320Succinate/fumarate +30Ubiquinone red/ox +40Cyt c2+/Cyt c3+ +2202H2O/O2 + 4H+ +820

CH

C OHH

C OHH

C OHH

CH O P

O– –

O

O P

O

O

O

N

NO

N

NH

O

H

2

C

H C

H C

H C

Flavin Adenine Dinucleotide (FAD)Flavin Mononucleotide (FMN)

Isoalloxazine

FAD

FMN

ON Nicotinamide

Ribose

HC

O

NH2

2

2

2 2

2

2

2

+

NH

N

N

N

N

H

CH O

HO OH

H HH

O

P OO

P OO

OAdenine

Ribose

R

Nicotinamide-Adenine Dinucleotide (NAD); R HNicotinamide-Adenine Dinucleotide Phosphate (NADP); R PO 3

H

CH O

HO O

H HH

Ubiquinone

Plastoquinone (PQ)(quinone form)

[CHO

CH

H

C

CH

CH ]n H

O

Plastoquinol (PQH )(hydroquinone form)

Ubiquinol

Oxidized Form Reduced FormA.

B.

C.

NH

N

N

N

N

H

CH O

HO OH

H HH

ON

HHC

O

NH–

NH

N

N

N

N

H

CH O

HO OH

H HH

O

P OO

P OO

O

R

H

CH O

HO O

H HH

+ [H ]

++ 2e + 2H

CH

C OH

H

H

C OHH

H

C OHH

CH O P

O

O

O P

O

O

O

N

NO

N

NH

O

H C

H C

NH

N

N

N

N

H

CH O

HO OH

H HH

2e + 2H

O

3

H C

H C [CH CH

H

C

CH

CH ]n HOH

OH

OH

OH

H3CO

H3CO [CH

H

+

2e + 2H+

–

–

–

–

CH C

CH

CH ]n H

H CO

H CO [CH

HO

CH C

CH

CH ]n H

2

2

22

2

3

3

3

3

2

2

2

2

––

–

–

+

2 2 2 2

2

2 2

3

3

3 3

3

3

–

+

2

3

3

3

–

FIGURE 5.3 The chemical structures of some common biological redox agents in oxidized and reduced states. (A) Nicoti-namide adenine dinucleotide (NAD) and nicotinamide adenine dinucleotide phosphate (NADP). Note that only thenicotinamide ring is changed by the reaction. The nicotinamide ring accepts two electrons but only one proton. Arrowindicates where the electrons are added to the nicotinamide ring. (B) Flavin adenine dinucleotide (FAD) consists ofadenosine (adenine plus ribose) and riboflavin (ribitol plus isoalloxazine). Flavin mononucleotide (FMN) consists ofriboflavin alone. Reduction occurs on the isoalloxazine moiety, which accepts two electrons and two protons. (C)Quinones. A quinone ring is attached to a hydrocarbon chain composed of five-carbon isoprene units. The value of nis usually 9 for plastoquinone, found in chloroplast thylakoid membranes, and 10 for ubiquinone, found in the innermembrane of mitochondria. Reduction of the quinone ring is a two-step reaction. The transfer of one electron producesthe partially reduced, negatively charged semiquinone (not shown). Addition of a second electron plus two protons yieldsthe fully reduced hydroquinone form.

84

5.3 Energy Transduction and the Chemiosmotic Synthesis of ATP 85

In addition to allowing us to predict the direction ofelectron transfer, redox potentials also permit the calcu-lation of Gibbs free energy changes for electron-transferreactions. This can be done using the following rela-tionship:

�G◦′ = −nF�Em (5.21)where n is the number of electrons transferred andF is the Faraday constant (96 500 coulombs mol−1).Biological electron transfers may involve either singleelectrons or pairs, but energy calculations are almostalways based on n = 2. �Em is the redox interval throughwhich the electrons are transferred and is determined as

�Em = Em(acceptor) − Em(donor) (5.22)Thus, for a coupled transfer of electrons from water(the donor) to NADP+ (the acceptor) as occurs in pho-tosynthetic electron transport in chloroplasts, �Em =(−320) − (+820) = −1140 mV = −1.14 V. Substitut-ing this in Equation 5.21, the value of �G◦′ for atwo-electron transfer from H2O to NADP+ is +220 kJmol−1. Note that the sign of �G◦′ is positive, indicatingthat this electron transfer will not occur spontaneously.In photosynthesis, light energy is used to drive thiscoupled endergonic reaction. In contrast, mitochondriatransfer electrons from NADH to O2. For this coupledelectron-transfer reaction, the value of � Em is +1.14 Vand consequently �G◦′ is −220 kJ mol−1, indicating thatthis electron transfer in mitochondria is exergonic andthus will occur spontaneously. It is important to notethat the molecular mechanisms by which these elec-trons are transferred through complex processes suchas photosynthesis, respiration, and nitrogen assimilationare not in the purview of bioenergetics. The details ofthe underlying mechanisms will be addressed later inChapters 7, 8, 10, and 11, respectively.

5.3 ENERGY TRANSDUCTIONAND THE CHEMIOSMOTICSYNTHESIS OF ATP

It has been known for many years that the threeprincipal energy-transducing membrane systems (inbacteria, chloroplasts, and mitochondria) were able tolink electron transport with the synthesis of ATP. Themechanism, however, was not understood until PeterMitchell proposed his chemiosmotic hypothesis in1961. Although not readily accepted by many bio-chemists in the beginning, Mitchell’s hypothesis is nowfirmly supported by experimental results. In honor ofhis pioneering work, Mitchell was awarded the Nobelprize for chemistry in 1978.

Mitchell’s hypothesis is based on two fundamentalrequirements. First, energy-transducing membranes areimpermeable to H+. Second, electron carriers are orga-nized asymmetrically in the membrane. The result is

that, in addition to transporting electrons, some carriersalso serve to translocate protons across the membraneagainst a proton gradient. The effect of these protonpumps is to conserve some of the free energy of electrontransport as an unequal or nonequilibrium distributionof protons, or �pH, across the membrane.

5.3.1 CHLOROPLASTS ANDMITOCHONDRIA EXHIBITSPECIFIC COMPARTMENTS

In plants, chloroplasts and mitochondria are the mainenergy-transducing organelles. The biochemical mech-anism by which ATP is synthesized is directly relatedto the specific compartments that exist in each of theseorganelles. The structure (Figure 5.4) and developmentor biogenesis of chloroplasts have been studied exten-sively (Box 5.1). The number of chloroplasts per cellvaries from species to species. For example, the unicel-lular green alga, Chlamydomonas reinhardtii, exhibits asingle chloroplast while a single mesophyll cell in theleaves of many terrestrial plants can exhibit in excess of200 chloroplasts.

Within a mature chloroplast, we recognize fourmajor structural regions or compartments: (1) a pairof outer limiting membranes, collectively known as theenvelope, (2) an unstructured background matrix orstroma, (3) a highly structured internal system of mem-branes, called thylakoids, and (4) the intrathylakoidspace, or lumen (Figure 5.7A). The envelope defines theouter limits of the organelle. These membranes are 5.0to 7.5 nm thick and are separated by a 10 nm intermem-brane space. Because the inner envelope membrane

G

S

S ECW

G

P

FIGURE 5.4 Electron micrograph of a mesophyll chloro-plast of maize (Zea mays). S, stroma; G, granum; P,peripheral reticulum; E, envelope membrane; CW, cellwall.

86 Chapter 5 / Bioenergetics and ATP Synthesis

G

S

SFCW

G

P

BOX 5.1PLASTIDBIOGENESIS

Plastids are a family of double membrane-bound, semi-autonomous organelles common to plant cells. Plastidsarise from small, vesicular organelles called proplastids(0.2 to 1.0 μm) (Figure 5.5), which are carried from onegeneration to the next through the embryo and main-tained in the undifferentiated state in the dividing cellsof plant meristems. Plastids are prokaryotic in originand arose by endosymbiosis. This theory states thatsometime during plant evolution, an ancestral eukary-ote host cell engulfed a photosynthetic bacterium whichestablished a stable association within the host. As aconsequence, plastids contain their own DNA in theform of double-stranded circular molecules very sim-ilar to prokaryotic DNA, and replicate by division ofexisting plastids. Thus, plastids are considered to besemiautonomous and are inherited maternally in most

Amyloplast

FruitsFloral petalsRoots

Chromoplast

Chloroplast

Proplastid

LeavesStems

LeavesStems

LeavesStems

Nonphotosyntheticstorage tissues

FIGURE 5.5 Plastids. A diagram illustrating some of theinterrelationships between various types of plastids.Colorless plastids, that is, plastids without pigments orany prominent internal structure, are called leucoplasts.Leucoplasts in nonphotosynthetic tissues are often thesites of starch accumulation, in which case they arecalled amyloplasts. In photosynthetic tissues, chloro-plasts may become amyloplasts by accumulating excessphotosynthetic product in the form of starch granules.When the starch is ultimately degraded, the plastid mayonce again resume its photosynthetic function. Chloro-plasts in maturing fruits and senescing leaves are com-monly converted to chromoplasts due to the simultane-ous loss of chlorophyll and accumulation of yellow orreddish-orange carotenoid pigments. The characteris-tic colors of tomato fruit and many autumn leaves aredue to such chromoplasts. Chromoplasts may also formin nonphotosynthetic tissues such as carrot roots. Thepigments in chromoplasts are often so concentrated thatthey form a crystalline deposit.

flowering plants but are paternally inherited in gym-nosperms. Several categories of plastids are found inplants and are named according to their color. Themost prominent plastids in the leaves of plants are thegreen photosynthetic chloroplasts, which synthesizechlorophyll. Leucoplasts are colorless plastids that syn-thesize volatile compounds called monoterpenes presentin essential oils. Amyloplasts are unpigmented plastidsspecialized for the biosynthesis and storage of starch andare prominent organelles in plant storage organs. Chro-moplasts are pigmented plastids that are responsible forthe colors for many fruit (tomatoes, apples, organges)and flowers (tulips, daffodils, marigolds). Their color is aconsequence of the particular combination of carotenesand xanthophylls which they accumulate.

During expansion in the dark, leaves are eitherwhite or pale yellow. Such leaves are called etiolatedleaves. In the development of etiolated leaves,proplastids are converted to etioplasts (Figure 5.6).The etioplast is not considered an intermediatestage in the normal development of a chloroplastbut rather a specialized plastid present in etiolatedleaves. Etioplasts lack chlorophyll but accumulate thecolorless chlorophyll precursor, protochlorophyllide.A prominent structural feature of etioplasts is ahighly ordered, paracrystalline structure called theprolamellar body (PLB). When exposed to light,the protochlorophyll is converted to chlorophyll and theprolamellar body undergoes a reorganization to formthe internal thylakoid membranes of the chloroplast.

The assembly of thylakoid membranes duringnormal chloroplast biogenesis is extremely complex.In angiosperms, chloroplast biogenesis is strictlylight-dependent but not in gymnosperms and somegreen algae. However, regardless of the species,chloroplast biogensis requires coordination betweengene expression in the nucleus as well as in theplastids. In addition, plastid protein synthesis has to becoordinated with cytosolic synthesis of nuclear encodedphotosynthetic proteins and their subsequent importfrom the cytosol to the chloroplast. For example, thegenes that encode light harvesting polypeptides ofphotosystem I and photosystem II (Chapter 7) areencoded by the nucleus and synthesized in the cytosolwhile proteins that form the reaction centers of thesesame photosystems are encoded by chloroplast DNAand synthesized in the stroma. Elucidation of the mech-anisms and regulation of protein import into cellularorganelles continues to be a major area of research. Theformation of thylakoid membranes during the greeningprocess occurs via the sequential appearance of PS I,followed by PS II, intersystem electron transport com-ponents, and lastly the assembly of the light harvestingcomplexes associated with PS I and PS II. Maximumrates of CO2 assimilation and O2 evolution occur oncethe biogenesis of the thylakoid membranes is complete.

5.3 Energy Transduction and the Chemiosmotic Synthesis of ATP 87

SL

PLBPT

Chloroplast

DARK

IM

Etioplast

Proplastid

LIGHT

LIGHT

GL

FIGURE 5.6 Chloroplast development inhigher plants. All chloroplasts are derivedfrom proplastids, which exist as a simple,double membrane vesicle with invagina-tions of the inner membrane (IM). Uponexposure to light, chlorophyll biosynthesisis initiated with the concomitant differen-tiation of the thylakoid membranes intogranal lamellae (GL; stacks) separatedby stromal lamellae (SL; unstacked). Ifseedlings are initially grown in the dark,the proplastid develops into an etioplast.The etioplast is characterized by the pres-ence of a prolamellar body (PLB) and pro-thylakoids (PT) extending out from theprolamellar bodies. Upon transfer of thedark grown etiolated seedlings to the light,the prolamellar body in the etioplast dis-perses and thylakoid membranes develop,producing a normal chloroplast.

In addition to the conversion of proplastids tochloroplasts, light also induces the conversion of etio-plasts into chloroplasts in etiolated plants. This processusually takes approximately 12 to 24 hours (Figure 5.5).Upon exposure to light, carotenoid biosynthesis is stim-ulated and chlorophyll biosynthesis is initiated by theconversion of protochlorophyllide to chlorophyllide a bythe enzyme NADPH:protochlorophyllide oxidore-ductase (POR), the major protein present in the PLB(Figure 5.6). Thus, exposure of etiolated plants to lightcauses their leaves to turn green in a process referredto as de-etiolation or greening. The photoreduction

of protochlorophyllide is associated with the disorgani-zation of the PLB, which leads to the production andassembly of normal, functional thylakoid membranes.

FURTHER READING

Baker, N. R., J. Barber. 1984. Chloroplast Biogenesis. Topics inPhotosynthesis: Vol. 5. Amsterdam: Elsevier Press.

Biswal, U. C., B. Biswal, M. K. Raval. 2004. ChloroplastBiogensis: From Plastid to Gerontoplast. Dordrecht: KluwerAcademic Publishers.

is selectively permeable, the envelope also serves toisolate the chloroplast and regulate the exchange ofmetabolites between the chloroplast and the cytosolthat surrounds it. Experiments with spinach chloroplastshave shown that the intermembrane space is freely acces-sible to metabolites in the cytoplasm. Thus it appears

that the outer envelope membrane offers little by wayof a permeability barrier. It is left to the inner enve-lope membrane to regulate the flow of molecular trafficbetween the chloroplast and cytoplasm.

The envelope encloses the stroma, a predomi-nantly protein solution. The stroma contains all of the

88 Chapter 5 / Bioenergetics and ATP Synthesis

Chloroplast

Proton Circuit

MitochondrionOuter envelope

Inner envelope

Granum

Lumen

Thylakoid

Stroma

Outer membrane

Intermembrane space

Crista

Inner membrane

Matrix

Matrix

Intermembrane spaceLumen

Stroma

H+

H+

ADP + Pi ATP

e–

A.

C.

B.

Thylakoid Crista

Redoxcarriers

FIGURE 5.7 Using various compartments within chloroplasts and mitochondria, elec-tron transport is coupled to ATP synthesis through the establishment of a protoncircuit. (A) A typical chloroplast illustrating the major compartments (stroma, thy-lakoid membrane, lumen) involved in the establishment of a proton circuit for thechemiosmotic synthesis of ATP in chloroplasts. (B) A typical mitochondrion illus-trating the major compartments (matrix, cristae, intermembrane space) involved inthe establishment of a proton circuit for the chemiosmotic synthesis of ATP in mito-chondria. (C) A model illustrating the similarity in the mechanism by which electrontransport is coupled to the establishment of a pH gradient, which is, in turn, con-sumed in the synthesis of ATP in both chloroplasts and mitochondria. However,the biochemical composition of the redox carriers involved in photosynthetic andrespiratory electron transport are quite different.

enzymes responsible for photosynthetic carbon reduc-tion, including ribulose-1,5-bisphosphate carboxy-lase/oxygenase, generally referred to by the acronymrubisco (Chapter 8). Rubisco, which accounts for fullyhalf of the total chloroplast protein, is no doubt theworld’s single most abundant protein. In addition torubisco and other enzymes involved in carbon reduc-tion, the stroma contains enzymes for a variety of othermetabolic pathways as well as DNA, RNA, and thenecessary machinery for transcription and translation.

Embedded within the stroma is a complex system ofmembranes often referred to as lamellae (Figure 5.7A).This system is composed of individual pairs of par-allel membranes that appear to be joined at the end,a configuration that in cross-section gives the mem-branes the appearance of a flattened sack, or thylakoid

(Gr., sacklike). In some regions adjacent thylakoidsappear to be closely appressed, giving rise to membranestacks known as grana. The thylakoids found withina region of membrane stacking are called granathylakoids. Some thylakoids, quite often every secondone, extend beyond the grana stacks into the stromaas single, nonappressed thylakoids. These stromathylakoids most often continue into another granastack, thus providing a network of interconnectionsbetween grana. While the organization of thylakoidsinto stacked and unstacked regions is typical, it is by nomeans universal. One particularly striking example is thechloroplast in cells that surround the vascular bundles inC4 photosynthetic plants and that have no grana stacks(Chapter 15). Here the thylakoids form long, unpairedarrays extending almost the entire diameter of the

5.3 Energy Transduction and the Chemiosmotic Synthesis of ATP 89

chloroplast. The thylakoid membranes contain thechlorophyll and carotenoid pigments and are the siteof the light-dependent, energy-conserving reactions ofphotosynthesis.

The interior space of the thylakoid is known as thelumen. The lumen is the site of water oxidation and,consequently, the source of oxygen evolved in photo-synthesis. Otherwise it functions primarily as a reservoirfor protons that are pumped across the thylakoid mem-brane during electron transport and that are used todrive ATP synthesis. It is generally assumed that thethylakoids represent a single, continuous network ofmembranes. This means, of course, that the lumen ofeach thylakoid seen in cross-section also represents buta small part of a single, continuous system.

Plant mitochondria are typically spherical or shortrods approximately 0.5 μm in diameter and up to 2 μm inlength. The number of mitochondria per cell is variablebut generally relates to the overall metabolic activity ofthe cell. In one study of rapidly growing sycamore cellsin culture, 250 mitochondria per cell was reported. Themitochondria accounted for about 0.7 percent of totalcell volume and contained 6 to 7 percent of the total cellprotein. In other metabolically more active cells, such assecretory and transfer cells, the number is even higherand may exceed a thousand per cell!

Mitochondria, like the other energy-transducingorganelle, the chloroplast, are organized into severalultrastructural compartments with distinct metabolic

functions: an outer membrane, an inner mem-brane, the intermembrane space, and the matrix(Figure 5.7B). The composition and properties ofthe two membranes are very different. The outermembrane, which has few enzymatic functions, is richin lipids and contains relatively little protein. The innermembrane, on the other hand, contains over 70 percentprotein on a dry-weight basis. The outer membrane isalso highly permeable to most metabolites. It containslarge channel-forming proteins, known as porins(Chapter 1), which allow essentially free passage ofmolecules and ions with a molecular mass of 10000 Daor less. The permeability of the inner membrane, likethat of the chloroplast, is far more selective—it isfreely permeable only to a few small molecules suchas water, O2, and CO2. Like the chloroplast thylakoidmembrane, the permeability of the mitochondrial innermembrane to protons is particularly low, a significantfactor with respect to its role in ATP synthesis.

The inner membrane of the mitochondrion isextensively infolded. These invaginations form a densenetwork of internal membranes called cristae. The typi-cal biology textbook picture of cristae is based on animalmitochondria where the infoldings are essentially lamel-lar and form platelike extensions into the matrix. Thecommon pattern in plant mitochondria is less regular,forming a system of tubes and sacs (Figure 5.8). Theunstructured interior of the mitochondrion, or matrix, isan aqueous phase consisting of 40 to 50 percent proteinby weight. Much of this protein is comprised of enzymes

Crista

InnermembraneOutermembrane

FIGURE 5.8 The mitochondrion. Left: An electron micrograph of a mitochondrionfrom a maize (Zea mays) leaf cell (×10,000). Right: A diagram of the mitochondrionshown in the electron micrograph, to illustrate the essential features. Note that thecristae are continuous with the inner membrane (arrows).

90 Chapter 5 / Bioenergetics and ATP Synthesis

involved in carbon metabolism, but the matrix also con-tains a mitochondrial genome, including DNA, RNA,ribosomes, and the necessary machinery for transcribinggenes and synthesizing protein.

5.3.2 CHLOROPLASTS ANDMITOCHONDRIA SYNTHESIZEATP BY CHEMIOSMOSIS

In chloroplasts the protons are pumped across the thy-lakoid membrane, from the stroma into the lumen. Thedifference in proton concentration (�pH) across themembrane may be quite large—as much as three orfour orders of magnitude. Since protons carry a positivecharge, a �pH also contributes to an electrical potentialgradient across the membrane. A trans-thylakoid �pHof 3.5, for example, establishes a potential difference of207 mV at 25◦C. Together the membrane potential dif-ference (��) and the proton gradient (�pH) constitutea proton motive force (pmf).

pmf = −59�pH + �� (5.23)

In order to pump protons into the lumen against aproton motive force of this magnitude, a large amountof energy is required. This energy is provided by thenegative �G generated by photosynthetic intersystemelectron transport.

The direction of the proton motive force also favorsthe return of protons to the stroma, but the low pro-ton conductance of the thylakoid membrane will notallow the protons to simply diffuse back. In fact, thereturn of protons to the stroma is restricted to highlyspecific, protein-lined channels that extend through themembrane and that are a part of the ATP synthesiz-ing enzyme, ATP synthase (Figure 5.9). This large(400 kDa) multisubunit complex, also known as cou-pling factor or CF0—CF1, consists of two multipeptidecomplexes.

A hydrophobic complex called CF0 is largelyembedded in the thylakoid membrane. Attached to theCF0 on the stroma side is a hydrophilic complex calledCF1. The CF1 complex contains the active site forATP synthesis, while the CF0 forms an H+ channelacross the membrane, channeling the energy of theelectrochemical proton gradient toward the active siteof the enzyme. When the electron-transport complexesand the ATP-synthesizing complex are both operating,a proton circuit is established in chloroplasts as wellas mitochondria (Figure 5.7C). In chloroplasts, thephotosynthetic electron-transport complex pumps theprotons from the stroma into the lumen and thusestablishes the proton gradient. At the same time, theATP synthase allows the protons to return to thestroma. Some of the free energy of electron transportis initially conserved in the proton gradient. As theenergy-rich proton gradient collapses through the

α α

αβ

β

β

γ

δ

ε

Lumen

Stroma

ADP + Pi

CF1

CF0

ATP

3H+

b

c

3H+

a

FIGURE 5.9 A model of the chloroplast ATP synthase.CF0 is an integral membrane protein that forms a pro-ton channel through the thylakoid membrane. CF1 isattached to the stromal side of CF0 and contains theactive site for ATP synthesis. CF1 consists of five differ-ent subunits with a stoichiometry of α3, β3, γ , δ, ε. CF0

consists of four different subunits with a proposed stoi-chiometry of a, b, b′, c10. The proton channel lies at theinterface between subunit a and the ring of c subunits.The pmf rotates the ring of c subunits, which causes theCF1 to rotate and convert ADP and Pi to ATP. Largearrow indicates the direction of rotation.

CF0-CF1 complex, that conserved energy is availableto drive the synthesis of ATP. The light-dependentsynthesis of ATP by chemiosmosis in the chloroplast iscalled photophosphorylation. In mitochondria, therespiratory electron-transport complex pumps protonsfrom the matrix to the intermembrane space (IMS) toestablish a proton gradient. The potential energy ofthis proton gradient is consumed by the mitochondrialATP synthase to synthesize ATP (Figure 5.7C). Thechemiosmotic synthesis of ATP by mitochondria iscalled oxidative phosphorylation.

An essential element of Mitchell’s chemiosmotichypothesis is the reversibility of the ATP synthase reac-tion. This means that under appropriate conditions theCF0-CF1 and other similar complexes can use the neg-ative free energy of ATP hydrolysis to establish a protongradient. For example, both the plasma membrane andtonoplast contain ATPase proton pumps, which pumpprotons out of the cell or the vacuole, as the case maybe (Chapter 3). The energy of ATP is thus conservedin the form of a proton gradient that may then be cou-pled to various forms of cellular work. ATPase protonpumps are a principal means of utilizing ATP to provide

Further Reading 91

energy for the transport of other ions and small solutemolecules across cellular membranes (Chapter 3).

Bioenergetics is a fundamental science. Its studyis challenging, but this discussion has, of necessity,been restricted to general principles. With this briefbackground in mind, however, we can now proceedto a discussion of energy conservation through thelight-dependent and light-independent reactions ofphotosynthesis and subsequently examine how plantcells unlock the stored chemical energy neededfor growth, development, and the maintenance ofhomeostasis through the processes of glycolysis andrespiration. In addition, we have seen that bioenergeticprinciples were important in our previous discussionof osmosis and water relations (Chapters 1 and 2) aswell as ion transport associated with nutrient uptake(Chapter 3).

SUMMARY

The application of thermodynamic laws to the study ofenergy flow through living organisms is called bio-energetics. There are two forms of energy, one thatis available to do work (free energy) and one that is not(entropy). In a biochemical system such as living orga-nisms, free energy is related to chemical equilibrium.The further a reaction is held away from equilibrium,the more work can be done. Cells utilize this principleto link or couple energy-yielding reactions withenergy-consuming reactions. In fact, life exists be-cause cells are able to avoid equilibrium, that is,avoid maximum entropy. Most energy-exchangereactions in the cell are mediated by phosphory-lated intermediates, especially ATP and relatedmolecules. ATP is useful in this regard becauseit has a large equilibrium constant and is highlymobile within the cell. Because ATP is turnedover rapidly, it is maintained far from equilibrium.By exploiting specific compartments in chloroplasts(stroma vs. lumen) and mitochondria (matrix vs.intermembrane space), the chemiosmotic synthesis of

ATP is linked or coupled to an energy-rich protongradient (a nonequilibrium proton distribution) acrossthe energy-transducing thylakoid of the chloroplastor the inner membrane (cristae) of the mitochondria.The free energy of electron transport is used toestablish the proton gradient and ATP is synthesizedas the protons return through transmembrane ATPsynthesizing complex.

CHAPTER REVIEW

1. The second law of thermodynamics states that thefree energy of the universe tends toward a mini-mum or that entropy tends toward a maximum.This idea is sometimes referred to as entropicdoom. Explain what is meant by entropic doom.

2. British writer C. P. Snow has written that under-standing the second law of thermodynamicsis as much a mark of the literate individual ashaving read a work of Shakespeare. Can youoffer an explanation of what he means by this?

3. Explain the relationship between free energy andchemical equilibria.

4. Explain the relationship between free energy andredox potential.

5. What is the major role of coupled reactions inbiology? Give an example of a coupled phosphatetransfer reaction and a coupled redox reaction.

6. Compare the compartmentation required for thechemiosmotic synthesis of ATP through pho-tophosphorylation and oxidative phosphorylation.

FURTHER READING

Nicholls, D. G. S., S. J. Ferguson. 2002. Bioenergetics 2. NewYork: Academic Press.

Schrodinger, E. 2000. What Is Life? Cambridge: CambridgeUniversity Press.

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Direction ofpropagation

E

E

H

H

1 wavelengthλ

6The Dual Role of Sunlight:

Energy and Information

Sunlight satisfies two very important needs of biologicalorganisms: energy and information. On the one hand,radiant energy from the sun maintains the planet’s sur-face temperature in a range suitable for life and, throughthe process of photosynthesis, is the ultimate source ofenergy that sustains most of life in our biosphere. Radia-tion, primarily in the form of light, also provides criticalinformation about the environment—information thatis used by plants to regulate movement, trigger devel-opmental events, and mark the passage of time. Theimportance of light in the life of green plants is reflectedin the study of photobiology, which encompasses notonly phenomena such as photosynthesis (which reflectsthe role of sunlight as an energy source) but alsophenomena such as photomorphogenesis and pho-toperiodism, where sunlight provides the necessaryinformation for proper plant development and the mea-surement of daylength, respectively.

In order to fully appreciate the pervasive importanceof light to plants, it is necessary to understand some-thing of the physical nature of light and the moleculeswith which light interacts in plants. In this chapter,we will

• explore the physical nature of light and how lightinteracts with matter,

• discuss some of the terminology used in describinglight and methods for measuring it,

• discuss briefly the characteristics of light in thenatural environment of plants, and

• review the principal pigments and pigment systemsfound in plants.

The various ways in which light is used by plants topower photosynthesis and regulate development will bediscussed throughout many of the subsequent chapters.

6.1 THE PHYSICAL NATUREOF LIGHT

6.1.1 LIGHT IS ELECTROMAGNETICENERGY, WHICH EXISTSIN TWO FORMS

What is light? As Johnson recognized more than 200years ago, ‘‘we all know what light is, but it is not easyto tell what it is.’’ The simplest answer is that lightis a form of radiant energy, a narrow band of energywithin the continuous electromagnetic spectrum

93

94 Chapter 6 / The Dual Role of Sunlight: Energy and Information

λ = 400 500 600 700 800 nm

Cosmic rays

Violet Blue Green Yellow Orange Red Infrared

10-14 10-12 10-10 10-8 10-6 10-4 10-2 1 102 104 106 cm

X rays

Gamma rays

Infrared

Solar rays

Visible light

Radio wavesUltra-violet Sound

FIGURE 6.1 The electromagnetic spectrum. Visible radiation, or light, representsonly a very small portion of the total electromagnetic spectrum.

of radiation emitted by the sun (Figure 6.1). The term‘‘light’’ describes that portion of the electromagneticspectrum that causes the physiological sensation ofvision in humans. In other words, light is defined by therange of wavelengths—between 400 and approximately700 nanometers—capable of stimulating the receptorslocated in the retina of the human eye. Strictly speaking,those regions of the spectrum we perceive as red, green,or blue are called light, whereas the ultraviolet andinfrared regions of the spectrum, which our eyes cannotdetect (although they may have significant biologicaleffects), are referred to as ultraviolet or infraredradiation, respectively. While the following discussionwill focus on light, it is understood that the principlesinvolved apply to radiant energy in the broader sense.

Like other forms of energy, light is a bit of an enigmaand is difficult to define. It is more easily described not bywhat it is but by how it interacts with matter. Physicistsof the late nineteenth and early twentieth centuriesresolved that light has attributes of both continuouswaves and discrete particles. Both of these attributes areimportant in understanding the biological role of light.

6.1.2 LIGHT CAN BE CHARACTERIZEDAS A WAVE PHENOMENON

The propagation of light through space is characterizedby regular and repetitive changes, or waves, in its electri-cal and magnetic properties. Electromagnetic radiationactually consists of two waves—one electrical and onemagnetic—that oscillate at 90◦ to each other and to thedirection of propagation (Figure 6.2). The wave proper-ties of light may be characterized by either wavelengthor frequency. The distance in space between wavecrests is known as the wavelength and is representedby the Greek letter lambda (λ). Biologists commonlyexpress wavelengths in units of nanometers (nm), where1 nm = 10−9 m. Frequency, represented by the Greekletter nu (ν), is the number of wave crests, or cycles,passing a point in space in one second. Frequency is thus

related to wavelength in the following way:

ν = c/λ (6.1)

where c is the speed of light (3 × 108 m s−1). Biologistsmost commonly use wavelength to describe light andother forms of radiation, although frequency is usefulin certain situations. Wavelengths of primary inter-est to photobiologists fall into three distinct ranges:ultraviolet, visible, and infrared (Table 6.1).

6.1.3 LIGHT CAN BE CHARACTERIZEDAS A STREAM OF DISCRETEPARTICLES

When light is emitted from a source or interacts withmatter, it behaves as though its energy is divided intodiscrete units or particles called photons. The energycarried by a photon is called a quantum (pl. quanta), toreflect the fact that the energy can be quantized, that is,it can be divided into multiple units.

The energy carried by a photon (Eq) is relatedto wavelength and frequency in accordance with thefollowing relationship:

Eq = hc/λ = hν (6.2)

Direction ofpropagation

E

E

H

H

1 wavelengthλ

FIGURE 6.2 Wave nature of light. Electric vectors (E) andmagnetic vectors (H) oscillate at 90◦ to each other.

6.1 The Physical Nature of Light 95

TABLE 6.1 Radiation of Principal Interest to Biologists.

Wavelength Average EnergyColor Range (nm) (kJ mol−1 photons)

Ultraviolet 100–400UV-C 100–280 471UV-B 280–320 399UV-A 320–400 332

Visible 400–740Violet 400–425 290Blue 425–490 274Green 490–550 230Yellow 550–585 212Orange 585–640 196Red 640–700 181Far-red 700–740 166

Infrared longer than 740 85

where h is a proportionality constant, called Planck’sconstant. The value of h is 6.62 × 10−34 J s photon−1.Accordingly, the quantum energy of radiation is inverselyproportional to its wavelength or directly proportional toits frequency. The symbol hν (pronounced ‘‘h nu’’) iscommonly used to represent a photon in figures anddiagrams.

Since both h and c are constants, the energy ofa photon is easily calculated for any wavelength ofinterest. The following example illustrates a calculationof the energy content of red light, with a representativewavelength of 660 nm (6.6 × 10−7 m).

Eq = (6.62 × 10−34J s photon−1)(3 × 108m s−1)/6.6 × 10−7m

(6.3)

Solving for Eq:

Eq = 3.01 × 10−19J photon−1 (6.4)For blue light, with a representative wavelength of435 nm (4.35 × 10−7 m),

Eq = (6.62 × 10−34J s photon−1)(3 × 108m s−1)/4.35 × 10−7m

(6.5)

Again, solving for Eq:

Eq = 4.56 × 10−19J photon−1 (6.6)As the above numbers indicate, the energy content

of a single photon is a very small number. However, theEinstein-Stark law of photochemical equivalence states thatone photon can interact with only one electron. Thus,in any irreversible photochemical reaction, the energyof one photon may be used to convert one molecule ofreactant A to one molecule of product B.

A + hν → B (6.7)Since one mole of any substance contains Avo-

gadro’s number (N) of molecules (N = 6.023 × 1023

molecules mol−1), to convert one mole of reactant A

to one mole of product B would require N number ofphotons. Thus, for practical purposes it is convenientto multiply the energy of a single photon by Avogadro’snumber, which gives the value of energy for a mole ofphotons. The energy carried by a mole of photons of redlight, for example, is 181,292 J mol−1, or 181 kJ mol−1

(Table 6.1). The energy carried by a mole of photons ofblue light is correspondingly 274 kJ mol−1. The conceptof a mole of photons is more useful than dealing withindividual photons. For example, as will become appar-ent in the following section, the law of photochemicalequivalence states that a mole of photons of a partic-ular wavelength would be required to excite a mole ofpigment molecules.

6.1.4 LIGHT ENERGY CAN INTERACTWITH MATTER

For light to be used by plants, it must first be absorbed.The absorption of light by any molecule is a photo-physical event involving internal electronic transitions(Figure 6.3A). The Gotthaus-Draper principle tells usthat only light that is absorbed can be active in a pho-tochemical process. In contrast to photophysical events,photochemistry refers to any chemical reaction whichutilizes absorbed light to convert reactants to products,that is, any light-dependent reaction (Equation 6.7).Therefore, any photobiological phenomenon requiresthe participation of a molecule that absorbs light. Sucha molecule may be defined as a pigment. Plants containa variety of pigments that are prominent visual featuresand important physiological components of virtuallyall plants. The characteristic green color of leaves, forexample, is due to a family of pigments known as thechlorophylls. Chlorophyll absorbs the light energy usedin photosynthesis. The pleasing colors of floral petals

96 Chapter 6 / The Dual Role of Sunlight: Energy and Information

λ

GROUND STATE

EN

ER

GY

LEVE

L

RelativeAbsorbance(nm)

400

500

600

700T1

F

P

2 1

S2

S1

S0

a. b.

λ

λ

FIGURE 6.3 The absorption light by a molecule. (A) Anenergy level diagram depicting the various possible tran-sitions when light is absorbed. A nonexcited moleculeis said to be in the ground state (S0). Upon absorptionof light of wavelength of either λ1 or λ2, a molecule canundergo an electronic transition (solid arrows) to a sin-glet excited state represented by either S1 or S2, respec-tively. Within each singlet excited state exist variousinternal energy states representing vibrational and rota-tional states (smaller horizontal lines). Dashed arrowsrepresent radiationless decay through which energy isgiven up primarily as heat. Fluorescence (F) is the emis-sion of light from the lowest excited singlet state. T1

represents the metastable excited triplet state. Energyfrom the triplet excited state may be lost by radiationlessdecay or by delayed emission of light known as phospho-rescence (P). The triplet state is sufficiently long-livedto allow for photochemical reactions to occur. (B) Anabsorption spectrum (solid line) is a graph in whichabsorbance is plotted as a function of wavelength. Peaksor absorption bands correspond to the principal excita-tion levels illustrated in the energy diagram. Also shownis a fluorescence emission spectrum (dashed line) thatcorresponds to the emission of the absorbed energy aslight from the lowest excited singlet state (S1).

are due to the anthocyanin pigments that serve to attractinsects as pollen vectors. Other pigments, such as phy-tochrome, are present in quantities too small to be visiblebut nonetheless serve important roles in plant morpho-genesis. These and other important plant pigments willbe described later in this chapter.

What actually happens when a pigment moleculeabsorbs light? Absorption of light by a pigment moleculeis a rapid, photophysical, electronic event, occurringwithin a femtosecond (fs = 10−15 s). In accordancewith the First Law of Thermodynamics (Chapter 5),the energy of the absorbed photon is transferred to anelectron in the pigment molecule during that extremelyshort period of time. The energy of the electron is thus

elevated from a low energy level, the ground state, to ahigher energy level known as the excited, or singlet,state. This change in energy level is illustrated graphi-cally in Figure 6.3. Like photons, the energy statesof electrons are also quantized, that is, an electron canexist in only one of a series of discrete energy levels.A photon can be absorbed only if its energy contentmatches the energy required to raise the energy of theelectron to one of the higher, allowable energy states.

In the same way that quanta cannot be sub-divided, electrons cannot be partially excited. Although,according to the Einstein-Stark law of photochemicalequivalence, a single photon can excite only one elec-tron, complex pigment molecules, such as chlorophyll,will have many different electrons, each of whichmay absorb a photon of a different energy level and,consequently, different wavelength. Moreover, eachsinglet excited state in which an electron may exist maybe subdivided into a variety of smaller but discreteinternal energy levels called vibrational and rotationallevels. This broadens even further the number ofphotons that may be absorbed (Figure 6.3). Pigmentmolecules such as chlorophyll, when exposed to whitelight, will thus exhibit many different excited states atone time.

An excited molecule has a very short lifetime (onthe order of a nanosecond, or 10−9 s) and, in the absenceof any chemical interaction with other molecules in itsenvironment, it must rid itself of any excess energy andreturn to the ground state. Dissipation of excess energymay be accomplished in several ways.

1. Thermal deactivation occurs when a moleculeloses excitation energy as heat (Figure 6.3A). Theelectron will very quickly drop or relax to the lowestexcited singlet state. The excess energy is givenoff as heat to its environment. If the electron thenreturns to the ground state, that energy will also bedissipated as heat.

2. Fluorescence is the emission of a photon of lightas an electron relaxes from the first singlet excitedto ground state. Since the rate of relaxation throughfluorescence is much slower than the rate of relax-ation through thermal deactivation, fluorescenceemission occurs only as a consequence of relaxationfrom the first excited singlet state (Figure 6.3A).Consequently, the emitted photon has a lowerenergy content, and, in accordance with Equation6.1, a longer wavelength, than the exciting photon.In the case of the photosynthetic pigment chloro-phyll, for example, peak fluorescent emission falls tothe long-wavelength side of the red absorption band(Figure 6.3B). This is true regardless of whether thepigment was excited with blue light (450 nm, 262 kJmol−1) or red light (660 nm, 181 kJ mol−1). Forpigments such as chlorophyll in solution, a return

6.1 The Physical Nature of Light 97

to the ground state by emission of red light is oftenthe only available option.

3. Energy may be transferred between pigmentmolecules by what is known as inductive reso-nance or radiationless transfer. Such transferswill occur with high efficiency, but require thatthe pigment molecules are very close togetherand that the fluorescent emission band of thedonor molecule overlaps the absorption band ofthe recipient. Inductive resonance accounts formuch of the transfer of energy between pigmentmolecules in the chloroplast (Chapter 7).

4. The molecule may revert to another type of excitedstate, called the triplet state (Figure 6.3A). The dif-ference between singlet and triplet states, related tothe spin of the valence electrons, is not so importanthere. It is sufficient to know that the triplet stateis more stable than the singlet state—it is consid-ered a metastable state. The longer lifetime of themetastable triplet state (on the order of 10−3 s) issufficient to allow for photochemical reactions tooccur. This could take the form of an oxidationreaction in which the energetic electron is actuallygiven up to an acceptor molecule. When this occurs,the pigment is said to be photooxidized and theacceptor molecule becomes reduced.

6.1.5 HOW DOES ONE ILLUSTRATETHE EFFICIENCY OF LIGHTABSORPTION AND ITSPHYSIOLOGICAL EFFECTS?

Figure 6.4B illustrates a graph whereby the absorptionof light by a pigment (in this case, chlorophyll) mea-sured as relative absorbance is plotted as a function ofwavelength. The resulting graph is known as an absorp-tion spectrum which emphasizes the correspondencebetween possible excitation states of the molecule andthe principal bands in the absorption spectrum. Theenergy level diagram in Figure 6.3B represents, in abroad sense, absorption of light by chlorophyll. In thiscase, λ1 would represent red light and λ2 would representblue light.

An absorption spectrum is in effect a probabilitystatement. The height of the absorption curve (or width,as presented in Figure 6.4B) at any given wavelengthreflects the probability by which light of that energylevel will be absorbed. More importantly, an absorp-tion spectrum is like a fingerprint of the molecule.Every light-absorbing molecule has a unique absorptionspectrum that is often a key to its identification. Forexample, there are several different variations of thegreen pigment chlorophyll. The pattern of the absorp-tion spectrum for each variation generally resembles thatshown in Figure 6.4B, yet each variation of chlorophyll

b.

a.

400 500 600 700

Wavelength (nm)

Rel

ativ

e R

ate

of P

hoto

sybt

hesi

s (A

)

Rel

ativ

e A

bsor

banc

e (B

)

FIGURE 6.4 A typical action spectrum for leaf photosyn-thesis (A) compared with the absorption spectrum (B) ofa pigment extract from a leaf containing primarily chloro-phyll. The action spectrum has peaks in the blue and redregions of the visible spectrum that correspond to theprincipal absorption peaks for the pigment extract.

differs with respect to the precise wavelengths at whichmaximum absorbance occurs.

Because light must first be absorbed in order tobe effective in a physiological process, it follows thatthere must be a pigment that absorbs the effectivelight. One of the first tasks facing a photobiologistwhen studying a light-dependent response is to iden-tify the responsible pigment. One important piece ofinformation is called an action spectrum. An actionspectrum is a graph that shows the effectiveness oflight in inducing a particular process plotted as afunction of wavelength. The underlying assumptionis that light most efficiently absorbed by the responsi-ble pigment will also be most effective in driving theresponse. In other words, the action spectrum for alight-dependent response should closely resemble theabsorption spectrum of the pigment or pigments thatabsorb the effective light. A comparison of an actionspectrum with the absorption spectra of suspected pig-ments can therefore provide useful clues to the identityof the pigment responsible for a photosensitive process.As an example, a typical action spectrum for photo-synthesis in a green plant is shown in Figure 6.4A. Itis compared with the absorption spectrum for a leafextract that contains primarily chlorophyll and somecarotenoid (Figure 6.4B). Note that the action spectrumhas pronounced peaks in the red and blue regions ofthe spectrum and that these action maxima correspondto the absorption maxima for chlorophyll. This is part

98 Chapter 6 / The Dual Role of Sunlight: Energy and Information

of the evidence that identifies a role for chlorophyll inphotosynthesis.

6.1.6 ACCURATE MEASUREMENTOF LIGHT IS IMPORTANT INPHOTOBIOLOGY

Given the manifold ways in which light can influencethe physiology and development of plants, it shouldnot be too surprising that proper measurement anddescription of light and light sources has become asignificant component of many laboratory and fieldstudies. Many experiments are now conducted in con-trolled environment-rooms or chambers that allow theresearcher to control light, temperature, and humidity.To permit others to interpret the experiments or repeatthem in their own laboratories, it is essential that lightsources and conditions be fully and accurately described.The spectral distribution of light emitted from fluores-cent lamps is very different from that emitted fromtungsten lamps or from natural skylight. Because ofthese and many other factors, each light source mayhave demonstrably different effects on plant develop-ment and behavior. Even natural light changes in qualityfrom dawn through midday to dusk, or between shadyand sunny habitats or cloudy and open skies. Under-standing photobiology thus requires an understandingof how light is measured and what those measurementsmean. Also required is a consistent terminology that isunderstood by everyone working in the field.

There are three parameters of primary concernwhen describing light. The first is light quantity—howmuch light has the plant received? The second is thecomposition of light with respect to wavelength, knownas light quality, spectral composition, or spectralenergy distribution (SED). The third factor is timing.What are the duration and periodicity of the lighttreatment?

The measure of light quantity most widely acceptedby plant photobiologists is based on the concept of flu-ence. Fluence is defined as the quantity of radiant energyfalling on a small sphere, divided by the cross-sectionof the sphere. Since light is a form of energy that canbe emitted or absorbed as discrete packets or photons,fluence can be expressed in terms of either the numberof photons or quanta (in moles, mol) or the amountof energy (in joules, J). Photon fluence (units = molm−2) refers to the total number of photons incident onthe sphere while energy fluence (units = J m−2) refersto the total amount of energy incident on the sphere.The corresponding rate terms are photon fluence rate(units = mol m−2 s−1) and energy fluence rate (units =J m−2 s−1 or W m−2). The term irradiance is frequentlyused interchangeably with energy fluence rate, althoughin principle the two are not equivalent. Irradiance refersto the flux of energy on a flat surface rather than a sphere.

Many instruments for measuring radiation actuallymeasure total energy, including energy outside the visi-ble portion of the spectrum, such as infrared, which isnot directly relevant to photobiological processes. Inorder to avoid such complications, instruments are nowcommercially available that are limited to that portionof the spectrum between 400 nm and 700 nm. Thisrange of light is broadly defined as photosyntheticallyactive radiation (PAR). Thus photon fluence rates,expressed as mol photons m−2 s−1 PAR, or energyfluence rates, expressed as Watts (W) m−2 PAR, arewidely accepted for routine laboratory work in plantphotobiology. The only serious limitation to PARmeasurements is that they exclude light in the 700to 750 nm range—light which, although inactive inhigher plant photosynthesis, plays a significant role inregulating plant development (Chapter 16).

The term ‘‘light quality’’ refers to spectral compo-sition and is usually defined by an emission or incidence

300 400 500

Incandescent

a.

600 700 800 900 1,000

Rel

ativ

e S

pect

ral I

nten

sity

Wavelength (nm)

Fluorescent

12,000

10,000

8,000

6,000

4,000

2,000

0.32 0.50

b.

0.79 1.26 2 3.16 μm

0.40 0.63 1 1.59 2.51 3.98 μm

Erg

s pe

r sq

. cm

. pe

r se

c.

Outside theatmosphere

After passing throughthe atmosphere

Ultra-violet

Visible Infrared

FIGURE 6.5 Spectral energy distribution of sunlight (A)compared with fluorescent and incandescent light (B).Note the difference in wavelength scale between A andB. About 20 percent of the incoming energy from thesun, particularly in the infrared region, is absorbed byatmospheric gases (primarily CO2 and H2O). The solarspectrum was drawn from measurements made in clearweather from an observatory in Australia by C. W. Allen.(From H. H. Lamb, Climate: Present, Past and Future,London: Methuen & Co., 1972.)

6.2 The Natural Radiation Environment 99

spectrum. SED is measured with a spectroradiometer,an instrument capable of measuring fluence rate overnarrow-wavelength bands. Depending on the instru-ment, either spectral photon fluence rate (units = molphotons m−2 s−1 nm−1) or spectral energy fluence rate(units = W m−2 nm−1) is plotted against wavelength.In practice, spectroradiometers are also equipped withflat surface detectors that measure spectral irradiance(W m−2 nm−1).

SED can vary depending on the nature of the lightsource and a number of other factors (Figure 6.5).The SED of natural sunlight, for example, can varydepending on the quality of the atmosphere, cloud cover,and the time of day (Figure 6.5A). The SED of artificialsources, such as incandescent and fluorescent lamps, aresignificantly different from natural light (Figure 6.5B).Fluorescent light has a relatively high emission in theblue but drops off sharply in the red. Incandescent light,on the other hand, contains relatively little blue lightbut high emissions in the far-red and infrared.

6.2 THE NATURAL RADIATIONENVIRONMENT

A relatively small proportion of the radiation origi-nating in the sun reaches the earth’s atmosphere andeven less actually reaches the surface (Figure 6.5A).However, both the quantity and spectral distributionof radiant energy that reaches (or fails to reach) earthmay have a significant impact on the physiology of theplant. As well, radiant energy is central to several prob-lems of more immediate and profound consequencesfor man.

Significant amounts of infrared radiation are absor-bed by the water vapor and carbon dioxide and othergases present in the earth’s atmosphere (Figure 6.5A),giving rise to a phenomenon known as the green-house effect (Figure 6.6). Although public awareness ofthe greenhouse effect has increased markedly in recentyears, it is not a phenomenon restricted to the latetwentieth century. Indeed, the greenhouse effect hasbeen with us since the beginnings of life on earth. With-out it, life as we know it would not be possible. Infraredradiation is of low frequency (or long wavelength) andtherefore low energy. Its principal effect is to increasevibrational activity in molecules—that is, heat. Absorp-tion of infrared by atmospheric water vapor and carbondioxide creates a ‘‘thermal blanket’’ that helps to pre-vent extreme variations in temperature such as occur onthe lunar surface, where these gases are absent. Simi-lar, although less extreme, temperature variations arecharacteristic of dry, desert regions on earth wherehigh daytime temperatures alternate with very coolnights.

(A)

(B)

(C)

Earth

FIGURE 6.6 Diagram representing the greenhouse effect.Radiation from the sun warms both the atmosphereand the earth. (A) The earth then reradiates infrared(heat) back into the atmosphere. (B) Here the infraredradiation is either reflected back to earth or absorbedby atmospheric gases, such as CO2, H2O vapor, andmethanol, thus preventing its escape. (C) Some of thetrapped infrared is reradiated back to earth, giving rise toincreased temperatures.

Public concern about the greenhouse effect arisesfrom evidence that, since the beginnings of the indus-trial revolution, our prodigious consumption of fossilfuels has contributed to a steady increase in atmosphericcarbon dioxide and other so-called ‘‘greenhouse gases.’’Many believe that continued release of carbon dioxidewill lead to greater heat retention in the atmosphere andglobal warming. This could result in partial melting ofpolar ice caps with extensive flooding of low-lying landareas and major shifts in plant biodiversity and agri-cultural productivity. A scenario commonly proposedis that higher carbon dioxide levels will stimulate pho-tosynthesis and increase the amount of plant materialon earth. However, this is an overly simplistic view ofthe effects of CO2 concentrations on photosynthesis.Increases in both global temperatures and global CO2concentrations can have negative effects on photosyn-thetic rates.

At the other end of the spectrum, ultraviolet radi-ation is characterized by short wavelength, high fre-quency, and high energy levels (Table 6.1). Absorptionof ultraviolet radiation creates highly reactive molecules,often causing the ejection of an electron, or ionization ofthe molecule. Such ionizations usually have deleteriouseffects on organisms. A principal action of UV-C (about254 nm), for example, is to induce thiamine dimers(hence, mutations) in deoxyribonucleic acid. In the nat-ural environment, UV-induced mutation is not normallya major problem because little far-ultraviolet radiationreaches the surface. Virtually all of the UV-C and most

100 Chapter 6 / The Dual Role of Sunlight: Energy and Information

of the UV-B is absorbed by ozone (O3) and aerosols (dis-persed particles of solids or liquids) in the stratosphere.If, however, the atmospheric ozone concentration wereto be lowered, there would be an increased potentialfor harmful effects to all organisms. In recent years,just such a depletion of the stratospheric ozone layer,leading to increases in UV-B radiation reaching theearth’s surface, has become a matter of some concern.Data compiled over the past two decades have revealedthat approximately one-half of the plant species stud-ied are adversely affected by elevated UV-B radiation.It is perhaps not surprising that plants most sensitiveto UV-B radiation are those native to lower elevationswhere UV-B fluxes are normally low.

With respect to both frequency and energy level,visible light falls between UV and infrared radiation.Absorption of visible light raises the energy level ofvalence electrons of the absorbing molecule and thus hasthe potential for initiating useful photochemical reac-tions. Moreover, the fluence rate and spectral quality ofvisible light are constantly changing, often predictably,throughout the day or season. These variations conveyinformation about the environment—information thatthe plant can use to its advantage.

The two most significant changes in visible light ona daily basis are seen in the fluence rate and in spectraldistribution. Typically at midday under full sun, thefluence rate approaches 2000 μmol m−2 s−1. At twilight,just before the sun sets below the horizon, the fluencerate will have dropped to the order of 10 μmol m−2 s−1

or less. During the period known as dusk, fluence ratefalls rapidly—by as much as one order of magnitudeevery 10 minutes.

Falling light levels at end of day are accompaniedby shifts in spectral quality (see Chapter 24). Normaldaylight consists of direct sunlight and diffuse sky-light. Diffuse skylight is enriched with blue wavelengthsbecause the shorter wavelengths are preferentially scat-tered by moisture droplets, dust, and other componentsof the atmosphere. Consequently, normal daylight isenriched with blue (hence, blue skies!). At twilight,often defined as a solar elevation of 10◦ or less fromthe horizon, a combination of scattering and refractionof the sun’s rays as they enter the earth’s atmosphereat a low angle enriches the light with longer red andfar-red wavelengths. This is because, at twilight, thepath traversed by sunlight through the atmosphere toan observer on earth may be up to 50 times longer thanit is when the sun is directly overhead. Much of theviolet and blue light is thus scattered out of the line ofsight, leaving predominantly the longer red and orangeto reach the observer.

Atmospheric factors, such as clouds and air pollu-tion, also influence the spectral distribution of sun-light. Cloud cover reduces irradiance and increasesthe proportion of scattered (i.e., blue) light. Airborne

pollutants will cause scattering, but will also absorb cer-tain wavelengths. Plants growing under a canopy mustcope with severe reduction in red and blue light asit is filtered through the chlorophyll-containing leavesabove, or with sunflecks—spots of direct sunlight thatsuddenly appear through an opening in the canopy.These sudden changes in irradiance may have a signif-icant impact on the photosynthetic capacity of a plant(Chapter 14).

It is clear that plants are exposed to an ever-changinglight environment. Many of these changes, such as cloudcover, are unpredictable but others such as daily changesin fluence rate and spectral energy distribution occurwith great regularity. The more regular changes conveyprecise information about the momentary status of theenvironment as well as impending changes (see Chapters13 and 14). It is perhaps not surprising that plantshave evolved sophisticated means for interpreting thisinformation as a matter of survival.

6.3 PHOTORECEPTORS ABSORBLIGHT FOR USE IN APHYSIOLOGICAL PROCESS

Photoreceptors are defined as pigment molecules thatprocess the energy and informational content of lightinto a form that can be used by the plant. A pigmentthat contains protein as an integral part of the moleculeis known as a chromoprotein. Thus, photorecep-tors typically are chromoproteins. The chromophore(Gr. phoros, bearing) is that portion of the chromo-protein molecule responsible for absorbing light and,hence, color. The protein portion of a chromopro-tein molecule is called the apoprotein. The completemolecule, or holochrome, consists of the chromophoreplus the protein. The principal photoreceptors found inplants are described here. Their roles in various phys-iological processes will be discussed in detail in laterchapters.

6.3.1 CHLOROPHYLLS ARE PRIMARILYRESPONSIBLE FOR HARVESTINGLIGHT ENERGY FORPHOTOSYNTHESIS

As noted earlier, chlorophyll is the pigment primar-ily responsible for harvesting light energy used inphotosynthesis. The chlorophyll molecule consists oftwo parts, a porphyrin head and a long hydrocarbon,or phytol tail (Figure 6.7). A porphyrin is a cyclictetrapyrrole, made up of four nitrogen-containing pyr-role rings arranged in a cyclic fashion. Porphyrinsare ubiquitous in living organisms and include theheme group found in mammalian hemoglobin and the

6.3 Photoreceptors Absorb Light for Use in a Physiological Process 101

H

CH3

C2H5

MgNN

N N

CH3HCH

CH2

H3C

CHO

HH3C

H

H2C H

COOCH3

OH

IV III

I II

O

CH

CH2

C O

H3C C

H3C HC

7

8

H2C

H2C

H2C

CH2

CH2

CH2

CH2

CH2

CH2

CH3

CH2

CHO

O

H3C HC

H3C HC

FIGURE 6.7 Chemical structure of chlorophyll a. Chloro-phyll b is similar except that a formyl group replacesthe methyl group on ring II. Chlorophyll c is similar tochlorophyll a except that it lacks the long hydrocarbontail. Chlorophyll d is similar to chlorophyll a except thata —O—CHO group is substituted on ring I as shown.

photosynthetic and respiratory pigments, cytochromes(Chapters 7, 10). Esterified to ring IV of the porphyrinin chlorophyll is a 20-carbon alcohol, phytol. This long,lipid-soluble hydrocarbon tail is a derivative of the5-carbon isoprene. Isoprene is the precursor to a varietyof important molecules, including other pigments (thecarotenes), hormones (the gibberellins), and steroids(Chapter 19).

Completing the chlorophyll molecule is a magne-sium ion (Mg2+) chelated to the four nitrogen atoms inthe center of the ring. Loss of the magnesium ion fromchlorophyll results in the formation of a nongreen prod-uct, pheophytin. Pheophytin is readily formed duringextraction under acidic conditions, but small amountsare also found naturally in the chloroplast where it servesas an early electron acceptor (Chapter 7).

Four species of chlorophyll, designated chlorophylla, b, c, and d, are known. The chemical structure ofchlorophyll a, the primary photosynthetic pigment inall higher plants, algae, and the cyanobacteria, is shownin Figure 6.7. Chlorophyll b is similar except that aformyl group (—CHO) substitutes for the methyl groupon ring II. Chlorophyll b is found in virtually all higherplants and green algae, although viable mutants deficientof chlorophyll b are known. The principal differencebetween chlorophyll a and chlorophyll c (found in thediatoms, dinoflagellates, and brown algae) is that chloro-phyll c lacks the phytol tail. Finally, chlorophyll d, foundonly in the red algae, is similar to chlorophyll a exceptthat a (—O—CHO) group replaces the (—CH CH2)group on ring I.

When grown in the dark, angiosperm seedlingsdo not accumulate chlorophyll (Chapter 5, Box 1).Their yellow color is primarily due to the presence ofcarotenoids. Dark-grown seedlings do, however, accu-mulate significant amounts of protochlorophyll a, theimmediate precursor to chlorophyll a. The chemicalstructure of protochlorophyll differs from chlorophyllonly by the presence of a double bond between carbons 7and 8 in ring IV (Figure 6.7). The reduction of this bondis catalyzed by the enzyme NADPH:protochlorophylloxidoreductase. In angiosperms this reaction requireslight, but in gymnosperms and most algae chlorophyllcan be synthesized in the dark. There is a general consen-sus among investigators that chlorophyll b is synthesizedfrom chlorophyll a.

Note that the respective chlorophylls exhibit gen-erally a similar shape to their absorption spectra inorganic solvents, but exhibit absorption maxima at dis-tinctly different wavelengths, in both the blue and thered regions of the spectrum (Figure 6.8). These shiftsin the absorbance maxima illustrate that subtle chemicalchanges in the porphyrin ring of chlorophyll (Figure 6.7)have significant effects on the absorption properties ofthis pigment. This is evidence that it is the porphyrinring of chlorophyll that actually absorbs the light andnot the phytol tail. Note also that chlorophyll does notabsorb strongly in the green region of the visible lightspectrum (490–550 nm). The strong absorbance in theblue and red and transmittance in the green is what giveschlorophyll its characteristic green color.

The presence of the long hydrocarbon phytolexerts a dominant effect on the solubility of chlorophyll,rendering it virtually insoluble in water. In the plant,

102 Chapter 6 / The Dual Role of Sunlight: Energy and Information

300 400 500

Chl b

Chl a

600 700

Abs

orba

nce

Wavelength (nm)

FIGURE 6.8 Absorption spectra of chlorophyll a (brokenline) and chlorophyll b (solid line) in acetone.

chlorophyll is found exclusively in the lipid domainof the chloroplast membranes, where it formsnoncovalent associations with hydrophobic proteins.Only an extremely small percentage of the chlorophyllfound in vivo is ever free chlorophyll, that is, notbound to proteins. The absorption spectra of thesechlorophyll-protein complexes are markedly differentfrom that of free pigment in solution. For example,chlorophyll a-protein absorbs primarily in the regionof 675 nm as opposed to 663 nm for chlorophyll a inacetone. Conjugation of chlorophylls with protein in themembrane is important for three reasons. One is that ithelps to maintain the pigment molecules in the preciserelationship required for efficient absorption andenergy transfer. A second reason is that it provides eachpigment with a unique environment that in turn giveseach molecule a slightly different absorption maximum.These slight absorbance differences are an importantfactor in the orderly transfer of energy throughthe pigment bed toward the reaction center wherephotochemical conversion actually occurs (Chapter 7).Third, the presence of excess free chlorophyll wouldphotosensitize plants, which would lead to thedestruction of chloroplast structure. Chlorophyll inthe unbound state is less efficient in photosyntheticinductive energy transfer but reacts more efficiently

CH3

BH

NH

ONH

A

NH

CO2HCO2H

C

+ NH

D

CH2

O

FIGURE 6.9 The open-chain tetrapyrrole chro-mophore of phycocyanin. Compare with thecyclic tetrapyrrole group in the chlorophyllmolecule (Figure 6.7).

with O2 to generate to highly dangerous and reactiveoxygen species such as oxygen free radicals, hydroxylradicals, and singlet oxygen. These reactive oxygenspecies may destroy the chloroplast. Although lightenergy is essential for life, clearly it can be a verydangerous form of energy especially in an aerobicenvironment! Later we will examine the mechanismsthat photosynthetic organisms have evolved to protectthemselves against this potential danger (Chapter 14).

6.3.2 PHYCOBILINS SERVE ASACCESSORY LIGHT-HARVESTINGPIGMENTS IN RED ALGAE ANDCYANOBACTERIA

Phycobilins are straight-chain or open-chain tetrapyr-role pigment molecules present in the eukaryotic redalgae and the prokaryotic cyanobacteria (Figure 6.9).The prefix, phyco, designates pigments of algal origin.Four phycobilins are known. Three of these are involvedin photosynthesis and the fourth, phytochromobilin,is an important photoreceptor that regulates variousaspects of growth and development (Chapter 22).

The three photosynthetic phycobilins are phycoery-thrin (also known as phycoerythrobilin), phycocyanin(phycocyanobilin), and allophycocyanin (allophyco-cyanobilin). In addition to the open-chain tetrapyrrole,the phycobilin pigments differ from chlorophyllin that the tetrapyrrole group is covalently linkedwith a protein that forms a part of the molecule. Inthe cell, phycobiliproteins are organized into largemacromolecular complexes called phycobilisomes.

With the exception of phytochromobilin, phyco-bilin pigments are not found in higher plants butoccur exclusively in the cyanobacteria and the red algae(Rhodophyta) where they assume a light-harvestingfunction in photosynthesis. Phycobilins, and in par-ticular phycoerythrin, are useful as light harvesters forphotosynthesis because they absorb light energy in thegreen region of the visible spectrum where chloro-phyll does not absorb (Figure 6.10). The red algae,for example, appear almost black because the chloro-phyll and phycoerythrin together absorb almost all ofthe visible radiation for use in photosynthesis (compareFigure 6.10 with Figure 6.8).

6.3 Photoreceptors Absorb Light for Use in a Physiological Process 103

300 400 500 600 700

Rel

ativ

e A

bsor

banc

e

Wavelength (nm)

FIGURE 6.10 Absorption spectra of phycocyanin (solidline) and phycoerythrin (broken line) in dilute buffer.Compare with the absorption spectra of chlorophyll(Figure 6.8). Note that the phycobilins, phycoerythrinin particular, absorb strongly in the 500–600 nm rangewhere chlorophyll absorption is minimal.

The fourth phycobiliprotein, of particular signifi-cance to higher plants, is phytochrome, a receptor thatplays an important role in many photomorphogenicphenomena. Its chromophore structure and absorptionspectrum are similar to that of allophycocyanin. Phy-tochrome (literally, plant pigment) is unique because itexists in two forms that are photoreversible. The formP660 (or Pr) absorbs maximally at 660 nm. However,absorption of 660 nm light converts the pigment to asecond, far-red-absorbing form P735 (or Pfr). Absorp-tion of far-red light by Pfr converts it back to thered-absorbing form. Pfr is believed to be an activeform of the pigment that is capable of initiating a widerange of morphogenetic responses. Phytochrome willbe discussed in more detail in Chapter 22.

6.3.3 CAROTENOIDS ACCOUNTFOR THE AUTUMN COLORS

Carotenoids comprise a family of orange and yellowpigments present in most photosynthetic organisms.Found in large quantity in roots of carrot and tomatofruit, carotenoid pigments are also prominent in greenleaves. In the fall of the year, the chlorophyll pigmentsare degraded and the more stable carotenoid pigmentsaccount for the brilliant orange and yellow colors socharacteristic of autumn foliage.

Carotenoid pigments are C40 terpenoids biosyn-thetically derived from the isoprenoid pathway describedin Chapter 19. Because the carotenoids are predomi-nantly hydrocarbons, they are lipid soluble and foundeither in the chloroplast membranes or in specializedplastids called chromoplasts. The concentration of pig-ment in chromoplasts may reach very high levels, to the

extent that the pigment actually forms crystals. Thecarotenoid family of pigments includes carotenes andxanthophylls (Figure 6.11). Carotenes are predomi-nantly orange or red-orange pigments. β-carotene isthe major carotenoid in algae and higher plants. Notethat in β-carotene and α-carotene (a minor form), bothends of the molecule are cyclized. Other forms, such asγ -carotene, found in the green photosynthetic bacteria,have only one end cyclized. Lycopene, the principal pig-ment of tomato fruit, has both ends open. The yellowcarotenoids, xanthophylls, are oxygenated carotenes.Lutein and zeaxanthin, for example, are hydroxylatedforms of α-carotene and β-carotene, respectively.

Like chlorophyll, β-carotene in the chloroplast iscomplexed with protein. β-carotene, which absorbsstrongly in the blue region of the visible spectrum(Figure 6.12), is known to quench both the triplet excitedchlorophyll as well as the highly reactive singlet excitedoxygen, which can be generated by the reaction of tripletchlorophyll with ground state oxygen. Thus, β-caroteneprotects chlorophyll from photooxidation.

6.3.4 CRYPTOCHROME ANDPHOTOTROPIN AREPHOTORECEPTORS SENSITIVETO BLUE LIGHT AND UV-ARADIATION

A wide range of plant responses to blue light and UV-Aradiation have been known or suspected for a longtime. Cryptochrome was the name given initially tothe blue light/UV-A photoreceptor because blue lightresponses appeared to be prevalent in cryptograms, anold primary division of plants which do not exhibit trueflowers and seeds and included ferns, mosses, algae andfungi. In addition, the molecular nature of the blue lightphotoreceptor remained unknown, and thus, cryptic(secret or hidden) for many years. However, recentresearch has established that cryptochromes are foundthroughout the plant kingdom. The action spectrumfor cryptochrome exhibits two peaks, one in the UVAregion (320-400 nm) and one in the blue region ofthe visible spectrum (400-500 nm). The chromophorefor cryptochrome is a flavin. The three most commonflavins are riboflavin (Figure 6.13) and its two nucleotidederivatives, flavin mononucleotide (FMN) and flavinadenine dinucleotide (FAD). The flavins may occur freeor complexed with protein, in which case they are calledflavoprotein. However, those flavins that function asphotoreceptors probably constitute a very small portionof a much larger pool. Both FMN and FAD, for example,are important cofactors in cellular oxidation–reductionreactions (Chapter 5).

Arabidopsis has two genes for cryptochrome (CRY1and CRY 2) whereas tomato has at least three genesthat encode this photoreceptor. Mosses and ferns exhibit

Lutein

Violaxanthin

XANTHOPHYLLS

Zeaxanthin

CH

CHH3 3

3 3

3

C CH

HHO

CC

H H

H H H H H H HCH

CH3 CH3

33

3

H C CH

O

OH

H C

H

CC

H

CC

H

CC

H

CC

CC

CC

CC

CC

CH

CHH3 3

3 3

3

C CH

H

CC

H H

H H H H H H HCH

CH3 CH3

33

3

H C CH

H C

H

CC

H

CC

H

CC

H

CC

CC

CC

CC

CC

CH

CHH3 3

3 3

3

C CH

H

CC

H H

H H H H H H HCH

CH3 CH3

33

3

H C CH

H C

H

CC

H

CC

H

CC

H

CC

CC

CC

CC

CCO

CAROTENES

-Carotene

CH

CHH3 3

3 3

3

C CH

H

CC

H H

H H H H H H HCH

CH3 CH3

33

3

H C CH

-Carotene

Lycopene

C H

H C

H

CC

H

CC

H

CC

H

CC

CC

CC

CC

CC

CH

CHH3 3

3 3

3

C CH

H

CC

H H

H H H H H H HCH

CH3 CH3

33

3

H C CH

H C

H

CC

H

CC

H

CC

H

CC

CC

CC

CC

CC

CH

CHH3 3

3 3

3

C CH

H

CC

H H

H

HH H H H H HCH

CH3 CH3

33

3

H C CH

H C

H

CC

H

CC

H

CC

H

CC

CC

CC

CC

CC

�

�

C

HO

HO

OH

OH

FIGURE 6.11 The chemical structures of representative carotenes and xanthophylls. The principal distinction between the two is that xanthophylls contain oxygenand carotenes do not. Carotenes are generally orange while xanthophylls are yellow.

6.3 Photoreceptors Absorb Light for Use in a Physiological Process 105

300 400 500 600 700

Abs

orba

nce

Wavelength (nm)

FIGURE 6.12 Absorption spectra of α-carotene (solid line)and β-carotene (broken line).

two and five genes for cryptochrome, respectively.Interestingly, the sequence of the CRY1 protein issimilar to photolyase, a unique class of flavoproteinsthat use blue light to stimulate repair of UV-induceddamage to microbial DNA. Photolyases contain twochromophores; one a flavin (FAD) and one a pterin(Figure 6.13). Although the precise nature of the CRY1chromophores remains to be determined, it appears thatone is FAD and the second is likely to be a pterin. Cryp-tochromes are cytoplasmic proteins with a mass of about75 kDa and, together with phytochrome, mediate photo-morphogenic responses such as photoperiod-dependent

H3C N O

H3CNH

O

O

N

N

CH2

OHH C

OHH C

OHH C

CH2OH

N

NR

NH2

A B C

N

NH

A.

B.

FIGURE 6.13 The structure of riboflavin (A) and pterin(B). Note the similarity between the pterin structure andthe B and C rings of riboflavin. See Chapter 5 for thestructures of riboflavin derivatives, FMN and FAD.

control of flowering, stimulation of leaf expansion andthe inhibition of stem elongation.

Phototropins (PHOT) are a second class of bluelight photoreceptor that was first discovered in the late1980s. Arabidopsis exhibits two phototropin genes des-ignated PHOT1 and PHOT2. Like cryptochrome,phototropin is also a flavoprotein with two FMNmolecules as chromophores. The molecular mass of thisphotoreceptor is about 120 kDa and is localized tothe plasmamembrane. Phototropins are not involvedin photomorphogenic responses. Analyses of mutantsdeficient in either PHOT1 or PHOT2 indicate that thesetwo genes exhibit partial overlapping roles in the regu-lation of phototropism. Furthermore, phototropins playimportant roles in optimizing photosynthetic efficiencyof plants such as the regulation of stomatal openingfor CO2 gas exchange as well as chloroplast avoidancemovement to protect the photosynthetic apparatusfrom photoinhibition due to exposure to excess light(Chapter 13). The biochemical nature and physiologicalroles of these blue light/UV-A photoreceptors arediscussed in more detail in Chapter 22.

6.3.5 UV-B RADIATION MAY ACTAS A DEVELOPMENTAL SIGNAL

More recently, a small number of responses, such asanthocyanin synthesis in young milo seedlings (Sorghumvulgare) and suspension cultures of parsley or carrot cellshave been described with an action spectrum peak near290 nm and no action at wavelengths longer than about350 nm. These findings would seem to indicate thepresence of one or more UV-B (280–320 nm) receptorsin plants, although the nature of the photoreceptors hasyet to be identified with certainty.

The impact of ultraviolet radiation, especiallyUV-B, on plants is receiving increasing attentionbecause of concerns about the thinning of theatmospheric ozone layer. A reduction in the ozone layerresults in an increase in UV-B radiation, specificallybetween 290 and 314 nm, which can cause damageto nucleic acids, proteins, and the photosyntheticapparatus and lead to shorter plants and reducedbiomass. The UV-B receptor also appears to modulateresponses to phytochrome in some systems. It has yetto be identified.

6.3.6 FLAVONOIDS PROVIDE THEMYRIAD FLOWER COLORS ANDACT AS A NATURAL SUNSCREEN

Although the plant world is predominantly green, it isthe brilliant colors of floral petals, fruits, bracts, andoccasionally leaves that most attracts humans and a vari-ety of other animals to plants. These various shades ofscarlet, pink, purple, and blue are due to the presence of

106 Chapter 6 / The Dual Role of Sunlight: Energy and Information

C CC

FIGURE 6.14 Flavonoids are phenylpropane derivativeswith a basic C6—C3—C6 composition.

pigments known as anthocyanins. Anthocyanins belongto a larger group of compounds known as flavonoids.Other classes of flavonoids (e.g., chalcones and aurones)contribute to the yellow colors of some flowers. Yetothers (the flavones) are responsible for the whiteness offloral petals that, without them, might appear translu-cent. The flavonoids are readily isolated and, becauseof their brilliant colors, have been known since antiq-uity as a source of dyes. Consequently, the flavonoidshave been extensively studied since the beginnings ofmodern organic chemistry and their chemistry is wellknown. The biosynthesis of flavonoids is discussed inChapter 28.

Flavonoids are phenylpropane derivatives with abasic C6—C3—C6 composition (Figure 6.14). The moststrongly colored of the flavonoids are the anthocyani-dins and anthocyanins. Anthocyanins are the glycosidederivatives of anthocyanidins. Unlike chlorophyll, theanthocyanins are water-soluble pigments and are foundpredominantly in the vacuolar sap. They are readilyextracted into weakly acidic solution. The color ofanthocyanins is sensitive to pH: both anthocyanidins andanthocyanins are natural indicator dyes. For example,the color of cyanidin changes from red (acid) to violet(neutral) to blue (alkaline). The deep violet extract ofboiled red cabbage will turn a definitely unappetizingblue-green if boiled in alkaline water!

Anthocyanins in leaves such as Coleus and red-leaved cultivars of maple (Acer sps.) are found in thevacuoles of the epidermal cells, where they appearto mask the chlorophylls. However, the anthocyaninsabsorb strongly between 475 nm and 560 nm whiletransmitting both blue and red light. Consequently, thepresence of anthocyanins does not interfere with photo-synthesis in the chloroplasts of the underlying mesophyllcells.

Virtually all flavonoids absorb strongly in the UV-Bregion of the spectrum (Figure 6.15). Since these com-pounds also occur in leaves, one possible function of theflavonoids is thought to be protection of the underlyingleaf tissues from damage due to ultraviolet radiation.Thus, the accumulation of UV-B absorbing flavonoidsacts as a natural sunscreen for plants, green algae, andcyanobacteria. As flower pigments, the flavonoids attractinsect pollinators. Many insects can detect ultravioletlight and thus can perceive patterns contributed by thecolorless flavonoids as well as the colored patterns visibleto humans. The synthesis of anthocyanins is stimulatedby light, both UV and visible, as well as by nutrient stress

600500400300

Wavelength (nm)

Abs

orba

nce

FIGURE 6.15 Absorption spectrum of the anthocyanin,pelargonin.

(especially nitrogen and phosphorous deficiencies) andlow temperature.

At least one group of flavonoids, the isoflavonoids,have become known for their antimicrobial activities.Isoflavonoids are one of several classes of chemicals ofdiffering chemical structures, known as phytoalexins,that help to limit the spread of bacterial and fungalinfections in plants. Phytoalexins are generally absentor present in very low concentrations, but are rapidlysynthesized following invasion by bacterial and fungalpathogens. The details of phytoalexin metabolism arenot yet clear. Apparently a variety of small polysaccha-rides, glycoproteins and proteins of fungal or bacterialorigin, serve as elicitors that stimulate the plant tobegin synthesis of phytoalexins. Studies with soybeancells infected with the fungus Phytophthora indicate thatthe fungal elicitors trigger transcription of mRNA forenzymes involved in the synthesis of isoflavonoids. Theproduction of phytoalexins appears to be a commondefense mechanism. Isoflavonoids are the predominantphytoalexin in the family Leguminoseae, but otherfamilies, such as Solanaceae, appear to use terpenederivatives.

6.3.7 BETACYANINS AND BEETS

The prominent red pigments of beet root andBougainvillea flowers are not flavonoids (as was longbelieved), but a more complex group of glycosylatedcompounds known as betalains or betacyanins.Betacyanins and the related betaxanthins (yellow)are distinguished from anthocyanins by the fact thatthe molecules contain nitrogen. They appear to berestricted to a small group of closely related familiesin the order Chenopodiales, including the goosefoot,cactus, and portulaca families, which are not known toproduce anthocyanins.

Further Reading 107

SUMMARY

Sunlight provides plants with energy to drive photo-synthesis and critical information about the environ-ment. Light is a form of electromagnetic energy thathas attributes of continuous waves and discrete parti-cles. The energy of a particle of light (a quantum) isinversely proportional to its wavelength.

Light is absorbed by pigments, and pigments thatabsorb physiologically useful light are called photo-receptors. All pigments have a characteristic absorptionspectrum that describes the efficiency of light absorp-tion as a function of wavelength. Because only light thatis absorbed by pigments can be effective in a physiolog-ical or biochemical process, a comparison of absorptionspectra with the action spectrum for a process helpsto identify the responsible pigment. When light is ab-sorbed, the pigment becomes excited, or unstable. Theexcess energy must be dissipated as heat, reemitted aslight, or used in a photochemical reaction, thus allow-ing the pigment to return to its stable, ground state.

Regular and predictable changes in fluence rateand spectral energy distribution provide plants withinformation about the momentary status of their envi-ronment as well as impending changes. The biochem-ical characteristics of the principal plant pigments ofphysiological interest are described.

CHAPTER REVIEW

1. Although, as Samuel Johnson said, it is not easy totell what light is, what is it? Describe the variousparameters of light and how it can be measured.

2. Describe the relationship between an absorptionspectrum and an action spectrum. Of what signifi-cance is an action spectrum to the plant physiolo-gist?

3. When is a pigment a photoreceptor? Make alist of the major plant pigments and identifyone or more principal functions of each.

4. Chlorophylls and carotenoids are found predom-inantly in cellular membranes while anthocyaninsare located in vacuoles. What does this distributiontell you about the chemistry of these pigments?

5. Assume you are writing a paper in which you re-port the effects of artificial light on the growthand photosynthesis of plants. How would youdescribe the light environment so that a readercould attempt to repeat your experiments in his/her own laboratory?

6. Describe how light energy is absorbed and dissi-pated by a pigment.

FURTHER READING

Batschauer, A. (ed.). 2003. Photoreceptors and Light Signaling.Cambridge: Royal Society of Chemistry.

Bova, B. 2001. The Story of Light. Naperville IL: SourcebooksInc.

Briggs, W. L., J. L. Supdich (eds.). 2005. Handbook of Photo-sensory Receptors. Weinheim Germany: Wiley-VCH.

Clegg, B. 2001. Light Years. An Exploration of Mankind’sEnduring Fascination with Light. London: PiatkusPublishers Ltd.

Goodwin, T. W. (ed.). 1988. Plant Pigments. London andNew York: Academic Press.

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PS II Cytochromecomplex

2H+

2H+

NADP + NADPH

PS I

LHCII

LHCI

CP47CP43

D2QA

D1

P6 80

Pheo

OEC

Mn2+PC PC

P700

A

fd

2H2O

QB

PQH2PQH2

PQ

PQPool

PQ

O2 + 4H+

Cyt b6

Cyt fFe-S

H +

ATPADP+ Pi

STROMA SIDE

LUMEN SIDE

7Energy Conservation in Photosynthesis:

Harvesting Sunlight

Photosynthesis is the fundamental basis of competitivesuccess in green plants and the principal organ of photo-synthesis in higher plants is the leaf. From the delicate,pastel hues of early spring through the brilliant reds andoranges of autumn, foliage leaves are certainly one of thedominant features of terrestrial plants. The biologist’sinterest in leaves, however, goes far beyond their aes-thetic quality. Biologists are interested in the structureof organs and how those structures are adapted to carryout effectively certain physiological and biochemicalfunctions. Leaves provide an excellent demonstration ofthis structure–function relationship. While some leavesmay be modified for special purposes (for example,tendrils, spines, and floral parts), the primary functionof leaves remains photosynthesis. In order to absorblight efficiently, a typical leaf presents a large surfacearea at approximately right angles to the incoming sun-light. From this perspective, the leaf may be viewed asa photosynthetic machine—superbly engineered to carryout photosynthesis efficiently in an extremely hostileenvironment.

Photosynthesis occurs not only in eukaryotic organ-isms such as green plants and green algae but also inprokaryotic organisms such as cyanobacteria and certaingroups of bacteria. In higher plants and green algae

the reactions of photosynthesis occur in the chloroplast,which is, quite simply, an incredible thermodynamicmachine. The chloroplast traps the radiant energy ofsunlight and conserves some of it in a stable chemi-cal form. The reactions that accomplish these energytransformations are identified as the light-dependentreactions of photosynthesis. Energy generated by thelight-dependent reactions is subsequently used to reduceinorganic carbon dioxide to organic carbon in the formof sugars. Both the carbon and the energy conservedin those sugars are then used to build the order andstructure that distinguishes living organisms from theirinorganic surroundings.

The focus of this chapter is the organization ofleaves with respect to the exploitation of light asthe primary source of energy and its conversion tothe stable, chemical forms of ATP and NADPHby the chloroplast. We will discuss

• the structure of terrestrial plant leaves with respectto the interception of light,

• photosynthesis as the reduction of carbon dioxideto carbohydrate,

• the photosynthetic electron transport chain, itsorganization in the thylakoid membrane, and its

109

110 Chapter 7 / Energy Conservation in Photosynthesis: Harvesting Sunlight

role in generating reducing potential and ATP,and

• the use of herbicides that specifically interact withphotosynthetic electron transport.

7.1 LEAVES AREPHOTOSYNTHETICMACHINES THAT MAXIMIZETHE ABSORPTION OF LIGHT

The architecture of a typical higher plant leaf is par-ticularly well suited to absorb light. Its broad, laminarsurface serves to maximize interception of light. In addi-tion, the bifacial nature of the leaf allows it to collectincident light on the upper surface and diffuse (both scat-tered and reflected) light on the lower surface. Grossmorphology is not, however, the only factor enhancinginterception of light—internal cellular arrangementsalso play an important role.

The anatomy of a typical dicotyledonous meso-morphic leaf is shown in Figure 7.1A,C. The leaf is

Palisade mesophyll

Upper epidermis

Lower epidermis

Hair(trichome)

Spongymesophyll

Stoma

C.

Epidermis

Mesophyll cells

StomataVascularbundle

Epidermis

D.

FIGURE 7.1 The structure of leaves shown in cross-section. (A,C) A dicotyledonousleaf Acer sp. (B,D) A monocotyledonous leaf (Zea mays), showing a section betweentwo major veins. (A,B: From T. E. Weier et al., Botany, 6th ed. New York, Wiley,1982. Used by permission of authors.)

sheathed with an upper and lower epidermis. Theexposed surfaces of the epidermal cells are coated with acuticle. The photosynthetic tissues are located betweenthe two epidermal layers and are consequently identi-fied as mesophyll (meso, middle; phyll, leaf) tissues. Theupper photosynthetic tissue generally consists of one tothree layers of palisade mesophyll cells. Palisade cellsare elongated, cylindrical cells with the long axis per-pendicular to the surface of the leaf. Below is the spongymesophyll, so named because of the prominent air spacesbetween the cells. The shape of spongy mesophyll cellsis somewhat irregular but tends toward isodiametric.The plan of a monocotyledonous leaf is similar exceptthat it lacks the distinction between palisade and spongymesophyll (Figure 7.1B,D).

Palisade cells generally have larger numbers ofchloroplasts than spongy mesophyll cells. In leaves ofCamellia, for example, the chlorophyll concentrationof the palisade cells is 1.5 to 2.5 times that of the spongymesophyll cells. The higher number of chloroplasts inthe palisade cells no doubt reflects an adaptation tothe higher fluence rates for photosynthetically activelight generally incident on the upper surfaces of theleaf.

7.1 Leaves are Photosynthetic Machines that Maximize the Absorption of Light 111

In spite of the relatively large number of chloroplastsin the palisade layers of a dicotyledonous leaf, there is asignificant proportion of the cell volume that does notcontain chloroplasts. Because the absorbing pigmentsare confined to the chloroplast, a substantial amount oflight may thus pass through the first cell layer withoutbeing absorbed. This has been called the sieve effect.Multiple layers of photosynthetic cells is one way ofincreasing the probability that photons passing throughthe first layer of cells will be intercepted by successivelayers (Figure 7.2).

The impact of the sieve effect on the efficiency oflight absorption is to some extent balanced by factorsthat change the direction of the light path within theleaf. Light may first of all be reflected off the manysurfaces associated with leaf cells. Second, light that isnot reflected but passes between the aqueous volume ofmesophyll cells and the air spaces that surround them(especially in the spongy mesophyll) will be bent byrefraction. Third, light may be scattered when it strikesparticles or structures with diameters comparable to its

AC

B D

Upper epidermis

Palisade mesophyll

Spongy mesophyll

Lower epidermis

FIGURE 7.2 A simplified diagram illustrating how theoptical properties of leaves help to redistribute incom-ing light and maximize interception by chlorophyll. (A)Photon strikes a chloroplast and is absorbed by chloro-phyll. (B) The sieve effect—a photon passes through thefirst layer of mesophyll cells without being absorbed. Itmay be absorbed in the next layer of cells or pass throughthe leaf to be absorbed by another leaf below. (C) Theplanoconvex nature of epidermal cells creates a lens ef-fect, redirecting incoming light to chloroplasts along thelateral walls of the palisade cells. (D) The light-guideeffect. Because the refractive index of cells is greater thanthat of air, light reflected at the cell–air interfaces may bechanneled through the palisade layer(s) to the spongymesophyll below.

wavelengths. In the leaf cell, for example, both mito-chondria and the grana structures within chloroplastshave dimensions (500–1000 nm) similar to the wave-lengths active in photosynthesis. Both organelles willscatter light. These three factors—reflection, refrac-tion, and scattering—combine to increase the effectivepath length as light passes through the leaf. The longerlight path increases the probability that any given pho-ton will be absorbed by a chlorophyll molecule before itcan escape from the leaf (Figure 7.2).

Careful studies of the optical properties of leaveshave shown that, in spite of their scattering properties,palisade cells do not appear to absorb as much light asmight be expected. That is to say that the palisade cellshave a lower than expected efficiency of light attenua-tion. This is apparently because they also act to someextent as a light guide. Some of the incident light ischanneled through the intercellular spaces between thepalisade cells in much the same way that light is trans-mitted by an optical fiber (Figure 7.2). It is probablethat photosynthesis in the uppermost palisade layer isfrequently light saturated. Any excess light would bewasteful and could, in fact, give rise to photoinhibi-tion and other harmful effects that we will discuss inmore detail later in this chapter. Thus, the increasedtransmission of light to the lower cell layers resultingfrom both scattering and the light-guide effect would nodoubt be advantageous by contributing to a more effi-cient allocation of photosynthetic energy throughoutthe leaf.

Not all leaves are designed like the ‘‘typical’’dicotyledonous mesomorphic leaf described above.Leaves may be modified in many ways to fit particularenvironmental situations. Pine leaves (or needles), forexample, are more circular in cross-section. Theircapacity for light interception has been compromisedin favor of a reduced surface-to-volume ratio, amodification that helps to combat desiccation whenexposed to dry winter air. In other cases, such as dryland or desert species, the leaves are much thickerin order to provide for storage of water. In extremecases, such as the cacti, the leaves have been reduced tothorns and the stem has taken over the dual functionsof water storage and photosynthesis. These and othermodifications to leaf morphology will be discussedmore fully in Chapter 14 and 15.

Within the leaf mesophyll cells of plants, the chloro-plast is the organelle that transforms light energy intoATP and NADPH to convert CO2 to sugars. The struc-ture of a typical chloroplast was discussed in Chapter 5.ATP is synthesized by chemiosmosis, whereas NADPHis the product of coupled electron transfer reactions inthe chloroplast thylakoid membranes. The enzymaticreactions involved in the conversion of CO2 to sugarstakes place in the chloroplast stroma (Chapter 8).

112 Chapter 7 / Energy Conservation in Photosynthesis: Harvesting Sunlight

7.2 PHOTOSYNTHESIS IS ANOXIDATION–REDUCTIONPROCESS

Although it may not be obvious at first glance, photosyn-thesis is fundamentally an oxidation–reduction reaction.This can be seen by examining the summary equationfor photosynthesis:

6CO2 + 12H2O → C6H12O6 + 6O2 + 6H2O (7.1)

Here photosynthesis is shown as a reaction betweenCO2 and water to produce glucose, a six-carbon car-bohydrate or hexose. Although glucose is not the firstproduct of photosynthesis, it is a common form of accu-mulated carbohydrate and provides a convenient basisfor discussion. Note that equal molar quantities of CO2and O2 are consumed and evolved, respectively. This isconvenient for the experimenter since it means that pho-tosynthesis can be measured in the laboratory as eitherthe uptake of CO2 or the evolution of O2. However,it is important to note that the ratio of CO2 fixed/O2evolved = 1 only under conditions where photorespira-tion is suppressed. We will discuss photorespiration andits impact on photosynthesis in more detail in Chapter 8.For simplicity, we can reduce equation 7.1 to

CO2 + 2H2O → (CH2O) + O2 + H2O (7.2)

where the term (CH2O) represents the basic buildingblock of carbohydrate. Equation 7.2 can be interpretedas a simple redox reaction, that is, a reduction of CO2to carbohydrate, where H2O is the reductant and CO2is the oxidant. But it might also be interpreted as ahydration of carbon (e.g., carbohydrate), as it was inearly studies of photosynthesis. How do we know thatit is one and not the other? And why is it necessaryto write the equation with two molecules of water asreactant (and one as product) when one would appearto suffice? These questions can best be answered byreviewing some of the early studies on photosynthesis(see Box 7.1. Historical Perspective—The Discovery ofPhotosynthesis).

One of the earliest clues to the redox nature ofphotosynthesis was provided by studies of C. B. vanNiel in the 1920s. As a microbiologist, van Niel wasinterested in the photosynthetic sulfur bacteria thatuse hydrogen sulfide (H2S) as a reductant in place ofwater. Consequently, unlike algae and higher plants,the photosynthetic sulfur bacteria do not evolve oxygen.Instead, they deposit elemental sulfur according to thefollowing equation:

CO2 + 2H2S → (CH2O) + 2S + H2O (7.3)

The reaction in equation 7.3 can also be written as twopartial reactions:

2H2S → 4e− + 4H+ + 2S (7.4)

CO2 + 4e− + 4H+ → (CH2O) + H2O (7.5)

Equations 7.4 and 7.5 describe photosynthesis in thepurple sulfur bacteria as a straightforward oxidation–reduction reaction. C. B. van Niel adopted a com-parative biochemistry approach and argued that themechanisms for oxygenic (i.e., oxygen-evolving)photosynthesis in green plants and anoxygenic (i.e.,non-oxygen-evolving) photosynthesis in the sulfur bac-teria both followed the general plan:

2H2A + CO2 → 2A + (CH2O) + H2O (7.6)

In this equation, A can represent either oxygen or sulfur,depending on the type of photosynthetic organism.According to equation 7.6, the O2 released in oxygenicphotosynthesis would be derived from the reductant,water. Correct stoichiometry would therefore requirethe participation of four electrons and hence twomolecules of water.

A second important clue was provided by R. Hillwho, in 1939, was first to demonstrate the partialreactions of photosynthesis in isolated chloroplasts. InHill’s experiments with chloroplasts, artificial electronacceptors, such as ferricyanide, were used. Under theseconditions, no CO2 was consumed and no carbohydratewas produced, but light-driven reduction of the electronacceptors was accompanied by O2 evolution:

4Fe3+ + 2H2O → 4Fe2+ + O2 + 4H+ (7.7)

Hill’s experiments confirmed the redox nature of greenplant photosynthesis and added further support for theargument that water was the source of evolved oxy-gen. Direct evidence for the latter point was finallyprovided by S. Ruben and M. Kamen in the early1940s. Using either CO2 or H2O labeled with 18O, aheavy isotope of oxygen, they showed that the labelappeared in the evolved oxygen only when suppliedas water (H2

18O), not when supplied as C18O2. If theevolved O2 is derived from water, then two molecules ofwater must participate in the reduction of each moleculeof CO2.

Based on these results, photosynthesis can be viewedas a photochemical reduction of CO2. The energy of lightis used to generate strong reducing equivalents fromH2O—strong enough to reduce CO2 to carbohydrate.These reducing equivalents are in the form of reducedNADP+ (or, NADPH + H+). Additional energy forcarbon reduction is required in the form of ATP, whichis also generated at the expense of light. The principalfunction of the light-dependent reactions of photosynthesisis therefore to generate the NADPH and ATP required forcarbon reduction. This is accomplished through a series ofreactions that constitute the photosynthetic electrontransport chain.

7.2 Photosynthesis is an Oxidation–Reduction Process 113

BOX 7.1HISTORICALPERSPECTIVE—THEDISCOVERY OFPHOTOSYNTHESIS

Photosynthesis assumes a role of such dominant propor-tions in the organization and development of plants, notto mention feeding world populations, it is somewhatsurprising that so little was known about the processbefore the final decades of the eighteenth century. Thepractice of agriculture was already several thousandyears old and practical discussions of crop productionhad been written at least 2,000 years before. The originsof plant nutrition as a science can be traced as far backas Aristotle and other Greek philosophers, who taughtthat plants absorbed organic material directly from thesoil. This theory, known as the humus theory, prevailedin agricultural circles until the late nineteenth century,long after the principles of photosynthesis had beenestablished.

The first suggestions of photosynthesis appear inthe writings of Stephen Hales, an English clergymanand naturalist who is considered ‘‘the father of plantphysiology.’’ In 1727, Hales surmised that plants obtaina portion of their nutrition from the air and wondered,as well, whether light might also be involved. Hales’sinsights were remarkably prescient, contrary as theywere to the long-established humus theory. However,chemistry had yet to come of age as a science and Hales’sideas were not provable by experiment or by referenceto any well-established chemical laws.

Rabinowitch and Govindjee (1969) date the ‘‘dis-covery’’ of photosynthesis as 1776, the year JosephPriestly published his two-volume work entitled Exper-iments and Observations on Different Kinds of Air. Butas with many other phenomena in science, there wasno one moment of discovery. The story gradually fellinto place through the cumulative efforts of severalclergy, physicians, and chemists over a period of nearly75 years. J. Priestly (1733–1804) was an English minis-ter whose nonconformist views on religion and politicsled to his emigration to the United States in 1794. Hewas also a scientist engaged in pioneering experimentswith gases and is perhaps best known for his discov-ery of oxygen. Priestly’s experiments, begun in 1771and first published in 1772, led him to observe that air‘‘contaminated’’ by burning a candle could not supportthe life of a mouse. He then found that the air couldbe restored by plants—a sprig of mint was introducedinto the contaminated air and ‘‘after eight or nine days Ifound that a mouse lived perfectly in that part of the airin which the sprig of mint had grown’’ (Priestly, 1772).

Priestly failed to recognize the role of light in hisexperiments and it was perhaps serendipitous that hislaboratory was well enough lighted for the experimentsto have succeeded at all. In 1773, Priestly’s experimentscame to the attention of Jan Ingen-Housz (1730–1799),a physician to the court of Austrian Empress MariaTheresa. During a visit to London, Ingen-Housz heardPriestly’s experiments described by the President of theRoyal Society. He was intrigued by these experimentsand six years later returned to England to conduct exper-iments. In the course of a single summer, Ingen-Houszperformed and had published some 500 experiments onthe purification of air! He observed that plants couldpurify air within hours, not days as observed by Priestly,but only when the green parts of plants were exposedto sunlight. Together, Priestly and Ingen-Housz hadconfirmed Hales’s guesses made some 52 years earlier.

Although Priestly continued his experiments—in1781 he agreed with Ingen-Housz on the value oflight and green plant parts—neither Priestly norIngen-Housz recognized the role of ‘‘fixed air,’’ as CO2was known at the time. This was left to the Swiss pastorand librarian Jean Senebier (1742–1809). In 1782,Senebier published a three-volume treatise in which hedemonstrated that the purification of air by green plantsin the light was dependent on the presence of ‘‘fixed’’air. It is interesting to note that all three scientistshad emphasized the purification of air in relation toits capacity to support animal life—plant nutritionwas not a central theme. At the same time chemistsacross Europe, including Priestly in England, Scheelein Germany, and Lavoisier in France, were activelyinvestigating the chemical and physical properties ofgases. By 1785 Lavoisier had identified ‘‘fixed’’ air asCO2 and by 1796 Ingen-Housz had correctly deducedthat CO2 was the source of carbon for plants.

Another important component in the equation ofphotosynthesis was added by the work of a Genevachemist, N. T. de Saussure (1767–1845). It was de Saus-sure who first approached photosynthesis in a sound,quantitative fashion. In his book Recherches Chimiquessur la Vegetation (1804) he showed that the weight oforganic matter plus oxygen formed by photosynthesiswas substantially larger than the weight of CO2 con-sumed. He thus concluded that the additional weightwas provided by water as a reactant. The equation forphotosynthesis, using the new language of chemistryfounded by Lavoisier, could now be written:

CO2 + H2O → O2 + organic matter

Finally, it remained for a German surgeon, JuliusMayer (1814–1878), to clarify the energy relationshipsof photosynthesis. In 1845, he correctly deduced, forthe first time, that the energy used by plants and animals

114 Chapter 7 / Energy Conservation in Photosynthesis: Harvesting Sunlight

in their metabolism is derived from the energy of the sunand that it is transformed by photosynthesis from theradiant to the chemical form. Thus by the middle of thenineteenth century the general outline of photosynthesiswas complete. Despite the importance of the process,however, it would be almost another century beforethe structural and chemical details of photosynthesiswould yield to modern methods of microscopic andradiochemical analysis.

FURTHER READING

Priestly, J. 1772. Observations on different kinds of air.Philosophical Transactions of the Royal Society of London62:166–170.

Rabinowitch, E. & Govindjee. 1969. Photosynthesis. NewYork: Wiley.

7.3 PHOTOSYNTHETICELECTRON TRANSPORT

7.3.1 PHOTOSYSTEMS ARE MAJORCOMPONENTS OF THEPHOTOSYNTHETIC ELECTRONTRANSPORT CHAIN

The key to the photosynthetic electron transportchain is the presence of two large, multimolecular,pigment-protein complexes known as photosystem I(PSI) and photosystem II (PSII) (Figure 7.3) PSIconsists of 18 distinct subunits whereas PSII consists of31 individual subunits! These two photosystems operatein series linked by a third multiprotein aggregate calledthe cytochrome complex. Overall, the effect of thechain is to extract low-energy electrons from waterand, using light energy trapped by chlorophyll, raisethe energy level of those electrons to produce a strongreductant NADPH (see Box 7.2: The Case for TwoPhotosystems).

The composition, organization, and function ofthe photosynthetic electron transport chain have beenan area of active study and rapid progress in recentyears. This interest has led to the development of avariety of experimental methods for the study of PSI,PSII, and other large-membrane protein aggregates.Most significant among these are techniques for theremoval of the complexes from the thylakoid mem-branes by first solubilizing the membrane with a range

H20

1/202 + 2H+

Cyt PS I2e

hν

PS II

hν

NADPH + H+

NADP+

+ 2H+

FIGURE 7.3 A linear representation of the photosynthetic electron-transport chain. Asequential arrangement of the three multimolecular membrane complexes extractslow-energy electrons from water and, using light energy, produces a strong reduc-tant, NADPH + H+.

of detergents. The different photosystems or classes ofmolecular aggregates can then be separated from eachother by centrifugation. If the detergents and the con-ditions under which the treatments are carried out arecarefully selected, not only can complexes be isolated butalso individual complexes can be further subdivided intosmaller aggregates that retain varying parts of the overallactivity. These purified complexes or subunits may thenbe analyzed for their composition with respect to pig-ments, protein, or other components or assayed for theircapacity to carry out specific photochemical or electrontransport reactions. Most recently this approach has ledto the crystallization of PSII and PSI reaction centers.By exposing these crystals to X-rays and analyzing theresulting diffraction patterns, scientists have been ableto determine the precise three-dimensional location ofall the pigment molecules and redox components of PSIand PSII reaction centers. The Nobel Prize in chem-istry was awarded to Diesenhoffer, Michel, and Huberin 1987 for the first successful crystallization and X-raydiffraction of bacterial reaction centers. This representsthe second Nobel Prize given for research in photosyn-thesis. See Chapter 8 for research that led to the firstNobel Prize in photosynthesis.

Such fractionation studies have revealed thatPSI and PSII each contain several different proteinstogether with a collection of chlorophyll and carotenoidmolecules that absorb photons. The bulk of thechlorophyll in the photosystem functions as antennachlorophyll (Figure 7.4). The association of chlorophyll

7.3 Photosynthetic Electron Transport 115

hνhν

Antenna

ReactionCenter

ElectronAcceptor

Q−

e−

A+

A Q

ElectronDonor

FIGURE 7.4 A photosystem contains antenna and a reac-tion center. Antenna chlorophyll molecules absorb in-coming photons and transfer the excitation energy tothe reaction center where the photochemical oxida-tion–reduction reactions occur.

with specific proteins forms a number of differentchlorophyll-protein (CP) complexes, which can beseparated by gel electrophoresis and identified on thebasis of their molecular mass and absorption spectrum(Figure 7.5). The core antenna for photosystem II, forexample, consists of two chlorophyll-proteins (CP)known as CP43 and CP47 (Figure 7.6). These two CPcomplexes each contain 20 to 25 molecules of chloro-phyll a. The core antenna chlorophyll a absorb light butdo not participate directly in photochemical reactions.However, protein-bound antenna chlorophylls lie veryclose together such that excitation energy can easilypass between adjacent pigment molecules by inductiveresonance or radiationless energy transfer (Chapter 6).The energy of absorbed photons thus migrates throughthe antenna complex, passing from one chlorophyllmolecule to another until it eventually arrives at thereaction center (Figure 7.4).

Each reaction center consists of a unique chloro-phyll a molecule that is thought to be present as adimer. This reaction center chlorophyll plus associ-ated proteins and redox carriers are directly involvedin light-driven redox reactions. The reaction centerchlorophyll is, in effect, an energy sink—it is thelongest-wavelength, thus the lowest-energy-absorbingchlorophyll in the complex. Because the reaction centerchlorophyll a is the site of the primary photochemicalredox reaction, it is here that light energy is actu-ally converted to chemical energy. The reaction centerchlorophyll a of PSI and PSII are designated as P700and P680, respectively. These designations identify thereaction center chlorophyll a, or pigment (P), with anabsorbance maximum at either 700 nm (PSI) or 680 nm(PSII).

35

15

0.0 0.5 1.0

20

25

4

Absorbance (671 nm)

Rel

ativ

e m

obili

ty (

mm

)

5

6

7

2

1

3

30

BA

2

3

4

5

6

7

1

FIGURE 7.5 Separation of thylakoid chlorophyll-proteincomplexes by nondenaturing polyacrylamide gel elec-trophoresis. (A) In the presence of specific deter-gents, chlorophyll-protein complexes are removedfrom thylakoid membranes structurally and function-ally intact. These pigment-protein-detergent complexesare charged and thus will migrate when an electricfield is applied. The porous matrix through which theelectric field is applied is a polyacrylamide gel. Thus,the physical separation of protein complexes througha polyacrylamide gel matrix by applying an electricfield is called polyacrylamide gel electrophoresis. Sincethese protein complexes still have chlorophyll boundto them, you can watch the chlorophyll-protein com-plexes separate through the gel matrix according totheir molecular mass right in front of your eyes! Thelargest complexes remain at the top of the gel and thesmallest complexes near the bottom of the gel. Theillustration shows such an electrophoretic separationof pigment-protein complexes from thylakoid mem-branes of Arabidopsis thaliana with the arrow indicat-ing the direction of migration. Typically, seven ‘‘greenbands’’ can be resolved. Bands 1 through 6 are individ-ual chlorophyll-protein complexes associated with PSIand PSII. The band exhibiting the greatest migration(band 7) is free pigment. (B) The relative amount repre-sented by each green band can be quantified by scan-ning the gel in a spectrophotometer. The peak areasprovide an estimate of the relative abundance of eachchlorophyll-protein complex. Clearly, green bands 1,2, and 3 are the most abundant pigment-protein com-plexes in this particular thylakoid sample. Biochemicaland spectroscopic analyses of each pigment-protein com-plex indicates that band 1 contains chlorophyll a andrepresents light harvesting complex (LHCI) associatedwith the core antenna complex (CP1) of PSI. Band 2is CP1 without its associated LCHI. Band 3 containschlorophyll a as well as chlorophyll b and represents thetrimeric form of the light harvesting complex (LHCII)associated with PSII. Band 4 and 6 represent the dimericand monomeric forms of LHCII. Band 5 is a chlorophylla pigment-protein complex designated as CPa and con-tains the core antenna of PSII (CP47, CP43) associatedwith the PSII reaction center, P680.

116 Chapter 7 / Energy Conservation in Photosynthesis: Harvesting Sunlight

PS II Cytochromecomplex

2H+

2H+

NADP + NADPH

PS I

LHCII

LHCI

CP47CP43

D2QA

D1

P6 80

Pheo

OEC

Mn2+PC PC

P700

A

fd

2H2O

QB

PQH2PQH2

PQ

PQPool

PQ

O2 + 4H+

Cyt b6

Cyt fFe-S

H +

ATPADP+ Pi

STROMA SIDE

LUMEN SIDE

FIGURE 7.6 The organization of the photosynthetic electron transport system in thethylakoid membrane. See text for details.

Tightly associated with the reaction centers, P680and P700, are core antenna complexes. CP47 and CP43are the core antenna of PSII whereas CP1 is the corecomplex of PSI (Figure 7.6). Also shown in Figure 7.6 aretwo additional chlorophyll-protein complexes, depictedin close association with PSII and PSI—light-harvestingcomplex II (LHCII) and light-harvesting complex I(LHCI), respectively. LHCII is associated with PSIIand LHCI is associated with PSI. As their names imply,the light-harvesting complexes function as extendedantenna systems for harvesting additional light energy.LHCI and LHCII together contain as much as 70 per-cent of the total chloroplast pigment, including virtuallyall of the chlorophyll b. LHCI is relatively small, has achlorophyll a/b ratio of about 4/1, and appears rathertightly bound to the core photosystem. LHCII, on theother hand, contains 50 to 60 percent of the total chloro-phyll and, with a chlorophyll a/b ratio of about 1.2, mostof the chlorophyll b. LHCII also contains most of thexanthophyll. The function of the light-harvesting com-plexes and the core antenna are to absorb light andtransfer this energy to the reaction centers (Figure 7.4).

The principal advantage of associating a single reac-tion center with a large number of light harvesting andcore antenna chlorophyll molecules is to increase effi-ciency in the collection and utilization of light energy.Even in bright sunlight it is unlikely that an individ-ual chlorophyll molecule would be struck by a photonmore than a few times every second. Since events atthe reaction center occur within a microsecond timescale, any reaction center that depended on a singlemolecule of chlorophyll for its light energy would nodoubt lie idle much of the time. Thus, the advantageof a photosystem is that while the reaction center is

busy processing one photon, other photons are beingintercepted by the antenna molecules and funneled tothe reaction center. This increases the probability thatas soon as the reaction center is free, more excita-tion energy is immediately available. The efficiencyof energy transfer through the light harvesting com-plexes and the core antenna complexes to the reactionis very high—only about 10 percent of the energy islost. Thus, it is important to appreciate that LCHIand LHCII are not necessarily absolute requirementsfor photosynthetic electron transport under light satu-rated conditions, that is, under conditions when lightis not limiting. Rather, the light harvesting complexesenhance photosynthetic efficiency under low light, thatis, under conditions where light limits photosynthesis.In fact, photosynthetic organisms modulate the struc-ture and function of the light harvesting complexes inresponse to changes irradiance. This will be discussedin more detail in Chapter 14. In addition, LHCII hasan important role in the dynamic regulation of energydistribution between the photosystems which will bediscussed in more detail in Chapter 13.

A schematic of the photosynthetic electron trans-port chain depicting the arrangement of PSI, PSII, andthe cytochrome b6/f complex in the thylakoid mem-brane is presented in Figure 7.6. A fourth complex—theCF0-CF1 coupling factor or ATP synthase—is alsoshown. All four complexes are membrane-spanning,integral membrane proteins with a substantial portionof their structure buried in the hydrophobic lipid bilayer.Note that the orientation of the complex and their indi-vidual constituents is not random—specific polypeptideregions will be oriented toward the stroma or lumenrespectively. Such a vectorial arrangement of proteins

7.3 Photosynthetic Electron Transport 117

is characteristic of all energy-transducing membranes, ifnot all membrane proteins, and is an essential element oftheir capacity to conserve energy through chemiosmosis(Chapter 5). One particularly significant consequence ofthis arrangement is the directed movement of protonsbetween the stroma and the thylakoid lumen as shown inFigure 7.6. Although PQ reduction and its concomitantprotonation occurs on the stromal side of the thylakoidmembrane, the oxidation of PQH2 by the cytochromeb6/f complex (Cyt b6/f ) requires the diffusion of PQH2from the stromal side to the lumen side of the thylakoidmembrane. It is this arrangement that gives rise tothe proton gradient necessary for ATP synthesis. Thisaspect of the electron transport chain will be revisitedlater. Another consequence of the vectorial arrangementis that the oxidation of water and reduction of NADP+occur on opposite sides of the thylakoid membrane.Water is oxidized and protons accumulate on the lumenside of the membrane where they contribute to thegradient, which drives ATP synthesis. However, bothNADPH and ATP are produced in the stroma wherethey are used in the carbon reduction cycle (Chapter 8)or other chloroplast activities (Chapter 11).

7.3.2 PHOTOSYSTEM II OXIDIZESWATER TO PRODUCE OXYGEN

Electron transport actually begins with the arrival ofexcitation energy at the photosystem II reaction centerchlorophyll, P680, which is located near the lumenalside of the reaction center. As illustrated in Figure 7.7,this excitation energy is required to change the redoxpotential of P680 from +0.8 eV to about −0.4 eV forP680*, the excited form of P680. As a consequence ofthis initial endergonic excitation process, P680* canrapidly (within picoseconds, 10−12 s) transfer elec-trons exergonically to pheophytin (Pheo). Pheophytin,

considered the primary electron acceptor in PSII, isa form of chlorophyll a in which the magnesium ionhas been replaced by two hydrogens. Since this initialoxidation of P680 is light dependent, this is called aphotooxidation event, which results in the formationof P680+ and Pheo−, a charge separation. Note thatthe energy of one photon results in the release of oneelectron, which is consistent with the Einstein-Stokeslaw (Chapter 6). This charge separation effectively storeslight energy as redox potential energy and represents theactual conversion of light energy to chemical energy. Itis essential that this charge separation be stabilized bythe rapid movement of the electron from P680 at thelumen side of the PSII reaction center to an electronacceptor molecule localized at the stromal side of thePSII reaction center (see discussion below). If the elec-tron were permitted to recombine with P680+, therewould be no forward movement of electrons, the energywould be wasted, and ultimately, carbon could not bereduced.

The role of the reaction proteins, D1 and D2, isto bind and to orient specific redox carriers of the PSIIreaction center in such a way as to decrease the prob-ability of charge recombination between P680+ andPheo−. How does this happen? First, within picosec-onds, pheophytin passes one electron on to a quinoneelectron acceptor called QA, resulting in the formationof [P680+ Pheo Q−

A ]. Now the PSII reaction centeris considered to be ‘‘closed,’’ that is, it is unable toundergo another photo-oxidation event (Figure 7.8).On a slower time scale of microseconds, the electronis passed from QA to plastoquinone(PQ), resulting inthe formation of [P680+ Pheo QA]. PQ is a quinone(see Figure 5.3C) that binds transiently to a bindingsite (QB) that is on the stromal side of the D1 reac-tion center protein (Figure 7.6). The reduction of PQto plastoquinol (PQH2) decreases its affinity for the

Strong Reductant

High

Low

NADP

fdP700∗

P680∗

PheoPQ

Cytb6 /f

PC

P700

2 hν > 680nm

PS I

PS II

2 hν ≤ 680 nm

P680

H20

StrongOxidant

Red

ox P

oten

tial

(eV

)

Rel

ativ

e E

nerg

y Le

vel

Direction of Electron Flow

0.6

0.4

0.2

0

0.2

0.4

0.6

0.8

1.0

(−)

(+)

1/2O2 + 2H+

FIGURE 7.7 The Z-scheme for photosyn-thetic electron transport. The redox com-ponents are arranged according to theirapproximate midpoint redox potentials (Em,Chapter 2).The vertical direction indicatesa change in energy level (�G, Chapter 5).The horizontal direction indicates elec-tron flow. The net effect of the processis to use the energy of light to generate astrong reductant, reduced ferredoxin (fd)from the low-energy electrons of water.The downhill transfer of electrons betweenP680* and P700 represents a negative freeenergy change. Some of this energy is usedto establish a proton gradient, which inturn drives ATP synthesis. Indicated redoxpotentials are only approximate.

118 Chapter 7 / Energy Conservation in Photosynthesis: Harvesting Sunlight

binding site on the D1 polypeptide. The plastoquinol isthus released from the reaction center, to be replacedby another molecule of PQ. PQH2 diffuses from theQB site and becomes part of the PQ pool present in thethylakoid membrane. Since PQ requires two electronsto become fully reduced to PQH2, reduction at the QBsite is considered a two-electron gate (Figure 7.8).

Second, the initial charge separation is further sta-bilized because P680+ is a very strong oxidant (perhapsthe strongest known in biological systems) and is able to‘‘extract’’ electrons from water. Thus P680+ is rapidlyreduced, again within picoseconds, to P680, resultingin the formation of [P680 Pheo QA]. Now the PSIIreaction center is said to be ‘‘open,’’ that is, it is readyto receive another excitation (Figure 7.8). On a bright,sunny summer day, the photon flux to which a leaf canbe exposed may reach 2000 μmol photons m−2 s−1. This

4 Photons

4[P680 Pheo QA]

4[P680+ Pheo QA]

2PQH2

2PQ + 4H+

2H2O

O2 + 4H+ + 4e

“Closed” PSIIreaction centers

“Open” PSIIreaction centers

_

FIGURE 7.8 The light-dependent, cyclic ‘‘opening’’ and‘‘closing’’ on gate of photosystem II reaction centers.The PSII reaction center polypeptides, D1 and D2, bindthe following redox components irreversibly: P680, thereaction chlorophyll a; pheophytin (Pheo) and QA, thefirst stable quinone electron acceptor of PSII reactioncenters. The transfer of absorbed light energy to ‘‘open’’PSII reaction centers causes the photooxidation of P680and converts it P680+. Subsequently, the electron lost byP680 is transferred very rapidly first to Pheo and then toconvert it to Q−

A. This results in a stable charge separa-tion [P680+ Pheo Q−

A]. The PSII reaction is now said tobe ‘‘closed,’’ that is, the PSII reaction center cannot bephotooxidized since P680 is already photooxidized. Exci-tation of a ‘‘closed’’ reaction center results in irreversibledamage. Conversion of a ‘‘closed’’ PSII reaction centerto an ‘‘open’’ PSII reaction center requires the concomi-tant reduction of P680+ through the oxidation of waterand the transfer of the electron from Q−

A to plastoquinone(PQ) bound to the QB site on the D1 polypeptide of PSII.Upon the complete reduction, plastoquinone is proto-nated to form plastoquinol (PQH2) and released fromthe QB site and subsequently reduces the Cyt b6/f com-plex. The scheme illustrated is balanced for 4 excitationsand for the evolution of one molecule of O2. Note thatthe absorption of one photon causes the photooxidationof one P680.

means that about 1019 charge separations per secondmay occur over a leaf surface area of 1 cm2!

The electrons that reduce P680+ are most imme-diately supplied by a cluster of four manganese ions as-sociated with a small complex of proteins called theoxygen-evolving complex (OEC). As the name im-plies, the OEC is responsible for the splitting (oxidation)of water and the consequent evolution of molecularoxygen. The OEC is located on the lumen side of thethylakoid membrane. The OEC is bound to the D1 andD2 proteins of the PSII reaction center and functionsto stabilize the manganese cluster. It also binds Cl−which is necessary for the water-splitting function.

2H2O ⇀ O2 + 4H+ + 4e (7.8)

According to equation 7.8, the oxidation of twomoles of water generates one mole of oxygen, fourmoles of protons, and four moles of electrons. It hasbeen determined that only one PSII reaction centerand OEC is involved in the release of a single oxygenmolecule. Thus, in order to complete the oxidation oftwo water molecules, expel four protons, and producea single molecule of O2, the PSII reaction center mustbe ‘‘closed’’ and then ‘‘opened’’ four times (Figure 7.8).This means that PSII must utilize the energy of four pho-tons in order to evolve one molecule of O2. Experimentsin which electron transport was driven by extremelyshort flashes of light—short enough to excite essentiallyone electron at a time—have demonstrated that theOEC has the capacity to store charges. Each excitationof P680 is followed by withdrawal of one electron fromthe manganese cluster, which stores the residual positivecharge. When four positive charges have accumulated,the complex oxidizes two molecules of water and releasesthe product oxygen molecule. As a consequence, organ-isms containing PSII and the OEC exhibit oxygenicphotosynthesis, that is, a photosynthetic process thatgenerates molecular oxygen (O2).

However, not all photosynthetic organisms are oxy-genic. Photosynthetic bacteria are anoxygenic, that is,photosynthesis in these prokaryotes does not generatemolecular O2. Why is this? First, photosynthetic purplebacteria such as Rhodopseudomonas viridis and Rhodobactersphaeroides contain only one reaction center, the special-ized bacterial reaction center, rather than two reactioncenters found in chloroplasts of plants and green algae.Second, the bacterial reaction center contains the bac-teriochlorophyll a, P870, rather than P680. Excitationof P870 does not generate a sufficiently positive redoxpotential to oxidize water. Thus, these bacteria can notuse water as a source of electrons to reduce P870+but rather utilize a variety of other electron donors(see equations 7.4 and 7.5) such as hydrogen sulfide(H2S) and molecular hydrogen (H2). As a consequencethese photosynthetic bacteria are restricted to environ-ments which specifically contain either H2S or H2 as

7.3 Photosynthetic Electron Transport 119

a source of reducing power. Since water is generallyvery abundant in our biosphere, the evolution of PSIIallowed oxygenic photosynthetic organisms to surviveand reproduce almost anywhere in our biosphere. Thus,the evolution of PSII and its associated OEC was a majorfactor determining the global distribution of oxygenicphotosynthetic organisms that fundamentally changedthe development of all life on Earth.

7.3.3 THE CYTOCHROME COMPLEXAND PHOTOSYSTEM I OXIDIZEPLASTOQUINOL

Following its release from PSII, plastoquinol diffuseslaterally through the membrane until it encounters acytochrome b6f complex (Figure 7.6). This is anothermultiprotein, membrane-spanning redox complexwhose principal constituents are cytochrome b6 (Cyt b6)and cytochrome f (Cyt f ). The cytochrome complexalso contains an additional redox component calledthe Rieske iron-sulfur (FeS) protein—iron-bindingproteins in which the iron complexes with sulfurresidues rather than a heme group as in the case of thecytochromes. Plastoquinol diffuses within the plane ofthe thylakoid membrane and passes its electrons first tothe FeS protein and then to Cyt f . Since the oxidationof plastoquinol is thought to be diffusion limited, thisis the slowest step in photosynthetic electron transportand occurs on a time scale of milliseconds (ms). TheRieske FeS protein and the heme of Cyt f are locatedon the lumenal side of the thylakoid membrane. FromCyt f , the electrons are picked up by a copper-bindingprotein, plastocyanin (PC). PC is a small peripheralprotein that is able to diffuse freely along the lumenalsurface of the thylakoid membrane.

In the meantime, a light-driven charge separationsimilar to that involving P680 has also occurred inthe reaction center of PSI. Excitation energy trans-ferred to the reaction center chlorophyll of PSI (P700)is used to change the redox potential of P700 fromabout +0.4 eV to about −0.6 eV for P700*, the excitedform of P700 (Figure 7.7). As a consequence of thisinitial endergonic excitation, P700* is rapidly photoox-idized to P700+ by the primary electron acceptor (A)in PSI, a molecule of chlorophyll a (Figure 7.6); theelectron is then passed through a quinone and addi-tional FeS centers and finally, on the stroma side ofthe membrane, to ferredoxin. Ferredoxin is anotherFeS-protein that is soluble in the stroma. Ferredoxinin turn is used to reduce NADP+, a reaction mediatedby the enzyme ferredoxin-NADP+-oxidoreductase.Finally, the electron deficiency in P700+ is satisfied bywithdrawing an electron from reduced PC (Figure 7.6).

The overall effect of the complete electron trans-port scheme is to establish a continuous flow of electronsbetween water and NADP+, passing through the two

separate photosystems and the intervening cytochromecomplex (Figure 7.6). The bioenergetics of this processare illustrated in Figure 7.7. In the overall process,electrons are removed from water, a very weak reductant(Em = 0.82 V ), and elevated to the energy level offerredoxin, a very strong reductant (Em = − 0.42 V).Ferredoxin in turn reduces NADP+ to NADPH(Em = − 0.32 V). NADPH, also a strong reductant, isa water-soluble, mobile electron carrier that diffusesfreely through the stroma where it is used to reduce CO2in the carbon reduction cycle (Chapter 8). Since twoexcitations—at PSII and PSI—are required for eachelectron moved through the entire chain, a substantialamount of energy is put into the system. Based on one680 nm photon (175 kJ per mol quanta) and one 700 nmphoton (171 kJ per mol quanta), 692 kJ are used to exciteeach mole pair of electrons [2 × (175 + 171)]. Onlyabout 32 percent of that energy is conserved in NADPH(218 kJ mol−1).

What happens to the other 68 percent of the energy?An additional portion of the redox free energy of elec-tron transport energy is conserved as ATP. This occursin part because transfer of electrons between PSII andPSI is energetically downhill—that is, it is accompaniedby a negative �G (Figure 7.7). In the process of movingelectrons between plastoquinone and the cytochromecomplex, some of that energy is used to move protonsfrom the stroma side of the membrane to the lumenside. These protons contribute to a proton gradient thatcan be used to drive ATP synthesis by chemiosmosis(Chapter 5).

The quantum requirement for oxygen evolutionis defined as the number photons required to evolveone molecule of O2. As discussed above, two excitations,one at PSII and one at PSI, are required to moveeach electron through noncyclic electron transport fromH2O to NADP+. From equation 7.8, the evolution ofone molecule of O2 by P680+ generates four electronsthat are eventually transferred through PSI to reduce2NADP+ to 2NADPH. Thus, to transfer four electronsfrom H2O to NADP+ requires 8 photons. Thereforethe minimal theoretical quantum requirement for O2evolution is 8 photons/molecule of O2 evolved.

Conversely, quantum yield of oxygen evolutionis the inverse of quantum requirement, that is, num-ber of molecules of O2 evolved per photon absorbed.Since this is also the definition of photosynthetic effi-ciency, the terms quantum yield and photosyntheticefficiency are interchangeable. Consequently, the maxi-mum theoretical quantum yield for oxygen evolution orphotosynthetic efficiency of O2 evolution must be 1/8 or0.125 molecules of O2 evolved/photon absorbed. Pho-tosynthetic efficiency or quantum yield for O2 evolutionwill vary depending on the environmental conditions towhich a plant is exposed. This will be discussed in moredetail in Chapters 13 and 14.

120 Chapter 7 / Energy Conservation in Photosynthesis: Harvesting Sunlight

7.4 PHOTOPHOSPHORYLATIONIS THE LIGHT-DEPENDENTSYNTHESIS OF ATP

In Chapter 5, we examined the bioenergetics of the light-dependent synthesis of ATP. However, by definition,thermodynamics does not provide specific informationwith respect to kinetics and biochemical mechanism.Here we discuss the molecular basis underlying thechemiosmotic synthesis of ATP in chloroplasts.

The ATP required for carbon reduction and othermetabolic activities of the chloroplast is synthesizedby photophosphorylation in accordance with Mitchell’schemiosmotic mechanism (Chapter 5). Light-drivenproduction of ATP by chloroplasts is known as photophosphorylation. Photophosphorylation is very impor-tant because, in addition to using ATP (along withNADPH) for the reduction of CO2, a continual supplyof ATP is required to support a variety of other meta-bolic activities in the chloroplast. These activities in-clude amino acid, fatty acid, and starch biosynthesis,the synthesis of proteins in the stroma, and the trans-port of proteins and metabolites across the envelopemembranes.

When electron transport is operating accordingto the scheme shown in Figures 7.6 and 7.7, electronsare continuously supplied from water and withdrawn asNADPH. This flow-through form of electron transportis consequently known as either noncyclic or linearelectron transport. Formation of ATP in associationwith noncyclic electron transport is known as noncyclicphotophosphorylation. However, as will be shownlater, PSI units and PSII units in the membrane are notphysically linked as implied by the Z scheme, but areeven segregated into different regions of the thylakoid.

One consequence of this heterogeneous distribution inthe membranes is that PSI units may transport electronsindependently of PSII, a process known as cyclicelectron transport. In terrestrial plants, the majorpathway for PSI cyclic electron transport is thought tooccur via P700 to ferredoxin (fd) which transfers theelectrons back to PQ via a recently discovered protein,PGR5 rather than to NADP+. The electron thenreturns to P700+, passing through the cytochrome b6/fcomplex and plastocyanin. Using a genetic approach inArabidopsis thaliana, the gene, PGR5, was shown toencode a small thylakoid polypeptide that is essential forPSI cyclic electron transport (Figure 7.9). However, theprecise role of PGR5 in the electron transfer processhas yet to be elucidated. Since these electrons also passthrough PQ and the cytochrome complex, cyclic elec-tron transport will also contribute to the establishmentof the pH gradient required to support ATP synthesis,a process known as cyclic photophosphorylation. It isthought that cyclic photophosphorylation is a source ofATP required for chloroplast activities over and abovethat required in the carbon-reduction cycle. Since non-cyclic photophosphorylation results in the productionof both ATP and NADPH whereas cyclic photophos-phorylation does not generate NADPH, switchingbetween cyclic and noncyclic photophosphorylationalso represents a mechanism by which the chloroplastcan regulate the stromal ATP/NADPH ratios, which isimportant in the maintenance of chloroplast metabolicactivity.

A key to energy conservation in photosynthetic elec-tron transport and the accompanying production of ATPis the light-driven accumulation of protons in the lumen.There are two principal mechanisms that account forthis accumulation of protons: the oxidation of water, inwhich two protons are deposited into the lumen for each

FIGURE 7.9 Cyclic electron trans-port. PSI units operating indepen-dently of PSII may return elec-trons from P700 through ferre-doxin (fd), and PGR5 to the thy-lakoid plastoquinone (PQ) pooland the cytochrome b6/f complex.In cyclic electron transport, theoxidation of PQ by the cytochromeb6/f complex generates a protongradient that can be used for ATPsynthesis but no NADPH is pro-duced.

STROMA SIDE

LUMEN SIDE

ADP

fdPGR5

PC PC

P700

ALHC

Cyt fFe-s

2H+

PQCyt b6

ATP

H+

+ Pi

PQH Pool2PQH2

PQ PQ I

NADPHNADP+

7.4 Photophosphorylation is the Light-Dependent Synthesis of ATP 121

water molecule oxidized, and a PQ-cytochrome protonpump. The energy of the resulting proton gradient isthen used to drive ATP synthesis in accordance withMitchell’s chemiosmotic hypothesis (Chapter 5).

The precise mechanism by which protons are movedacross the membrane by the cytochrome complex is notyet understood, although several models have been pro-posed. The most widely accepted model is known asthe Q-cycle, based on an original proposal by Mitchell.A simplified version of the Q-cycle during steady-stateoperation is shown in Figure 7.10. When PQ is reducedby PSII, it binds temporarily to the D1 protein (QB)as a semiquinone after it accepts the first electron fromQA. Subsequently, the QB semiquinone is convertedto the fully reduced plastoquinol (PQH2) after it hasaccepted another electron from QA, plus two protonsare picked up from the surrounding stroma. PQH2dissociates from the PSII complex and diffuses lat-erally through the membrane until it encounters thelumenal PQH2 binding site of the cytochrome b6/fcomplex. There, two PQH2 bind sequentially and arereoxidized to PQ through a semiquinone intermedi-ate (PQHz, not shown) by the combined action ofthe Rieske FeS-protein and the low reduction potentialform of cytochrome b6 (LP). Concomitantly, 4 H+ aretransferred to the lumen. One of the PQ moleculesreturns to the thylakoid PQ pool to be reduced again byPSII, while the other PQ molecule is transferred to the

PS II

2H+

CP43

PQH2

PQH2

2PQH2Pool

PQ

PQ

4H+

2H+

PQ

2e

2e

2e

2e

2e

2e2P700

+

2FeS2Cytf

2PC

2CytbLP

2CytbHP

STROMAL SIDE

LUMEN SIDE

Cytochromecomplex

Pheo

2H2O

O2+4H+

CP47

D2

P6 80

OEC

Mn2+

QB

D1QA

LHCII

FIGURE 7.10 The Q-cycle, a model for coupling electron transport from plastoquinol(PQH2) to the cytochrome b6/f complex (Cyt b6/f ) with the translocation of protonsacross the thylakoid membrane. Six protons are translocated for each pair of elec-trons that passes through the electron transport chain. Fe, Rieske FeS-center; Cyt f ,cytochrome f of the cytochrome b6/f complex; Cyt bLP, low reduction potential formof cytochrome b6 of the cytochrome b6/f complex; Cyt bHP, high reduction potentialform of cytochrome b6 of the cytochrome b6/f complex; PC, plastocyanin.

stromal binding site of the cytochrome b6/f complexwhere it becomes reduced by the high reduction poten-tial form of cytochrome b6 (HP) and is protonated using2 H+ from the stroma. This PQH2 molecule is thenreleased from the stromal binding site and recycled intothe thylakoid PQH2 pool.

Thus, for each pair of electrons passing from plasto-quinone through the Rieske FeS-center and cytochromef to plastocyanin, four protons are translocated from thestroma into the lumen of the thylakoid. If this schemeis correct, then each pair of electrons passing throughnoncyclic electron transport from water to NADP+contributes six protons to the gradient—four from theQ-cycle (Equation 7.9) plus two from water oxidation(Equation 7.10).

2PQH2 ⇀ PQ + PQH2 + 4H+ + 2e (7.9)

therefore 4H+/2e

H2O ⇀ 1/2O2 + 2H+ + 2e (7.10)

therefore 2H+ /2eFor cyclic electron transport, the number of protons

transferred per pair of electrons would be four.Since it is generally agreed that three protons

must be transported through the CF0-CF1 for eachATP synthesized (3H+/ATP), a pair of electrons pass-ing through noncyclic electron transport would beexpected to yield two ATP molecules for every NADPH

122 Chapter 7 / Energy Conservation in Photosynthesis: Harvesting Sunlight

produced (2ATP/NADPH). The precise stoichiometry,however, is difficult to determine, in part because ofuncertainty with regard to the relative proportions ofcyclic and noncyclic photophosphorylation occurring atany specific moment in time.

According to the chemiosmotic theory,pmf = −59�pH + �� (7.11)

As protons accumulate in the lumen relative to thestroma, divalent magnesium ions (Mg2+) released fromthe thylakoid membrane accumulate in the chloro-plast stroma. This minimizes the difference in electricalcharge between the stroma and the lumen. Thus, inchloroplasts, the �pH (i.e., the H+ concentration gradi-ent) is the major factor that contributes to chloroplasticpmf, whereas �� contributes minimally.

7.5 LATERAL HETEROGENEITYIS THE UNEQUALDISTRIBUTION OFTHYLAKOID COMPLEXES

In addition to the vectorial arrangement of electrontransport components across the membrane thataccounts for the simultaneous electron transfer withinthe thylakoid membrane and active transport of protonsfrom the stroma to the lumen, there is also a distinctlateral heterogeneity with respect to their distributionof the major protein complexes within the thylakoids(Figure 7.11). The result is that PSI and PSII, forexample, are spatially segregated, rather than arrangedas some kind of supercomplex that might be suggestedby the static representation in the previous figures.The PSI/LHCI complexes and the CF0-CF1 ATPaseare located exclusively in nonappressed regions of thethylakoid; that is, those regions where the membranesare not paired to form grana. These regions includethe stroma thylakoids, the margins of the grana stacks,and membranes at either end of the grana stacks,all of which are in direct contact with the stroma(Figure 7.11A). Virtually all of the PSII complexes andLHCII, on the other hand, are located in the appressedregions of the grana membranes (Figure 7.11B). Thecytochrome b6/f complexes are uniformly distributedthroughout both regions.

Spatial segregation also requires that the electrontransport complexes be linked with each other throughone or more mobile carriers that can deliver electronsbetween complexes. These carriers are plastoquinone(PQ), plastocyanin (PC), and ferredoxin. All three aremobile carriers that are not permanently part of any elec-tron transport complex. Plastoquinone is a hydrophobicmolecule and is consequently free to diffuse laterallywithin the lipid matrix of the thylakoid membrane. Itsestimated diffusion coefficient is 106 cm−2 s−1, whichmeans that it could travel more than the diameter ofa typical granum in less than one millisecond. The

Grana thylakoids Stroma thylakoid

Granum end membrane

Margins

Appressedregions

Non-appressedregions

PSI PSII Cyt b6/f CF0 - CF1 ATPaseB.

A.

FIGURE 7.11 Lateral heterogeneity in the thylakoid mem-brane. (A) Nonappressed membranes of the stroma thy-lakoids, grana end membranes, and grana margins areexposed to the stroma. Appressed membranes in theinterior of grana stacks are not exposed to the stroma. (B)PSII units are located almost exclusively in the appressedregions while PSI and ATP synthase units are locatedin nonappressed regions. The cytochrome b6/f com-plex, plastoquinone, and plastocyanin are uniformlydistributed throughout the membrane system.

lateral mobility of PQ allows it to carry electronsbetween PSII and the cytochrome complex. Plasto-cyanin is a small (10.5 kDa) peripheral copper-proteinfound on the lumenal side of the membrane. It read-ily diffuses along the lumenal surface of the membraneand carries electrons between the cytochrome complexand PSI. Ferredoxin, a small (9 kDa) iron-sulfur pro-tein, is found on the stroma side of the membrane. Itreceives electrons from PSI and, with the assistance ofthe ferredoxin-NADP oxidoreductase, reduces NADP+to NADPH.

An unequal number and spatial segregation ofPSI and PSII means that both cyclic and noncyclicphotophosphorylation can occur more or less simulta-neously. Thus the output of ATP and NADPH can beadjusted to meet the demands not only of photosynthe-sis but of other biosynthetic energy requirements withinthe chloroplast (see Chapter 8).

Are granal stacks required for oxygenic photosyn-thesis? The unequivocal answer to this question is no.Contrary to the illusion created by a transmission elec-tron micrograph of a chloroplast (see Figure 5.4), thestacking of thylakoid membranes into grana is not staticbut very dynamic. If granal stacks are not required foroxygenic photosynthesis, what regulates their forma-tion and why do they occur in most chloroplasts? First,

7.6 Cyanobacteria are Oxygenic 123

in vitro experiments with isolated thylakoids showedthat decreasing the concentration of monovalent cationssuch as K+ and Mg2+, respectively, in the surroundingthylakoid isolation buffer decreases stacking and resultsin the homogeneous distribution of the photosyntheticelectron protein complexes within the thylakoid mem-brane. This is reversible upon the re-addition of thesecations which induces stacking of the thylakoids andthe reestablishment of lateral heterogeneity. Later, itwas established that the N-terminal domain of themajor LHCII polypeptides mediated the stacking pro-cess. Wild type barley chloroplasts exhibit typical granalstacks whereas the chloroplasts of the chlorina f2 mutantof barley, which lacks both chlorophyll b and the majorLHCII polypeptides, does not exhibit granal stacks.Proteolytic cleavage of this N-terminal domain of themajor LHCII polypeptides also inhibited granal stack-ing. It was concluded that the presence of cations shieldsthe negative surface charges created by the exposedN-terminal domains of the major LHCII polypeptides

present thylakoid membranes. Thus, unstacking of thy-lakoid membranes is the result of electrostatic repulsionof the negative surface charges on thylakoid membranes.The possible functions of this remarkable process arestill not completely understood. We will return to thissubject in Chapter 13 when we discuss the regulation ofenergy distribution between PSII and PSI.

7.6 CYANOBACTERIA AREOXYGENIC

In contrast to photosynthetic bacteria, cyanobacteria(Figure 7.12A) are a large and diverse group of pro-karyotes which perform oxygenic photosynthesisbecause they exhibit PSI as well as PSII with itsassociated OEC and an intersystem electron transportchain comparable to that of eukayotic photoautotrophs.However, in contrast to the intrinsic, major light har-

FIGURE 7.12 (A) An electron micro-graph of a typical single cell cyanobac-terium, Synechocystis. Note thatthere are no granal stacks. The thy-lakoid membranes are arranged inconcentric rings in the cell cyto-plasm. (B) A confocal micrographof the cyanobacterium, Plectonemaboryanum. This is an example of afilamentous cyanobacterium. Each redcircle represents one cell and is visibledue to the red fluorescence emanatingfrom chlorophyll a. (C) General struc-ture of PSII and its associated phycobil-isome in cyanobacteria. Note that thephycobilisome is associated with PSII atthe surface of the thylakoid membrane.Phycoerythrin (red) and phycocyanin(dark blue) are the pigments bound tothe protein rod structure. The rods arebound to the allophycocyanin proteins(blue circles) which bind the phycobili-some to CP43 and CP47 of PSII.

OEC

Mn2+

2H2O

O2 + 4H+

LUMEN SIDE

CYTOPLASMSIDE

Phycobilisome

ThylakoidMembrane

PQ

PQH2 Pool

PQH2

PQ PQ

2H+

2H+

Cyt fFe-s

Cyt b6

A.A. B.

C.

thylakoids

CP43

PC

Pheo

CP47D2

P6 80

QB

D1QA

124 Chapter 7 / Energy Conservation in Photosynthesis: Harvesting Sunlight

vesting pigment-protein complex found in chloroplastthylakoid membranes of plants and green algae, thelight harvesting complex of cyanobacteria is an extrinsicpigment-protein complex called a phycobilisomewhich is bound to the outer, cytoplasmic surface ofcyanobacterial thylakoids (Figure 7.12B). Phycobili-somes (PBSs) are rod-shaped chromoproteins calledphycobiliproteins which may constitute up to 40 percentof the total cellular protein. The phycobiliproteinsusually associated with PBS include allophycocyanin(AP), phycocyanin (PC), and phycoerythrin (PE). Inaddition to PBS, PSII of cyanobacteria include the Chla core antenna CP47 and CP43 similar to that foundin eukaryotic organisms. Cyanobacteria are distinctfrom chloroplasts because the redox carriers involved inrespiratory as well as photosynthetic electron transportare located in the cyanobacterial thylakoid membraneswhere they share a common PQ pool and a commonCyt b6f complex. Because PBSs are large, extrinsicpigment-protein complexes, this prevents appression ofcyanobacterial thylakoid membranes and the formationof granal stacks characteristic of eukaryotic chloroplasts.This is further evidence that granal stacks are not aprerequisite for oxygenic photosynthesis.

7.7 INHIBITORS OFPHOTOSYNTHETICELECTRON TRANSPORT AREEFFECTIVE HERBICIDES

Since the dawn of agriculture, man has waged war againstweeds. Weeds compete with crop species for water,nutrients, and light and ultimately reduce crop yields.Traditional methods of weed control, such as crop rota-tion, manual hoeing, or tractor-drawn cultivators werelargely replaced in the 1940s by labor-saving chemicalweed control. Modern agriculture is almost completelydependent upon the intensive use of herbicides.

A wide spectrum of herbicides is now available thatinterfere with a variety of cell functions. Many of thecommercially more important herbicides, however, actby interfering with photosynthetic electron transport.Two major classes of such herbicides are derivativesof urea, such as monuron and diuron, and the triazineherbicides, triazine and simazine (Figure 7.13). Boththe urea and triazine herbicides are taken up by theroots and transported to the leaves. There they bindto the QB binding site of the D1 protein in PSII (alsoknown as the herbicide-binding protein). The herbicideinterferes with the binding of plastoquinone to thesame site and thus blocks the transfer of electrons toplastoquinone. Because of its action in blocking electrontransport at this point, DCMU is commonly used inlaboratory experiments where the investigator wishes toblock electron transport between PSII and PSI.

3-(3,4-Dichlorophenyl)-1,1-dimethylurea(common names: Diuron, DCMU)

33

2-Chloro-4-ethylamino-6-isopropylamino-s-triazine(common name: Atrazine)

Paraquat(methyl viologen)

++ NCHH CN

CH3

CH3

Cl

Cl

HN C

O

N

Cl

N

C2H5 CH(CH )2NH

N

N NH

FIGURE 7.13 The chemical structures of some commonherbicides that act by interfering with photosynthesis.

The triazine herbicides are used extensively to con-trol weeds in cornfields, since corn roots contain anenzyme that degrades the herbicide to an inactive form.Other plants are also resistant. Some, such as cotton,sequester the herbicide in special glands while othersavoid taking it up by way of root systems that pene-trate deep below the application zones. In many cases,however, weeds have developed triazine-resistant races,or biotypes. In several cases, the resistance has beentraced to a single amino acid substitution in the D1protein. The change in amino acid reduces the affinityof the protein for the herbicide but does not interferewith plastoquinone binding and, consequently, electrontransport.

The availability of herbicide-resistant genestogether with recombinant DNA technology hasstimulated considerable interest in the prospects fordeveloping additional herbicide-resistant crop plants.It is possible, for example, to transfer the gene forthe altered D1 protein into crop species and conferresistance to triazine herbicides. This approach willbe successful, however, only if weed species do notcontinue to acquire resistance to the same herbicidesthrough natural evolutionary change.

7.7 Inhibitors of Photosynthetic Electron Transport are Effective Herbicides 125

H20

1/202 + 2H+

Cyt PS I2e

hν

PS II

hν

NADPH + H+

NADP+

+ 2H+

BOX 7.2THE CASEFOR TWOPHOTOSYSTEMS

The photosynthetic unit of oxygenic photosyntheticorganisms is organized as two separate photosystemsthat operate in series. While the two-step series for-mulation, or ‘‘Z-scheme,’’ for photosynthesis is gen-eral knowledge today, the idea generated considerableexcitement when it was first proposed in the early 1960s.The two-step idea was based on a series of exper-iments conducted during the 1950s, which laid thefoundation for significant advances in our understand-ing of photosynthetic electron transport. The first ofthese experiments was centered around the concept ofquantum efficiency. Information about quantum effi-ciency is very useful when attempting to understandphotochemical processes. Quantum efficiency can beexpressed in two ways—either as quantum yield or asquantum requirement. Quantum yield (φ) expressesthe efficiency of a process as a ratio of the yieldof product to the number of photons absorbed. Inphotosynthesis, for example, product yield would bemeasured as the amount of CO2 taken up or O2evolved. Alternatively, the quantum requirement (1/φ)(sometimes referred to as quantum number) tells howmany photons are required for every molecule of CO2reduced or oxygen evolved. Equation 7.8 identifies thata minimum of four electrons are required for everymolecule of CO2 reduced. In Chapter 5 it was estab-lished that one photon is required for each electronexcited.

Therefore, the minimum theoretical quantumrequirement for photosynthesis is four. However, it hasbeen well established experimentally that the minimumquantum requirement for photosynthesis is eight toten photons for every CO2 reduced. If eight photonsare required (it is usual to assume the minimum) forfour electrons, then each electron must be excited twice!A second line of evidence, again from the laboratoryof R. Emerson, was based on attempts to determinethe action spectra for photosynthesis in Chlorella.Emerson and his colleague C. M. Lewis reported in1943 that the value of φ was remarkably constant overmost of the spectrum (Emerson and Lewis, 1943).This would indicate that any photon absorbed bychlorophyll was more or less equally effective in drivingphotosynthesis. However, there was an unexpecteddrop in the quantum yield at wavelengths greater than680 nm, even though chlorophyll still absorbed in that

Qua

ntum

Yie

ld (

Φ)

Abs

orba

nce

0.1

0.09

0.08

0.07

0.06

0.05

0.04

Wavelength (nm)

400 440 480 540 560 600 640 680 720

FIGURE 7.14 The Emerson ‘‘red drop’’ in the green algaChlorella. Lower curve: Absorption spectrum of photo-synthetic pigments. Upper curve: Action spectrum forquantum yield of photosynthesis. (Redrawn from the dataof R. Emerson, C. M. Lewis, American Journal of Botany30:165–178, 1943).

range (Figure 7.14). This puzzling drop in quantumefficiency in the long red portion of the spectrum wascalled the red drop.

In another experiment, Emerson and his colleaguesset up two beams of light—one in the region of650 to 680 nm and the other in the region of 700to 720 nm. The fluence rates of both beams wereadjusted to give equal rates of photosynthesis. Emersondiscovered that when the two beams were appliedsimultaneously, the rate of photosynthesis was two tothree times greater than the sum of the rates obtainedwith each beam separately! This phenomenon hasbecome known as the Emerson enhancement effect(Figure 7.15). The enhancement effect suggests thatphotosynthesis involves two photochemical eventsor systems, one driven by short-wavelength light(≤680 nm) and one driven by long-wavelength light(>680 nm). For optimal photosynthesis to occur, bothsystems must be driven simultaneously or in rapidsuccession.

In an attempt to explain conflicting informationabout the role of cytochromes and redox potential values,R. Hill and Fay Bendall, in 1960, proposed a newmodel for electron transport. The Hill and Bendallmodel involved two photochemical acts operating inseries—one serving to oxidize the cytochromes and oneserving to reduce them (Figure 7.16). The followingyear, L. Duysens confirmed the Hill and Bendall model,showing that cytochromes were oxidized in the presenceof long-wavelength light. The effect could be reversedby short-wavelength light.

126 Chapter 7 / Energy Conservation in Photosynthesis: Harvesting Sunlight

FIGURE 7.15 Schematic to illustrate theEmerson ‘‘enhancement effect.’’ Twobeams of light (660 nm and 710 nm)were presented either singly (A andB) or in combination (C). Beam ener-gies were adjusted to give equal ratesof oxygen evolution. When presentedsimultaneously, the rate of oxygen evo-lution exceeded the sum of the rateswhen each beam was presented singly.Up arrows indicate light on. Downarrows indicate light off. (Reproducedwith permission from the AnnualReview of Plant Physiology, Vol. 22,copyright 1971 by Annual Reviews,Inc.)

710nm660nm

02 E

volu

tion

660nm

710nm+

0A B C

A + B

Although the scheme has been significantly modi-fied and considerable detail has been added since itwas originally proposed, the Hill and Bendall schemeprovided the catalyst that has led to our presentunderstanding of photosynthetic electron transport and

0.6

0.4 0.41

0.2

0.0

0.2

0.4

0.0

0.375

0.6

0.8 0.81

1.0

NADPH

Volt

s

Cyt b6

Cyt f

H2O

FIGURE 7.16 The Z scheme as originally proposed byHill and Bendall. For a current version, see Figure 7.7.(Redrawn from Hill and Bendall, 1960).

oxygen evolution. As a consequence, today the Z-schemeis the prevailing paradigm for photosynthetic electrontransport in plants, algae, and cyanobacteria. However,Daniel Arnon spent part of his illustrious scientific careerchallenging the Z-scheme. Supported with experimentalevidence, he maintained until his death in 1995 that PSIand PSII could operate independently of one anotherand still support CO2 assimilation. He maintained thatPSI can operate in a cyclic mode to generate ATPand PSII could reduce NADP+ directly. Although thereduction of NADP+ by PSII can be shown in vitro, thequantum yield for this process appears to be very lowand this reaction has never been reported in vivo. If PSIIcould reduce NADP+ directly, one would expect to seeCO2 fixation in the absence of PSI. However, all exper-iments with mutants of the green alga, Chlamydomonasreinhardtii, that lack PSI indicate that these mutantsare unable to fix CO2. Although the consensus is thatthe Z-scheme reflects an accurate description of pho-tosynthetic electron flow in photosynthetic organismsgrown under optimal growth conditions, data continueto accumulate that indicate that the Z-scheme maynot explain photosynthetic electron flow during growthunder extreme conditions. For recent controversies sur-rounding the Z-scheme see the papers by Redding andPeltier (1998) as well as by Ivanov et al. (2000).

REFERENCES

Arnon, D. I. 1995. Divergent pathways of photosyntheticelectron transfer: The autonomous oxygenic andanoxygenic photosystems. Photosynthesis Research46:47–71.

Duysens, L. N. M., J. Amesz, B. M. Kamp. 1961. Twophotochemical systems in photosynthesis. Nature190:510–511.

Emerson, R., C. M. Lewis. 1943. The dependence of thequantum yield of Chlorella photosynthesis on wavelengthof light. American Journal of Botany 30:165–178.

Chapter Review 127

Emerson, R., R. Chalmers, C. Cederstrand. 1957. Somefactors influencing the long-wave length limit ofphotosynthesis. Proceedings of the National Academy ofScience USA 43:133–143.

Hill, R., F. Bendall. 1960. Function of the two cytochromecomponents in chloroplasts: A working hypothesis.Nature 186:136–137.

Ivanov, A. G., Y.-I. Park, E. Miskiewicz, J. A. Raven, N. P. A.Huner, G. Oquist. 2000. Iron stress restricts

photosynthetic intersystem electron transport inSynechococcus sp. PCC 7942. FEBS Lett. 485:173–177.

Redding, K., G. Peltier. 1998. Reexamining the validity ofthe Z-scheme: Is photosystem I required for oxygenicphotosynthesis in Chlamydomonas? In: J.-D. Rochaix,M. Goldschmidt-Clermont, S. Merchant (eds.), TheMolecular Biology of Chloroplasts and Mitochondria inChlamydomonas? Advances in Photosynthesis, Vol. 7,pp. 349–362. Dordrecht: Kluwer Academic.

Another class of herbicides are the bipyridyliumviologen dyes—paraquat (Figure 7.13)—which act byintercepting electrons on the reducing side of PSI. Theviologen dyes are auto-oxidizable, immediately reduc-ing oxygen to superoxide. Not only do the viologendyes interfere with photosynthetic electron transport,but the superoxide they produce causes additional dam-age by rapidly inactivating chlorophyll and oxidizingchloroplast membrane lipids. Because viologen herbi-cides are also highly toxic to animals, their use is bannedor tightly regulated in many jurisdictions.

Chemical herbicides have become an importantmanagement tool for modern agriculture, but theirvalue as a labor-saving device must be carefully weighedagainst potentially harmful ecological effects. Many ofthese herbicides are carcinogenic, and thus the potentialaccumulation of these hazardous compounds in watersupplies continues to be a major public concern. Inaddition, the overuse of herbicides promote herbicidetolerance in weeds, which exacerbates the weed problemin the long term.

SUMMARY

The function of the light-dependent reactions ofphotosynthesis is to generate the ATP and reducingpotential (as NADPH) required for subsequentcarbon reduction. The electron transport chain inthe thylakoid membranes of oxygenic photoautrophsis composed of two photosystems (PSI, PSII) and acytochrome b6f complex. The three complexes arelinked by plastoquinone and plastocyanin, mobilecarriers that freely diffuse within the plane of the mem-brane. Each photosystem consists of a reaction center,core antenna, and associated light-harvesting (LHC)complexes. Light energy gathered by the antenna andLHC is passed to the reaction center. In the reactioncenter, electron flow is initiated by a charge separation(photooxidation). As a result, electrons obtained fromthe oxidation of water are passed through PSII, thecytochrome b6f complex, and PSI to NADP+. Protonspumped across the membrane between PSII and PSIdrive photophosphorylation. The components of

the photosynthetic electron transport chain are notdistributed homogeneously throughout the thylakoidmembranes of eukaryotic chloroplasts but exhibitlateral heterogeneity. However, lateral heterogeneityis dynamic and is a consequence electrostatic shieldingof negative surface charges on thylakoid membranescreated by the exposed N-terminal domains of themajor LHCII polypeptides. Granal stacks are not anabsolute requirement for oxygenic photosynthesis.Although cyanobacteria exhibit PSI, PSII and areoxygenic, thylakoids of these prokaryotes do notexhibit granal stacks due to the presence of extrinsicpigment-protein complexes called phycobilisomes. Incontrast to eukaryotic photosynthetic organisms andcyanobacteria, photosynthetic bacteria contain onlyone specialized bacterial reaction center that oxidizesH2S or H2 and is incapable of oxidizing H2O. Severalclasses of economically important herbicides act byinterfering with photosynthetic electron transport.

CHAPTER REVIEW

1. ATP formation in chloroplasts is based on thestepwise conservation of energy. Trace theconservation of energy from the initial absorptionof light by an antenna chlorophyll moleculeto the final formation of a molecule of ATP.

2. Describe the concept of a photosystem and how itis involved in converting light energy to chemicalenergy.

3. Explain the difference between cyclic and non-cyclic electron transport. How can noncyclicphotosynthetic electron transport function if thePSII and PSI units are located in different re-gions of the thylakoid membrane?

4. How is lateral heterogeneity regulated?5. Explain the difference between oxygenic photo-

synthesis and anoxygenic photosynthesis. Whatrole did the evolution of oxygenic photosynthesisplay in the global distribution of photosyntheticorganisms?

128 Chapter 7 / Energy Conservation in Photosynthesis: Harvesting Sunlight

6. What is LHCII and where is it localized? Whatare phycobilisomes and where are they localized?

7. The herbicide DCMU is commonly used inlaboratory investigations of electron transportreactions in isolated chloroplasts. Can you suggestwhy DCMU might be useful for such studies?

FURTHER READING

Aro, E.-M., B. Andersson. 2001. Regulation of Photosynthesis.Advances in Photosynthesis and Respiration, Vol. 11. Dor-drecht: Kluwer.

Blankenship, R. E. 2002. Molecular Mechanisms of Photosynthe-sis. Williston: Blackwell Science.

Buchanan, B. B., W. Gruissem, R. L. Jones. 2000.Biochemistry and Molecular Biology of Plants. Rockville,MD: American Society of Plant Physiologists.

Merchant, S., M. R. Sawaya. 2005. The light reactions: Aguide to recent acquisitions for the picture gallery. PlantCell 17:648–663.

Nelson, N., C. F. Yocum. 2006. Structure and function ofphotosystems I and II. Annual Review of Plant Biology57:521–565.

Ort, D. T., C. F. Yokum (eds.). 1996. Oxygenic Photosynthesis:The Light Reactions. Advances in Photosynthesis and Respira-tion, Vol. 4. Dordrecht: Kluwer.

Shikanai, T. 2007. Cyclic electron transport around photo-system I: Genetic approaches. Annual Review of PlantBiology 58:199–217.

PR

R GP

CO2

CO2CO2

O2

O2O2

8Energy Conservation in Photosynthesis:

CO2 Assimilation

Chapter 7 showed how chloroplasts conserve lightenergy by converting it to reducing potential, in theform of NADPH and ATP. In this chapter, attention isfocused on how CO2 enters the leaf and subsequentlyis reduced through the utilization of the NADPH andATP produced by photosynthetic electron transport. Airis the source of CO2 for photosynthesis. Gas exchangebetween the leaf and the surrounding air is depen-dent upon diffusion and is controlled by the openingand closing of special pores called stomata. Stomatalmovements are very sensitive to external environmentalfactors such as light, CO2, water status, and tempera-ture. The reactions involved in the reduction of CO2have been traditionally designated the ‘‘dark reactions’’of photosynthesis, but this designation is quite mislead-ing, since it implies they can proceed in the absence oflight. However, several critical enzymes in the carbonreduction cycle are light activated; in the dark they areeither inactive or exhibit low activity. Consequently, car-bon reduction cannot occur in the dark, even if energycould be made available from some source other thanthe photochemical reactions.

Only a few decades ago, knowledge of carbonmetabolism was in its infancy and, as is to be expectedwhen opening up new areas of study, understand-ing of the process was somewhat unsophisticated.

Photosynthesis and respiration, long recognized asthe two major divisions of carbon metabolism, werethought to be separate and independent metabolicprocesses, neatly compartmentalized in the cell.Photosynthesis was localized in the chloroplast whilerespiration appeared restricted to the cytoplasm andthe mitochondrion. The task of photosynthesis was toreduce carbon and store it as sugars or starch. Whenrequired, these storage products could be mobilizedand exported to the mitochondrion where they wereoxidized to satisfy the energy and carbon needs ofthe cell through respiration. Thus, the relationshipsbetween photosynthesis and respiration appearedsimple and uncomplicated.

Over the past 40 years, however, knowledge andunderstanding of carbon metabolism has improved con-siderably and, with that, so has the apparent complexity.Photosynthetic carbon metabolism can no longer beexplained by a single, invariable cycle. It is no longerrestricted to just the chloroplast or even to a single cell.In addition to carbon reduction, photosynthetic energyis used to drive nitrogen assimilation, sulphate reduc-tion, and other aspects of intermediary metabolism. Therate of photosynthesis is influenced and even controlledby events occurring outside the chloroplast and else-where in the plant. These and other complexities of

129

130 Chapter 8 / Energy Conservation in Photosynthesis: CO2 Assimilation

metabolic integration within the cell and between dif-ferent parts of the plant are only beginning to becomeobvious. One thing is certain: a more holistic approachto carbon metabolism is called for. The traditional,compartmentalized vision of independent processes nolonger adequately explains carbon metabolism in plants.

This chapter will describe several interrelated vari-ations of photosynthetic carbon metabolism in higherplants, including

• leaf gas exchange through stomatal pores that pro-vide an efficient mechanism for the absorption ofCO2 along a shallow concentration gradient,

• the path of carbon, energetics, and regulation ofthe photosynthetic carbon reduction cycle, or C3metabolism—the pathway that all organisms ulti-mately use to assimilate carbon, and

• photorespiration and how limitations are imposedon carbon assimilation in C3 plants by the photo-synthetic carbon oxidation cycle

Later chapters will address carbon partitioning, res-piratory carbon metabolism, and factors influencing thedistribution of carbon throughout the plant as well as theecological significance of C4 and CAM (CrassulaceanAcid Metabolism) photosynthesis.

8.1 STOMATAL COMPLEXCONTROLS LEAF GASEXCHANGE AND WATERLOSS

The epidermis of leaves contains pores that provide forthe exchange of gases between the internal air spacesand the ambient environment. The opening, or stoma,is bordered by a pair of unique cells called guard cells(Figure 8.1). In most cases the guard cells are in turnsurrounded by specialized, differentiated epidermal cellscalled subsidiary cells. The stoma, together with itsbordering guard cells and subsidiary cells, is referred toas the stomatal complex, or stomatal apparatus.

The distinguishing feature of the stomatal complexis the pair of guard cells that functions as a hydrauli-cally operated valve. Guard cells take up water andswell to open the pore when CO2 is required for pho-tosynthesis, and lose water to close the pore whenCO2 is not required or when water stress overridesthe photosynthetic needs of the plant. The mechan-ical, physiological, and biochemical properties of theguard cells have attracted scholars almost since theiroccurrence was first reported by M. Malpighi in thelate seventeenth century. A continuing interest in stom-atal movement is understandable, given the foremostimportance of stomata in regulating gas exchange andconsequent effects on photosynthesis and productivity.

FIGURE 8.1 Stomata. (A) Elliptic type in the lower epi-dermis of Zebrina. In this picture the stoma is open(× 250). (B) Graminaceous type from the adaxial sur-face of maize (Zea mays) leaf. These stomata are closed(× 250).

More than 90 percent of the CO2 and water vaporexchanged between a plant and its environment passesthrough the stomata. Stomata are therefore involved incontrolling two very important but competing processes:uptake of CO2 for photosynthesis and, as discussed inChapter 2, transpirational water loss. It is important,therefore, to take into account stomatal function whenconsidering photosynthetic productivity and crop yields.

8.1 Stomatal Complex Controls Leaf Gas Exchange and Water Loss 131

More recently, additional interest in stomatal functionhas been prompted by recognition that airborne pollu-tants such as ozone (O3) and sulphur dioxide (SO2) alsoenter the leaf through open stomata.

Stomata are found in the leaves of virtually allhigher plants (angiosperms and gymnosperms) and mostlower plants (mosses and ferns) with the exceptionof submerged aquatic plants and the liverworts. Inangiosperms and gymnosperms they are found on mostaerial parts including nonleafy structures such as floralparts and stems, although they may be nonfunctional insome cases. The frequency and distribution of stomatais quite variable and depends on a number of factorsincluding species, leaf position, ploidy level (the num-ber of chromosome sets), and growth conditions. Afrequency in the range of 20 to 400 stomata mm−2

of leaf surface is representative, although frequenciesof 1000 mm−2 or more have been reported. Althoughthere are exceptions to every rule, the leaves of herba-ceous monocots such as grasses usually contain stomataon both the adaxial (upper) and abaxial (lower) sur-faces with roughly equal frequencies. Stomata occur onboth the upper and lower surfaces of herbaceous dicots’leaves, but the frequency is usually lower on the uppersurface. Most woody dicots and tree species have stom-ata only on the lower leaf surface while floating leaves ofaquatic plants (e.g., water lily) have stomata only on theupper surface. In most cases the stomata are randomlyscattered across the leaf surface, although in monocotswith parallel-veined leaves the stomata are arranged inlinear arrays between the veins.

The most striking feature of the stomatal complexis the pair of guard cells that border the pore. Thesespecialized epidermal cells have the capacity to undergoreversible turgor changes that in turn regulate the size

of the aperture between them. When the guard cellsare fully turgid the aperture is open, and when flaccid,the aperture is closed. While there are many variationson the theme, anatomically we recognize two basic typesof guard cells: the graminaceous type and the elliptictype (Figure 8.1).

Elliptic or kidney-shaped guard cells are so calledbecause of the elliptic shape of the opening. In surfaceview, these guard cells resemble a pair of kidney beanswith their concave sides opposed. In cross-section thecells are roughly circular in shape, with a ventral wallbordering the pit and a dorsal wall adjacent to thesurrounding epidermal cells (Figure 8.2). The matureguard cell has characteristic wall thickenings, mainlyalong the outer and inner margins of the ventral wall.These thickenings extend into one or two ledges thatprotect the throat of the stoma. In some plants, partic-ularly the gymnosperms and aquatic species, the innerledge may be small or absent. The outer ledge appearsto be an architectural adaptation that helps to preventthe penetration of liquid water from the outside intothe substomatal air space, which would otherwise havedisastrous consequences for gas exchange.

The graminaceous type of guard cell is largelyrestricted to members of the Gramineae and certainother monocots (e.g., palms). Often described asdumbbell-shaped, the graminaceous-type guard cellshave thin-walled, bulbous ends that contain most ofthe cell organelles (Figure 8.1). The ‘‘handle’’ of thedumbbell is characterized by walls thickened toward thelumen. The pore in this case is typically an elongatedslit. The guard cells are flanked by two prominentsubsidiary cells.

FIGURE 8.2 Guard cells seen in cross-section. (From K. Esau, Anatomy of Seed Plants,New York, Wiley, 1977. Reprinted by permission).

132 Chapter 8 / Energy Conservation in Photosynthesis: CO2 Assimilation

8.2 CO2 ENTERS THE LEAFBY DIFFUSION

Diffusion of CO2 into the leaf through the stoma is moreefficient than would be predicted on the basis of stomatalarea alone. A fully open stomatal pore typically measures5 to 15 μm wide and about 20 μm long. The combinedpore area of open stomata thus amounts to no morethan 0.5 to 2 percent of the total area of the leaf. Sinceleaves contain no active pumps, all of the CO2 takeninto the leaf for photosynthesis must enter by diffusionthrough these extremely small pores. One might thinkthat diffusion through such a limited area would beextremely restricted, yet it has been calculated that therate of CO2 uptake by an actively photosynthesizing leafmay approach 70 percent of the rate over an absorbingsurface with an area equivalent to that of the entire leaf!This extraordinarily high diffusive efficiency appears tobe related to the special geometry of gaseous diffusionthrough small pores.

The high efficiency of gaseous diffusion throughstomata can be demonstrated experimentally by mea-suring CO2 diffusion into a container of CO2-absorbingagent such as sodium hydroxide. The container iscovered with a thin membrane perforated with poresof known dimensions. Diffusion of CO2 through themembrane can be measured as the amount of carbon-ate present in the sodium hydroxide solution after, forexample, one hour. It was discovered that the rate ofCO2 diffusion through a perforated membrane varies inproportion to the diameter of the pores, not the area. Howcan these results be reconciled with Fick’s law, whichstates that

Rate of diffusion = v = D · A(dc/dx) (8.1)

where D is the diffusion coefficient, A is the surfacearea over which diffusion occurs, and (dc/dx) is theconcentration gradient over which diffusion occurs?Clearly, the rate of diffusion is directly proportionalto the surface area, A, and the concentration gradient(dc/dx).

The physical explanation for this paradox lies inthe pattern of diffusive flow as the gases enter and exitthe stomatal pore. This is illustrated schematically fora stoma in Figure 8.3. Note that in the aperture itself(i.e., in the throat of the stoma) CO2 molecules can flowonly straight through and diffusion is proportional tothe cross-sectional area of the throat as predicted byFick’s law of diffusion. But when the gas molecules passthrough the aperture into the substomatal cavity, theycan ‘‘spill over’’ the edge of the pore. The additional dif-fusive capacity contributed by spillover is proportionalto the amount of edge, or the perimeter of the pore.Because the area of a pore decreases by the square ofthe radius (r) while perimeter varies directly with the

Guardcell

Guardcell

Epidermis

Outside

Inside

FIGURE 8.3 The spillover effect for diffusion of CO2

through a stomatal pore. The dashed lines are isobars,representing regions of equivalent CO2 partial pressure.

diameter (2r), the relative contribution of the perime-ter effect increases as the pore size decreases. Thus invery small pores (e.g., the size of stomata) the bulk ofgas movement is accounted for by diffusion over theperimeter. Even this effect is exaggerated with respectto stomata. Because of their elliptical shape, the ratio ofperimeter to area is greater than for circular pores.

How is a high concentration gradient for CO2established and maintained? To ensure a constant dif-fusion of CO2 from the air into the leaf, the CO2concentration within the substomatal cavity and leaf airspaces must be less than the CO2 concentration in theair above the leaf. This CO2 concentration gradient(dc/dx) is established because, in the light, chloroplastscontinuously fix CO2, that is, chloroplasts within theleaf mesophyll cells continuously convert gaseous CO2into a stable, nongaseous molecule 3-phosphoglycerate(PGA) through the reductive pentose phosphate cycle(see below). Thus, this biochemical cycle constantlyremoves CO2 from intercellular air spaces of a leaf,thereby ensuring that the internal leaf CO2 concen-tration is less than the ambient CO2 concentrations inthe light. In the dark, photosynthesis stops but respi-ration generates CO2 such that the internal leaf CO2concentrations are greater than the ambient CO2 con-centrations, and thus CO2 diffuses out of a leaf in thedark. The rate of CO2 evolution from a leaf in the darkis a measure of the rate of leaf mitochondrial respiration.

The above arguments, of course, represent an idealsituation. In reality, the stomatal pore itself is not theonly barrier to gaseous diffusion between the leaf andits environment. A number of other factors—such asunstirred air layers on the leaf surface and the aqueouspath between the air space and the chloroplast—offerresistance to the uptake of CO2 into the leaf and com-plicate the actual situation. Nonetheless, stomata areremarkably efficient structures. They permit very high

8.3 How Do Stomata Open and Close? 133

rates of CO2 absorption, without which photosynthesiswould be severely limited. This creates a paradox. Asystem that is efficient for the uptake of CO2 is alsoefficient for the loss of water vapor from the internalsurfaces of the leaf (Chapter 2). Thus, the principalfunctional advantage offered by the stomatal apparatusis an ability to conserve water by closing the pore whenCO2 is not required for photosynthesis or when waterstress overrides the leaf’s photosynthetic needs.

8.3 HOW DO STOMATA OPENAND CLOSE?

This question may be answered by first asking whatmechanical forces are involved in guard cell movement.The driving force for stomatal opening is known to bethe osmotic uptake of water by the guard cells and theconsequent increase in hydrostatic pressure. The resultis a deformation of the opposing cells that increases thesize of the opening between them. In the case of ellipticguard cells the thickened walls become concave, whilein the dumbbell-shaped cells the handles separate butremain parallel. Stomatal closure follows a loss of water,and the consequent decrease in hydrostatic pressure andrelaxation of the guard cell walls.

Deformation of elliptic guard cells during openingis due to the unique structural arrangement of the guardcell walls. In normal cells, bands of cellulose microfibrilsencircle the cell at right angles to the long axis of the cell.Studies with polarized light and electron microscopyhave demonstrated that the microfibrils in the guard cellwalls are oriented in radial fashion, fanning out fromthe central region of the ventral wall (Figure 8.4). Addi-tional microfibrils are arranged longitudinally withinthe ventral wall thickenings, crosslinking with the radialbands and restricting expansion along the ventral wall.When the guard cells take up water, expansion followsthe path of least resistance—which is to push the rel-atively thin dorsal walls outward into the neighboringepidermal cells. This causes the cells to arch along theventral surface and form the stomatal opening. Thedumbbell-shaped guard cells of the grasses also dependon the osmotic uptake of water, but operate in a slightlydifferent way. In this case the bulbous ends of the cellspush against each other as they swell, driving the centralhandles apart in parallel and widening the pore betweenthem.

What controls stomatal opening and closure? Toanswer this question it is necessary instead to ask whatregulates the osmotic properties of the guard cells. Thisquestion has proven difficult to answer, partly becauseso many factors seem to be involved and partly because ithas been difficult to study guard cell metabolism free ofcomplications introduced by the surrounding epidermaland mesophyll cells. This problem has been partially

FIGURE 8.4 The role of microfibrils in guard cell move-ment. The orientation of the microfibrils (solid arrows)in elliptic guard cells allows expansion of the cells only inthe direction shown by the dashed arrows. This causesthe cells to buckle and thereby increase the size of theopening between their adjacent walls.

resolved by studying guard cell behavior in peeled stripsof epidermal cells. More recently, techniques for prepa-ration of guard cell protoplasts have become available,making it possible to study guard cell metabolism andion movement in isolation.

Over the years a variety of mechanisms have beenoffered to explain changing osmotic concentrationsof guard cells. Most have centered on the observa-tion that guard cells normally contain chloroplastsand were assumed to be photosynthetically competent.One way or another it was proposed that an accu-mulation of photosynthetic product—sugars and othersmall molecules—contributed directly to the observedosmotic changes in the guard cells. While it is truethat most guard cells do have chloroplasts, the numberof chloroplasts varies considerably. As well, the guardcells of some species (e.g., some orchids and variegatedregions of Pelargonium) have no chloroplasts but remainfully functional. Furthermore, investigators have beenunable to detect significant levels of Rubisco (the princi-pal carbon-fixing enzyme; see below) in the guard cellsof at least 20 species, leading to the conclusion thatthe carbon-fixing portion of photosynthesis does notoperate in guard cells. The conclusion is inescapable:photosynthetic carbon metabolism cannot be invoked asa general mechanism to explain guard cell movement.

In the late 1960s it became evident that K+ levelsare very high in open guard cells and very low in closedguard cells (Table 8.1). A variety of techniques, includ-ing electron microprobes and histochemical methodsspecific for K+, have confirmed that the K+ contentof closed guard cells is low compared with that of thesurrounding subsidiary and epidermal cells. Upon open-ing, large amounts of K+ move from the subsidiary and

134 Chapter 8 / Energy Conservation in Photosynthesis: CO2 Assimilation

TABLE 8.1 Potassium content of open and closed guard cells.

K+ Content

pmol/Guard Cell mM

Species Open Closed Open Closed

Vicia faba 2.72 0.55 552 112Commelina communis 3.1 0.4 448 95

Data from MacRobbie, 1987.

epidermal cells into the guard cells. Consequently, anaccumulation of K+ in guard cells is now accepted as auniversal process in stomatal opening. This work gaverise to the current hypothesis that the osmotic poten-tial of guard cells and, consequently, the size of thestomatal opening, is determined by the extent of K+accumulation in the guard cells.

Although we lack a thorough understanding of themechanisms involved, available information about guardcell metabolism and stomatal movements is summarizedin the general model shown in Figure 8.5. It is widelyaccepted that accumulation of ions by most plant cellsis driven by an ATP-powered proton pump locatedon the plasma membrane (Chapter 3). Two lines ofevidence indicate that K+ uptake by stomatal guardcells fits this general mechanism. First, the fungal toxinfusicoccin, which is known to stimulate active protonextrusion by the pump, stimulates stomatal opening.Second, vanadate (VO−

3 ), which inhibits the protonpump, also inhibits stomatal opening. This constitutes

H+ H+

2H+

K+ K+

CI– CI−

CO2 Malate 2−

PEP

ATP

PEPcase

Starch

Vacuole

FIGURE 8.5 A simplified model for ion flow associatedwith the guard cells during stomatal opening. Potas-sium uptake is driven by an ATPase-proton pump ofions located in the plasma membrane. The accumula-tion of ions in the vacuole lowers the water potential ofthe guard cell, thereby stimulating the osmotic uptake ofwater and increased turgor.

reasonably good evidence that proton extrusion is oneof the initial events in stomatal opening. By removingpositively charged ions, proton extrusion would tendto hyperpolarize the plasma membrane (i.e., lower theelectrical potential inside the cell relative to the outside)as well as establish a pH gradient. Hyperpolarization isthought to open K+ channels in the membrane, whichthen allows the passive uptake of K+ in response tothe potential difference or charge gradient across themembrane.

In order to maintain electrical neutrality, excess K+ion accumulated in the cells must be balanced by acounterion carrying a negative charge. According to themodel shown in Figure 8.5, charge balance is achievedpartly by balancing K+ uptake against proton extrusion,partly by an influx of chloride ion (Cl−), and partlyby production within the cell of organic anions suchas malate. In most species, malate production probablyaccounts for the bulk of the required counterion whilein others, such as corn (Zea mays), as much as 40 per-cent of the K+ moving into the cell is accompaniedby Cl−. In those few species whose guard cells lackchloroplast or starch, Cl− is probably the predominantcounterion.

In addition to its role in maintaining charge bal-ance, the accumulation of malate also helps to main-tain cellular pH during solute accumulation. Protonextrusion would tend to deplete the intracellular pro-ton concentration and increase cellular pH. However,because malate is an organic anion, each carboxyl group(—COO−) accumulated releases one proton into thecytosol. The synthesis of malate therefore tends toreplenish the supply of protons lost by extrusion andmaintain cellular pH at normal levels.

The evidence for malate as a counterion is quitestrong. To begin with, malate levels in guard cells ofopen stomata are five to six times that of closed stomata.Second, guard cells contain high levels of the enzymephosphoenolpyruvate carboxylase (PEPcase), which cat-alyzes the formation of malate (Figure 8.5). Third, thereis a decrease in the starch content of open stomata thatcorrelates with the amount of malate formed. Finally,factors that influence stomatal opening and closure alsoinfluence the activity of PEPcase. For example, fusic-occin, which induces stomatal opening, also causes an

8.4 Stomatal Movements are also Controlled by External Environmental Factors 135

increase in both malate concentration and the activity ofPEPcase. Conversely, the plant hormone abscisic acid,which normally induces stomatal closure, antagonizesthe effect of fusicoccin. The effect of fusicoccin is tostimulate the phosphorylation of PEPcase, a processwell known to activate a variety of enzymes and otherproteins in the cell.

The accumulation of K+, Cl−, and malate in thevacuoles of the guard cells would lower both the osmoticpotential and the water potential of the guard cells. Theconsequent uptake of water would increase the turgorand cause the stomata to open. At present, this remainsa working model for stomatal opening since many of thedetails have yet to be verified experimentally.

Stomatal closure has not received the same atten-tion that opening has, but it is generally assumed thatclosure is effected by a simple reversal of the eventsleading to opening. On the other hand, the rate of clo-sure is often too rapid to be accounted for simply bya passive leakage of ions from the guard cells, leadingto the suggestion that other specific metabolic pumpsare responsible for actively extruding ions upon closure.One possibility is that signals for stomatal closure stim-ulate the uptake of Ca2+ into the cytosol. Ca2+ uptakewould depolarize the membrane, thus initiating a chainof events that includes opening anion channels to allowthe release of Cl− and malate. According to this scenario,a loss of anions would further depolarize the membrane,opening K+ channels and allowing the passive diffu-sion of K+ into the adjacent subsidiary and epidermalcells.

What is the source of ATP that powers the guardcell proton pumps? The two most logical sources wouldbe either photosynthesis in the guard cell chloroplastsor cellular respiration. Although most guard cells docontain chloroplasts, they are generally smaller, lessabundant, and with fewer thylakoids than those ofunderlying mesophyll cells. As noted above, guardcell chloroplasts apparently lack the enzymatic machin-ery for photosynthetic carbon fixation. On the otherhand, although ATP production has not been measureddirectly, indirect evidence indicates that they are capa-ble of using light energy to produce ATP, a processknown as photophosphorylation (see Chapters 5 and7). Photosynthesis is probably not the only immediatesource of energy, however, since stomatal movementcan occur in the dark. An alternative source of energyis cellular respiration. Guard cells do have large num-bers of mitochondria and high levels of respiratoryenzymes. They may well be able to derive sufficientATP from the oxidation of carbon through oxida-tive phosphosphorylation (see Chapters 5 and 10).It appears that guard cells have more than adequatecapacity to produce, through either respiration or pho-tosynthesis, all the energy necessary to drive stomatalopening.

8.4 STOMATAL MOVEMENTSARE ALSO CONTROLLEDBY EXTERNALENVIRONMENTAL FACTORS

The major role of stomata is to allow entry of CO2into the leaf for photosynthesis while at the same timepreventing excessive water loss. In this sense, they evi-dently serve a homeostatic function; they operate tomaintain a constancy of the internal environment ofthe leaf. It should come as no surprise, then, to findthat stomatal movement is regulated by a variety ofenvironmental and internal factors such as light, CO2levels, water status of the plant, and temperature. Itmight be expected, for example, that stomata will openin the light in order to admit CO2 for photosynthesis orpartially close when CO2 levels are high in order to con-serve water while allowing photosynthesis to continue.On the other hand, conditions of extreme water stressshould override the plant’s immediate photosyntheticneeds and lead to closure, protecting the leaf againstthe potentially more damaging effects of desiccation. Ingeneral, these expectations have been verified by directobservation. Each of these factors can theoretically bestudied independently under the controlled conditionsof the laboratory, but the extent to which they interactunder natural conditions makes it far more difficult tostudy the effects of one relative to another. Moreover,it must be kept in mind that stomatal opening is not anall-or-none phenomenon. At any given time, the extentof stomatal opening and its impact on both photosyn-thesis and water loss will be determined by the sum ofall of these factors and not by any one alone.

8.4.1 LIGHT AND CARBON DIOXIDEREGULATE STOMATAL OPENING

Both light and CO2 appear to make a substantial contri-bution to the daily cycle of stomatal movements. Theireffects are also tightly coupled, which makes it verydifficult to distinguish their relative contributions. Ingeneral, low CO2 concentrations and light stimulateopening while high CO2 concentrations cause rapidclosure even in the light. The response of the stomata isto the intracellular concentration of CO2 in the guard cells.Recall that the outer surfaces of the epidermis, includingthe guard cells, are covered with the CO2-impermeablecuticle. Once induced to close by high CO2 treatment,stomata are not easily forced to open by treatment withCO2-free air. This is because the closed guard cellsremain in equilibrium with the high CO2 content of theair trapped in the substomatal chamber. Consequently,it is the CO2 content of the substomatal chamber ratherthan the ambient atmosphere that is most important in

136 Chapter 8 / Energy Conservation in Photosynthesis: CO2 Assimilation

regulating stomatal opening. The actual mechanism bywhich CO2 regulates stomatal opening is not under-stood.

Stomata normally open at dawn. As well, stomataclosed by exposure to high CO2 can be induced to openslowly if placed in the light. Both responses appear toresult from two separate effects of light; one indirect andone direct. The indirect effect requires relatively highfluence rates and is usually attributed to a reductionin intercellular CO2 levels due to photosynthesis inthe mesophyll cells. By the same argument, closureof the guard cells in the dark can be attributed tothe accumulation of respiratory CO2 inside the leaf.This interpretation is reinforced by the observationthat the action spectrum for moderate to high fluencerates resembles that for photosynthesis with peaks inboth the red and blue. Thus it appears that CO2 isa primary trigger and that, at least in intact leaves, theindirect effect of light may operate through regulation ofintercellular CO2 levels. A significant difficulty with thisinterpretation, however, is that similar action spectrahave been obtained for isolated epidermal peels. Sucha result in the absence of an intact leaf argues stronglyfor an important but yet undefined role of the guard cellchloroplasts.

Perhaps one of the more significant advances toemerge in recent years is the unequivocal demonstra-tion of a direct effect of low-fluence blue light onstomatal opening. If the stomata depended solely onphotosynthetically active light, it would likely sufferfrom two limitations. First, the guard cells would beunable to respond to light levels below the photosyn-thetic light compensation point (i.e., the minimumfluence rate at which photosynthesis exceeds respira-tion). Second, the system would be prone to extremeoscillations as the rate of photosynthesis fluctuated withrapid changes in PAR. A direct effect of blue lighton stomatal opening would seem to circumvent theselimitations.

The blue light effect has been demonstrated in avariety of ways. Although stomatal opening is promotedby both red and blue light, it is generally more sensitiveto blue light than to red. At low fluence rates, below 15μmol m−2 s−1, blue light will cause stomatal opening butred light is ineffective. At higher fluence rates stomatalopening under blue light (which presumably activatesboth systems) is consistently higher than under red atthe same fluence rate. The response of stomata to redlight is probably indirect, mediated by the guard cellchloroplasts and involving photosynthetic ATP produc-tion. The action spectrum of the blue light response, onthe other hand, is typical of other blue light responsesand is probably mediated by cryptochrome, a putativeblue light receptor (Chapter 6). The mode of actionof blue light is not certain, but blue light does causeswelling of isolated guard cell protoplasts. This result

indicates that blue light acts directly on the guard cells.Several investigators have reported that blue light acti-vates proton extrusion by the guard cells and stimulatesmalate biosynthesis; both are prerequisites to stomatalopening.

But what function does the blue light response serveunder natural conditions? One interesting and plausiblesuggestion is that it may have a role in the early morn-ing opening of stomata. Opening can often be observedbefore sunrise, when fluence rates are much lower thanthat required to drive photosynthesis. They may alsoremain open after sunset. The high sensitivity of theblue light response to low fluence rates together withthe relatively high proportion of blue light in sunlightat dawn and dusk suggests that the blue light responsecould function as an effective ‘‘light-on’’ signal. Froman ecophysiological standpoint, the blue light responseanticipates the need for atmospheric CO2 and drivesstomatal opening in preparation for active photosynthe-sis. Another possible role is to stimulate rapid stomatalopening in response to sunflecks—the sunfleck itselfwould be analogous to a blue light pulse—in order tomaximize the opportunity for photosynthesis under thisparticular condition (Chapter 14).

8.4.2 STOMATAL MOVEMENTSFOLLOW ENDOGENOUSRHYTHMS

Many biological processes undergo periodic fluctuationsthat persist under constant environmental conditions.This phenomenon, known as endogenous rhythm,is discussed further in Chapter 25. It was demon-strated that stomatal opening and closure in Tradescantialeaves persisted for at least three days, even thoughthe plants were maintained under continuous light. Aperiodicity of approximately 24 hours was maintained,although the timing of opening or closure could beshifted by a six-hour dark period. Results such as theseclearly indicate an involvement of an endogenous circa-dian rhythm in control of stomatal opening, althoughit is not clear how the rhythm interacts with otherstimuli.

8.5 THE PHOTOSYNTHETICCARBON REDUCTION (PCR)CYCLE

Now that we understand the processes involved in thecontrol of CO2 entry into a leaf, we will examinein some detail the biochemical mechanisms by whichchloroplasts fix this CO2 and convert it to stable phos-phorylated carbon intermediates.

8.5 The Photosynthetic Carbon Reduction (PCR) Cycle 137

ATPNADPH

Triose-P

RuBP PGA

CO2

ReductionRegeneration

ATP

Carboxylation

FIGURE 8.6 The three stages of the photosynthetic car-bon reduction cycle.

8.5.1 THE PCR CYCLE REDUCES CO2TO PRODUCE A THREE-CARBONSUGAR

The pathway by which all photosynthetic eukaryoticorganisms ultimately incorporate CO2 into carbohy-drate is known as carbon fixation or the photosyntheticcarbon reduction (PCR) cycle. It is also referred toas the Calvin cycle, in honor of Melvin Calvin, whodirected the research effort that elucidated the pathway.Mapping the complex sequence of reactions involvingthe formation of organic carbon and its conversion tocomplex carbohydrates represented a major advance inplant biochemistry. For his efforts and those of his asso-ciates, Calvin was awarded the Nobel Prize for chemistryin 1961.

The PCR cycle can be divided into three pri-mary stages (Figure 8.6): (1) carboxylation which fixesthe CO2 in the presence of the five-carbon acceptormolecule, ribulose bisphosphate (RuBP), and convertsit into two molecules of a three-carbon acid; (2) reduc-tion, which consumes the ATP and NADPH producedby photosynthetic electron transport to convert the

FIGURE 8.7 The carboxylation reaction ofthe photosynthetic carbon reduction cycle.

three-carbon acid to triose phosphate; and (3) regener-ation, which consumes additional ATP to convert someof the triose phosphate back into RuBP to ensure thecapacity for the continuous fixation of CO2.

8.5.2 THE CARBOXYLATION REACTIONFIXES THE CO2

Calvin’s strategy for unraveling the path of carbonin photosynthesis was conceptually very straightfor-ward: identify the first stable organic product formedfollowing uptake of radiolabeled CO2. In order toachieve this, cultures of the photosynthetic green algaChlorella were first allowed to establish a steady rate ofphotosynthesis. 14CO2 was then introduced and photo-synthesis continued for various periods of times beforethe cells were dropped rapidly into boiling methanol.The hot methanol served two functions: it denatured theenzymes, thus preventing any further metabolism, whileat the same time extracting the sugars for subsequentchromatographic analysis. When the time of photosyn-thesis in the presence of 14CO2 was reduced to as littleas two seconds, most of the radioactivity was found in athree-carbon acid, 3-phosphoglycerate (3-PGA). Thus3-PGA appeared to be the first stable product of photo-synthesis. Other sugars that accumulated the label laterin time were probably derived from 3-PGA. BecauseCalvin’s group determined that the first product wasa three-carbon molecule, the PCR cycle is commonlyreferred to as the C3 cycle. The next step was todetermine what molecule served as the acceptor—themolecule to which CO2 was added in order to makethe three-carbon product. Systematic degradation of3-PGA demonstrated that the 14C label was predomi-nantly in the carboxyl carbon. A two-carbon acceptormolecule would be logical, but the search was long andfutile. No two-carbon molecule could be found. Instead,Calvin recognized that the acceptor was the five-carbonketo sugar, ribulose-1,5-bisphosphate (RuBP). Thisturned out to be the key to the entire puzzle. The reac-tion is a carboxylation in which CO2 is added to RuBP,forming a six-carbon intermediate (Figure 8.7). The

138 Chapter 8 / Energy Conservation in Photosynthesis: CO2 Assimilation

intermediate, which is transient and unstable, remainsbound to the enzyme and is quickly hydrolyzed totwo molecules of 3-PGA. The carboxylation reactionis catalyzed by the enzyme ribulose-1,5-bisphosphatecarboxylase-oxygenase, or Rubisco. Rubisco is with-out doubt the most abundant protein in the world,accounting for approximately 50 percent of the solubleprotein in most leaves. The enzyme also has a high affin-ity for CO2 that, together with its high concentrationin the chloroplast stroma, ensures rapid carboxylation atthe normally low atmospheric concentrations of CO2.Thus, the reaction catalyzed by Rubisco maintains theCO2 concentration gradient (dc/dx) between the inter-nal air spaces of a leaf and the ambient air to ensure aconstant supply of this substrate for the PCR cycle.

8.5.3 ATP AND NADPH ARE CONSUMEDIN THE PCR CYCLE

The carboxylation reaction, with a �G of −35 kJ mol−1,is energetically very favorable. This poses an interestingquestion. If the equilibrium constant of the reactionfavors carboxylation with such a high negative freeenergy change, where is the need for an input of energyfrom the light reactions of photosynthesis? Energy isrequired at two points: first for the reduction of 3-PGAand second for regeneration of the RuBP acceptormolecule. Each of these requirements will be discussedin turn.

8.5.3.1 Reduction of 3-PGA In order for the chlo-roplast to continue to take up CO2, two conditions mustbe met. First, the product molecules (3-PGA) must becontinually removed and, second, provisions must bemade to maintain an adequate supply of the acceptormolecule (RuBP). Both require energy in the form ofATP and NADPH.

The 3-PGA is removed by reduction to the triosephosphate, glyceraldehyde-3-phosphate. This is atwo-step reaction (Figure 8.8) in which the 3-PGA isfirst phosphorylated to 1,3-bisphosphoglycerate, whichis then reduced to glyceraldehyde-3-phosphate (G3P).Both the ATP and the NADPH required in these two

steps are products of the light reactions and togetherrepresent one of two sites of energy input. The resultingtriose sugar-phosphate, G3P, is available for export tothe cytoplasm, probably after conversion to dihydroxy-acetone phosphate (DHAP) (Chapter 9).

8.5.3.2 Regeneration of RuBP In order to main-tain the process of CO2 reduction, it is necessary toensure a continuing supply of the acceptor molecule,RuBP. This is accomplished by a series of reactionsinvolving 4-, 5-, 6-, and 7-carbon sugars (Figures 8.9,8.10). These reactions include the condensation of a6-carbon fructose-phosphate with a triose-phosphate toform a 5-carbon sugar and a 4-carbon sugar. Anothertriose joins with the 4-carbon sugar to produce a7-carbon sugar. When the 7-carbon sugar is com-bined with a third triose-phosphate, the result is twomore 5-carbon sugars. All of the five-carbon sugar canbe isomerized to form ribulose-5-phosphate (Ru5P).Ru5P can, in turn, be phosphorylated to regenerate therequired ribulose-1,5-bisphosphate.

The net effect of these reactions is to recycle thecarbon from five out of every six G3P molecules, thusregenerating three RuBP molecules to replace thoseused in the earlier carboxylation reactions. The summaryreactions shown in Figures 8.9 and 8.10 include threemolecules of RuBP on each side of the equation. Thisis to emphasize that the cycle serves to regenerate theoriginal number of acceptor molecules and maintain asteady-state carbon reduction. Figures 8.9 and 8.10 showthat for every three turns of the cycle (i.e., the uptakeof three CO2) there is sufficient carbon to regeneratethe required number of acceptor molecules plus oneadditional triose phosphate, which is available for exportfrom the chloroplast. The stoichiometry in Figures 8.9and 8.10 was chosen to illustrate this point. Six turnsof the cycle would regenerate 6 molecules of RuBP,leaving the equivalent of one additional hexose sugar asnet product. Twelve turns would generate the equivalentof a sucrose molecule, and so on.

As a general rule it is necessary to show that therequired enzymes are present and active before a com-plex metabolic scheme can be accepted as fact. Calvin’s

FIGURE 8.8 Reduction of phosphoglyceric acid (PGA) to glyceraldehyde-3-phosphate (G3P).

8.6 The PCR Cycle is Highly Regulated 139

FIGURE 8.9 The photosynthetic carbon reduction (PCR) cycle. Numbers inbrackets indicate stoichiometry. Enzymes, indicated by circled numbers are: (1)ribulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco); (2) 3-phosphoglyceratekinase and glyceraldehyde-3-phosphate dehydrogenase; (3) aldolase; (4) fructose-1,6-bisphosphatase; (5) transketolase; (6) aldolase; (7) sedoheptulose-1,7- bisphos-phatase; (8, 9) ribulose-5-phosphate epimerase; (10) ribose-5-phosphate isomerase;(11) ribulose-5-phosphate kinase.

PCR cycle has met this criterion since all of the enzymesrequired by the scheme in Figure 8.9 have now beendemonstrated in the stroma. Moreover, all of the reac-tions have been demonstrated in vitro, at rates that wouldsupport maximal rates of photosynthesis.

8.5.4 WHAT ARE THE ENERGETICSOF THE PCR CYCLE?

Figure 8.9 shows that for three turns of the cycle,that is, the uptake of 3 molecules of CO2, a total of6 molecules of NADPH and 9 molecules of ATP arerequired. Therefore, the reduction of each molecule ofCO2 requires 2 molecules of NADPH and 3 moleculesof ATP for a ratio of ATP/NADPH of 3/2 or 1.5.Since each NADPH stores 2 electrons, we can see thata total of 4 electrons are required to fix each moleculeof CO2. This total represents an energy input of 529 kJmol−1 of CO2. Oxidation of one mole of hexose wouldyield about 2817 kJ, or 469 kJ mol−1 of CO2. Thus, thephotosynthetic reduction process represents an energystorage efficiency of about 88 percent. If we includethe energy consumed in the form of the three ATP

per CO2 (3 × 31.4 kJ mol−1 = 282 kJ mol−1) for theregeneration of RuBP, then energy storage efficiency isabout 58 percent. An important assumption underlyingthese simple calculations is that all of the CO2 fixed bythe PCR cycle actually remains fixed in the leaf. Laterin this chapter we will see that this assumption does notnecessarily hold under all conditions.

8.6 THE PCR CYCLE IS HIGHLYREGULATED

It was originally believed that the PCR cycle did notrequire a significant level of regulation, in part becauseearly in vitro studies of Rubisco suggested a low, andprobably rate-limiting, reactivity for this critical enzyme.(Its in vivo reactivity is now known to be much higher,although it may still be rate limiting.) In addition, plantswere widely believed to be opportunistic and would useavailable light, water, and CO2 to conduct photosyn-thesis at maximum rates. However, it is now recognizedthat photosynthesis does not operate in isolation and an

140 Chapter 8 / Energy Conservation in Photosynthesis: CO2 Assimilation

FIGURE 8.10 Summary reactions of the PCR cycle.Three turns of the cycle result in the regeneration of3 molecules of the acceptor ribulose-1,5-bisphosphate(RuBP) plus an additional molecule of glyceraldehyde-3-phosphate (G-3-P). Additional abbreviations are: PGA,3-phosphoglyceric acid; FBP, fructose-1,6-bisphosphate;F6P, fructose-6-phosphate; E-4-P, erythrose-4-phosphate; XuP, xylulose-5-phosphate; SBP,sedoheptulose-1,7-bisphosphate; R-5-P, ribulose-5-phosphate.

unregulated photosynthetic machinery is incompatiblewith an orderly and integrated metabolism. Chang-ing levels of intermediates between light and darkperiods and competing demands for light energy andcarbon with other cellular needs (nitrate reduction, forexample) demand some degree of regulation. The mosteffective control is, of course, at the level of enzymeactivities. Molecular biology combined with classicalenzyme kinetics (see Box 8.1) and structural informa-tion obtained through protein crystallization has begunto elucidate the sophisticated nature of photosyntheticenzyme regulation. A principal factor in the regula-tion of the PCR cycle is, perhaps not surprisingly,light.

8.6.1 THE REGENERATION OF RuBPIS AUTOCATALYTIC

The rate of carbon reduction is partly dependent on theavailability of an adequate pool of acceptor molecules,CO2 and RuBP. The PCR cycle can utilize newly fixedcarbon to increase the size of this pool, when neces-sary, through the autocatalytic regeneration of RuBP.During the night, when photosynthesis is shut downand carbon is required for other metabolic activities, theconcentrations of intermediates in the cycle (includingRuBP) will fall to low levels. Consequently, when pho-tosynthesis starts up again, the rate could be severelylimited by the availability of RuBP, the CO2 acceptormolecule. Normally the extra carbon taken in through

P

P

P

15 CO2

18 RuBP15 RuBP

30 PGA

SugarStarch

30 Triose −

25 Triose −

5 Triose −

FIGURE 8.11 Autocatalytic properties of PCR cycle.When required, carbon can be retained within the PCRcycle (dashed arrows) to build up the amount of receptormolecules and increase the rate of photosynthesis.

the PCR cycle is accumulated as starch or exported fromthe chloroplast. However, the PCR cycle has the poten-tial to augment supplies of acceptor by retaining thatextra carbon and diverting it toward generating increas-ing amounts of RuBP instead (Figure 8.11). In this waythe amount of acceptor can be quickly built up withinthe chloroplast to the level needed to support rapidphotosynthesis. Only after the level of RuBP has beenbuilt up to adequate levels will carbon be withdrawn forstorage or export. The time required to build up thenecessary levels of PCR cycle intermediates in the tran-sition from dark to light is called the photosyntheticinduction time. No other sequence of photosyntheticreactions has this capacity, which may help to explainwhy all photosynthetic organisms ultimately rely onthe C3 cycle for carbon reduction. How autocatalysisis regulated is not altogether clear. However, the mosteffective control would be to enhance the activities ofenzymes favoring recycling over those leading to starchsynthesis or export of product.

8.6.2 RUBISCO ACTIVITY ISREGULATED INDIRECTLYBY LIGHT

Rubisco activity declines rapidly to zero when the lightis turned off and is regained only slowly when the light isonce again turned on. Light activation is apparently indi-rect and involves complex interactions between Mg2+fluxes across the thylakoid, CO2 activation, chloroplastpH changes, and an activating protein.

As noted in the previous chapter, light-driven elec-tron transport leads to a net movement of protons intothe lumen of the thylakoids. The movement of protons

8.6 The PCR Cycle is Highly Regulated 141

across the thylakoid membrane generates a proton gra-dient equivalent to 3.0 pH units and an increase in thepH of the stroma from around pH 5.0 in the dark toabout pH 8.0 in the light. In vitro, Rubisco is gener-ally more active at pH 8.0 than at pH 5.0. The Mg2+requirement for Rubisco activity was noted some yearsago. Light also brings about an increase in the free Mg2+of the stroma as it moves out of the lumen to compensatefor the proton flux in the opposite direction.

Work in the laboratory of G. H. Lorimer, againusing isolated Rubisco in vitro, has shown that Rubiscouses CO2 not only as a substrate but also as an activator.The activating CO2 must bind to an activating site,called the allosteric site, that is separate and distinct fromthe substrate-binding site (see Box 8.1). Based on thesein vitro studies, Lorimer and Miziorko proposed a modelfor in vivo activation that takes into account all threefactors: CO2, Mg2+, and pH. According to this model,the CO2 first reacts with an ε-amino group of a lysineresidue in the allosteric site, forming what is known as acarbamate (Figure 8.12). Carbamate formation requiresthe release of two protons and, consequently, wouldbe favored by increasing pH. The Mg2+ then becomescoordinated to the carbamate to form a carbamate-Mg2+complex, which is the active form of the enzyme.

Further experiments, however, indicated that the invitro model could not fully account for the activationof Rubisco in leaves. In particular, measured values forin vivo Mg2+ and CO2 concentrations and pH differ-ences were not sufficient to account for more than halfthe expected activation level. This paradox was resolvedby the discovery of an Arabidopsis mutant that failed toactivate Rubisco in the light, even though the enzymeisolated from the mutant was apparently identical tothat isolated from the wildtype. Electrophoretic analysisrevealed that the rca mutant, as it was called, was missinga soluble chloroplast protein. Subsequent experimentsdemonstrated that full activation of Rubisco could be

hν

H+

H+ lumenpH = 5.0

Stroma pH = 8.0

Mg2+

Mg2+

Mg2+

COO−

NH

COO−

NHNH3

CO2

2H+

RubiscoRubiscoRubisco

FIGURE 8.12 Light-driven ion fluxes and activation ofRubisco. Activation of Rubisco is facilitated by theincrease in stromal pH and Mg2+ concentration thataccompanies light-driven electron transport.

restored in vitro simply by adding the missing pro-tein to a reaction mixture containing Rubisco, RuBP,and physiological levels of CO2. This protein has beennamed Rubisco activase to signify its role in promotinglight-dependent activation of Rubisco.

Rubisco activase is known to require energy in theform of ATP. The protein has been identified in atleast 10 genera of higher plants as well as the greenalga Chlamydomonas. It is clear that Rubisco activase hasa significant and probably ubiquitous role to play inregulating eukaryotic photosynthesis.

8.6.3 OTHER PCR ENZYMES ARE ALSOREGULATED BY LIGHT

Rubisco is not the only PCR cycle enzyme requiringlight activation. Studies with algal cells, leaves, andisolated chloroplasts have shown that the activities of atleast four other PCR cycle enzymes are also stimulatedby light. These include glyceraldehyde-3-phosphatedehydrogenase (G-3-PDH) (reaction 2, Figure 8.9),fructose-1,6-bisphosphatase (FBPase) (reaction 4,Figure 8.9), sedoheptulose-1,7-bisphosphatase (SBPase)(reaction 7, Figure 8.9), and ribulose-5-phosphatekinase (R-5P-K) (reaction 11, Figure 8.9).

The mechanism for light activation is different fromthat of Rubisco and is best demonstrated in the case ofFBPase. Light activation of FBPase can be blocked bythe electron transport inhibitor DCMU and agents thatselectively modify sulfhydryl groups. On the other hand,the enzyme can be activated in the dark by the reducingagent, dithiothreitol (DTT). It gradually emerged thatactivation requires the participation of both chloroplastferredoxin, a product of the light-dependent reactions,and thioredoxin (Figure 8.13). Like ferredoxin, thiore-doxin is a small (12 kDa) iron-sulphur protein, knownto biochemists for its role in the reduction of ribonu-cleotides to deoxyribonucleotides. It contains two cys-teine residues in close proximity that undergo reversiblereduction–oxidation from the disulphide (—S—S—)

P 700∗

fdred fdox

hν

ThioredoxinS

SThioredoxin

SH

SH

Reduced enzyme(active)

Oxidized enzyme(inactive)

P 700PC

PS I

FIGURE 8.13 The ferredoxin/thioredoxin system forlight-driven enzyme activation.

142 Chapter 8 / Energy Conservation in Photosynthesis: CO2 Assimilation

state to the sulfhydryl (—SH HS—) state. In the chloro-plast, PSI drives the reduction of ferredoxin, which inturn reduces thioredoxin. The reaction is mediated bythe enzyme ferredoxin-thioredoxin reductase. Thiore-doxin subsequently reduces the appropriate disulphidebond on the target enzyme, resulting in its activation.Subsequent deactivation of the enzymes in the dark isnot well understood, but clearly the sulfhydryl groupsare in some way reoxidized and the enzymes renderedinactive.

The traditional view of the PCR cycle was that itdid not require the direct input of light. These reactionswere consequently referred to as the ‘‘dark reactions’’of photosynthesis. In view of the fact that at least fivecritical enzymes in the cycle require light activation,such a designation is clearly not appropriate.

8.7 CHLOROPLASTS OF C3PLANTS ALSO EXHIBITCOMPETING CARBONOXIDATION PROCESSES

The most widely used method for assessing the rateof photosynthesis in whole cells (e.g., algae) or intactplants is to measure gas exchange—either CO2 uptakeor O2 evolution. This is, at best, a complicated pro-cess since there are several different and competingmetabolic reactions that contribute to the gas exchangeof an algal cell or a higher plant leaf. Cellular (or mito-chondrial) respiration (R) is an example of oppositegas exchange, since it results in an evolution of CO2and uptake of O2. Historically, it was assumed thatmitochondrial-based respiration and chloroplast-basedphotosynthesis were effectively independent and thattheir respective contributions to gas exchange couldalso be assessed independently. (One argument heldthat photosynthesis could supply the entire energy needof the leaf directly and the mitochondria would con-sequently ‘‘shut down’’ in the light!). We now knowthat measuring gas exchange is a far less certain pro-cess, complicated in part by oxidative metabolism andthe consequent evolution of CO2 directly associatedwith photosynthetic metabolism (Figure 8.14). Calledphotorespiration (PR), this process involves the reox-idation of products just previously assimilated in pho-tosynthesis. The photorespiratory pathway involves theactivities of at least three different cellular organelles(the chloroplast, the peroxisome, and the mitochon-drion) and, because CO2 is evolved, results in a net lossof carbon from the cell.

The measured CO2 uptake in the light is termedapparent or net photosynthesis (AP), since itrepresents photosynthetic CO2 uptake minus the CO2evolved from mitochondrial respiration plus photores-piration (Equation 8.2). True or gross photosynthesis

PR

R

R

GP

CO2

CO2

CO2

CO2

O2

O2

O2

O2

DarkA.

LightB.

FIGURE 8.14 Gas exchange observed in a C3 leaf in thedark (A) and in the light (B). GP, gross photosynthesis;PR, photorespiration; R, mitochondrial respiration.

(GP) is thus calculated by adding the amount ofmitochondrial-respired CO2 plus photorespired-CO2to that taken up in the light (Equation 8.3).

AP = GP − (R + PR) (8.2)

GP = AP + R + PR (8.3)

Early experiments based on discrimination betweencarbon isotopes suggested there were both qualitativeand quantitative differences between the process of res-piration (i.e., CO2 evolution) as it occurred in the darkand in the light. On this basis, CO2 evolution in the lightwas called photorespiration. Initially, the concept thatlight would alter the rate of respiration was, to say theleast, controversial. However, biochemical and molec-ular evidence has firmly established photorespirationas important process contributing to the gas exchangeproperties of C3 leaves.

8.7.1 RUBISCO CATALYZES THEFIXATION OF BOTH CO2 AND O2

While the legitimacy of photorespiration was beingestablished during the 1960s, the attention of sev-eral investigators was attracted to the synthesis andmetabolism of a two-carbon compound, glycolate. Itgradually emerged that glycolate metabolism was relatedto photorespiration and that the enzymes involved werelocated in peroxisomes and mitochondria as well as thechloroplast. The key to photorespiratory CO2 evolu-tion and glycolate metabolism is the bifunctional natureof Rubisco. In addition to the carboxylation reaction,

8.7 Chloroplasts of C3 Plants also Exhibit Competing Carbon Oxidation Processes 143

FIGURE 8.15 The RuBP oxygenase reaction.

Rubisco also catalyzes an oxygenase reaction, hence thename ribulose-1,5-bisphosphate carboxylase-oxygenase.With the addition of a molecule of oxygen, RuBP is con-verted into one molecule of 3-PGA and one moleculeof phosphoglycolate (Figure 8.15). The phosphoglyco-late is subsequently metabolized in a series of reactionsin the peroxisome and the mitochondrion that result inthe release of a molecule of CO2 and recovery of theremaining carbon by the PCR cycle (Figure 8.16).

The C2 glycolate cycle, also known as the pho-tosynthetic carbon oxidation (PCO) cycle, beginswith the oxidation of RuBP to 3-PGA and P-glycolate.The 3-PGA is available for further metabolism by thePCR cycle, but the P-glycolate is rapidly dephospho-rylated to glycolate in the chloroplast. The glycolate isexported from the chloroplast and diffuses to a perox-isome. Taken up by the peroxisome, the glycolate isoxidized to glyoxylate and hydrogen peroxide. The per-oxide is broken down by catalase and the glyoxylateundergoes a transamination reaction to form the aminoacid glycine. Glycine is then transferred to a mitochon-drion where two molecules of glycine (4 carbons) areconverted to one molecule of serine (3 carbons) plus oneCO2. Glycine is thus the immediate source of photorespiredCO2. The serine then leaves the mitochondrion, return-ing to a peroxisome where the amino group is given upin a transamination reaction and the product, hydrox-ypyruvate, is reduced to glycerate. Finally, glycerate isreturned to the chloroplast where it is phosphorylatedto 3-PGA.

The release of carbon as CO2 during the conver-sion of glycine to serine is accompanied by the releaseof an equivalent amount of nitrogen in the form ofammonia. During active photorespiration, the rate ofammonia release may be substantially greater than therate of nitrogen assimilation. This nitrogen is not lost,however, as the ammonia is rapidly reassimilated in thechloroplast, using the enzymes of the glutamate synthasecycle (Chapter 11).

The C2 glycolate pathway involves complex inter-actions between photosynthesis, photorespiration, andvarious aspects of nitrogen metabolism in at least three

(2) O2

(2) RuBP

PGAADPATP

PCR cycle

Glycerate

Glycerate

(2) P − Glycolate + (2) PGA

(2) Pi

(2) Glycolate

(2) Glycolate

NAD+

NADH

Hydroxy pyruvate (2) Glyoxylate

[−NH2][−NH2]

Serine (2) Glycine

CO2

NH3

(2) GlycineSerine

NAD+NADH

FIGURE 8.16 The photorespiratory glycolate pathway.

different cellular organelles. Much of the supportingevidence comes from labeling studies employing either14CO2 or specific intermediates, or 18O2, in which thefate of the label is followed through the various suspectedchemical transformations. As with the PCR cycle, all ofthe enzymes necessary to carry out the C2 glycolate cyclehave been demonstrated. The distribution of interme-diates between the three organelles, however, is notconclusively established. It is largely inferred from thelocation of the enzymes. All of the subcellular organellesinvolved have been isolated and shown to contain theappropriate enzymes.

8.7.2 WHY PHOTORESPIRATION?

In normal air (21% O2), the rate of photorespirationin sunflower leaves is about 17 percent of gross photo-synthesis. Every photorespired CO2, however, requiresan input of two molecules of O2 (Figure 8.16). Thetrue rate of oxygenation is therefore about 34 percentand the ratio of carboxylation to oxygenation is about3 to 1 (1.00/0.34). This experimental value agrees withsimilar values calculated for several species based on theknown characteristic of purified Rubisco. The ratio ofcarboxylation to oxygenation depends, however, on therelative levels of O2 and CO2 since both gases compete

144 Chapter 8 / Energy Conservation in Photosynthesis: CO2 Assimilation

for binding at the active site on Rubisco. As the concen-tration of O2 declines, the relative level of carboxylationincreases until, at zero O2, photorespiration is also zero.On the other hand, increases in the relative level ofO2 (or decrease in CO2) shifts the balance in favor ofoxygenation. An increase in temperature will also favoroxygenation, since as the temperature increases the sol-ubility of gases in water declines, but O2 solubility isless affected than CO2. Thus O2 will inhibit photosyn-thesis, measured by net CO2 reduction, in plants thatphotorespire. The inhibition of photosynthesis by O2was first recognized by Otto Warburg in the 1920s, but50 years were to pass before the bifunctional nature ofRubisco offered the first satisfactory explanation for thisphenomenon.

There is also an energy cost associated with pho-torespiration and the glycolate pathway. Not only isthe amount of ATP and NAD(P)H expended in theglycolate pathway following oxygenation (5 ATP + 3NADPH) greater than that expended for the reductionof one CO2 in the PCR cycle (3 ATP + 2 NADPH),but there is also a net loss of carbon. On the surface, then,photorespiration appears to be a costly and inefficientprocess with respect to both energy and carbon acquisi-tion. It is logical to ask, as many have, why should theplant indulge in such an apparently wasteful process?

This question is not easily answered, although sev-eral ideas have been put forward. One has it that theoxygenase function of Rubisco is inescapable. Rubiscoevolved at a time when the atmosphere contained largeamounts of CO2 but little oxygen. Under these condi-tions, an inability to discriminate between the two gaseswould have had little significance to the survival of theorganism. Both CO2 and O2 react with the enzyme at thesame active site, and oxygenation requires activation byCO2 just as carboxylation does. It is believed that oxygenbegan to accumulate in the atmosphere primarily due tophotosynthetic activity, but by the time the atmosphericcontent of O2 had increased to significant proportions,the bifunctional nature of the enzyme had been estab-lished without recourse. In a sense, C3 plants were thearchitect of their own problem—generating the oxy-gen that functions as a competitive inhibitor of carbonreduction. By this view, then, the oxygenase functionis an evolutionary ‘‘hangover’’ that has no useful role.However, this is an oversimplified view of photores-piration since photorespiratory mutants of Arabidopsisproved to be lethal under certain growth conditions,indicating the essential nature of the photorespiratorypathway in C3 plants. Clearly, any inefficiencies result-ing from photorespiration in C3 plants are apparentlynot severe. There is no evidence that selection pressureshave caused evolution of a form of Rubisco with loweraffinity for O2.

While most agree that oxygenation is an unavoid-able consequence of evolution, many have argued thatplants have capitalized on this apparent evolutionarydeficiency by turning it into a useful, if not essen-tial, metabolic sequence. The glycolate pathway, forexample, undoubtedly serves a scavenger function. Foreach two turns of the cycle, two molecules of phos-phoglycolate are formed by oxygenation. Of these fourcarbon atoms, one is lost as CO2 and three are returnedto the chloroplast. The glycolate pathway thus recovers75 percent of the carbon that would otherwise be lost asglycolate. The salvage role alone may be sufficient jus-tification for the complex glycolate cycle. There is alsothe possibility that some of the intermediates, serineand glycine, for example, are of use in other biosyn-thetic pathways, although this possibility is still subjectto some debate.

Recently, strong experimental support has beenprovided for the thesis that photorespiration couldalso function as a sort of safety valve in situationsthat require dissipation of excess excitation energy.For example, a significant decline in the photosyn-thetic capacity of leaves irradiated in the absence ofCO2 and O2 has been reported. Injury is prevented,however, if sufficient O2 is present to permit pho-torespiration to occur. Apparently the O2 consumedby photorespiration is sufficient to protect the plantfrom photooxidative damage by permitting continuedoperation of the electron transport system. This couldbe of considerable ecological value under conditionsof high light and limited CO2 supply, for example,when the stomata are closed due to moisture stress(Chapter 14). Indeed, photorespiratory mutants of Ara-bidopsis are more sensitive to photoinhibition than theirwildtype counterparts.

A claim made frequently in the literature is thatcrop productivity might be significantly enhanced byinhibiting or genetically eliminating photorespiration.As a result, substantial effort has been expended in thesearch for chemicals that inhibit the glycolate pathwayor selective breeding for low-photorespiratory strains.Others have surveyed large numbers of species in aneffort to find a Rubisco with a significantly lower affinityfor oxygen. All of these efforts have been unsuccessful,presumably because the basic premise that photorespi-ration is detrimental to the plant and counterproductiveis incorrect. Clearly, success in increasing photosynthe-sis and improving productivity lies in other directions.For example, a mechanism for concentrating CO2 inthe photosynthetic cells could be one way to suppressphotorespiratory loss and improve the overall efficiencyof carbon assimilation. That is exactly what has beenachieved by C4 and CAM plants and will be discussedfurther in Chapter 15.

8.7 Chloroplasts of C3 Plants also Exhibit Competing Carbon Oxidation Processes 145

8.7.3 IN ADDITION TO PCR,CHLOROPLASTS EXHIBITAN OXIDATIVE PENTOSEPHOSPHATE CYCLE

Although the oxidative pentose phosphate cycle(OPPC) is restricted to the cytosol in animals, thispathway is present in both the chloroplast (Figure 8.17)and the cytosol (Chapter 10) in plants. Furthermore,the chloroplastic OPPC shares several intermediateswith the PCR pathway and is closely integratedwith it (Figure 8.17). The first step in the oxidativepentose phosphate cycle is the oxidation of glucose-6-P(G-6-P) to 6-phosphogluconate (6-P-gluconate) bythe enzyme glucose-6-phosphate dehydrogenase(Figure 8.17, reaction 1). The glucose-6-phosphateand fructose-6-phosphate are components of the samestromal hexose phosphate pool that is shared with theRPPC (Figure 8.9). This reaction is highly exergonic(�G < 0), and thus is not reversible. As a consequence,this reaction is apparently the rate-determining stepfor the stromal OPPC. The second reaction in theOPPC involves the oxidation of 6-phosphogluconateto ribulose-5-phosphate (R-5-P) by the enzymegluconate-6-phosphate dehydrogenase with theproduction of one molecule of NADPH and one CO2(Figure 8.17, reaction 2).

The simultaneous operation of both the PCRpathway and the OPPC in the stroma would result inthe reduction of one molecule of CO2 to carbohydrate

CO2

CO2

2x PGA

2x Triose-P

F-6-P

G-6-P

2x 1,3-Bis PGA

2 NADPH

2 NADP+NADP+

NADP+

NADPH

NADPH

6-P-Gluconate

2 ATP

2 ADPATP

ADP

R-5-P

RuBP

1

2

FIGURE 8.17 The simultaneous operation of the PCRcycle and the oxidative pentose phosphate cycle (OPPC)illustrating the potential for the futile cycling of CO2 inthe chloroplast. Reaction 1 is catalyzed by the enzymeglucose-6-P dehydrogenase and reaction 2 by theenzyme phosphogluconate dehydrogenase.

at the expense of three ATP and two NADPH throughthe PCR pathway. Subsequently, the carbohydratewould be reoxidized to CO2 by the OPPC yieldingtwo NADPH. Thus, if both metabolic pathwaysoperate simultaneously in the stroma, three ATPwould be consumed with no net fixation of CO2.This would represent futile cycling of CO2 withthe net consumption of ATP. This would be terriblywasteful!

How do plants overcome the apparent conundrumcreated by the presence of both a reductive andan oxidative pentose phosphate cycle in the samecompartment? The potential for the futile cyclingof CO2 is overcome by metabolic regulation, whichensures that the key enzymes of the PCR cycle are activeonly in the light and inactive in the dark. In contrast, thekey regulatory enzymes of the OPPC are active only inthe dark. Figure 8.13 shows that key regulatory enzymesof the PCR cycle (FBPase, SBPase and Ru-5-P kinase)are converted by light from their inactive to their activeforms by reduced thioredoxin through the reducingequivalents generated by photosynthetic electrontransport. In contrast to stromal FBPase, SBPase, andRu-5-P kinase, which are active when their disulfidebonds are reduced by thioredoxin (—S—S— →— SH HS—), the key regulatory enzyme in theOPPC (glucose-6-P dehydrogenase; Figure 8.17,reaction 1) is active when its internal disulfide bondsare oxidized and inactive when they are reduced bythioredoxin. As a consequence, Rubisco (Figure 8.12),as well as stromal FBPase, SBPase, and Ru-5-Pkinase (Figure 8.13) are in their active states in thelight but phosphogluconate dehydrogenase is in theinactive state, whereas in the dark, phosphogluconatedehydrogenase is in its active state and the keyenzymes of the PRC pathway are inactive. Thus,this exquisite regulation ensures that photosynthesisresults in the net fixation of CO2 and conversion tocarbohydrate and prevents the wasteful consumption ofATP.

The OPPC is thought to be a means to gener-ate NADPH required to drive biosynthetic reactionssuch as lipid and fatty acid biosynthesis in plant mes-ophyll cells. The oxidative pentose phosphate cyclerepresents an important source of pentose phosphate,which serves as a precursor for the ribose and deoxyri-bose required in the synthesis of nucleic acids. Anotherintermediate of the oxidative pentose phosphate path-way with potential significance to plants is the 4-carbonerythrose-4-P, a precursor for the biosynthesis of aro-matic amino acids, lignin, and flavonoids. In addi-tion, the Ru-5-P generated by the OPPC in the darkcan be converted to RuBP in the light to providethe necessary acceptor molecule to get the RPPCstarted.

146 Chapter 8 / Energy Conservation in Photosynthesis: CO2 Assimilation

α α

αβ

β

β

γ

δ

ε

Lumen

Stroma

ADP + Pi

CF1

CF0

ATP

3H+

b

c

3H+

a

BOX 8.1ENZYMES

Living cells must carry out an enormous variety of bio-chemical reactions, yet cells are able to rapidly constructvery large and complicated molecules, or regulate theflow of materials through complex metabolic pathways,with unerring precision and accuracy. All this is madepossible by enzymes. Enzymes are biological catalysts;they facilitate the conversion of substrate molecules toproduct, but are not themselves permanently altered bythe reaction. Cells contain thousands of enzymes, eachcatalyzing a particular reaction.

Enzyme-catalyzed reactions differ from ordinarychemical reactions in four important ways:

1. High specificity. Enzymes are capable of recognizingsubtle and highly specific differences in substrateand product molecules, to the extent of discriminat-ing between mirror images of the same molecules(called stereoisomers or enantiomers) in the sameway you do not fit your right hand into your leftglove.

2. High reaction rates. The rates of enzyme-catalyzedreactions are typically 106 to 1012 greater than ratesof uncatalyzed reactions. Many enzymes are capa-ble of converting thousands of substrate moleculesevery second.

3. Mild reaction conditions. Enzyme reactions typicallyoccur at atmospheric pressure, relatively low tem-perature, and within a narrow range of pH nearneutrality. There are exceptions, such as certainprotein-degrading enzymes that operate in vacuoleswith a pH near 4.0, or enzymes of thermophilicbacteria that thrive in hot sulfur springs, wheretemperatures are close to 100◦C. Most enzymes,however, enable biological reactions to occur underconditions far milder than those required for mostchemical reactions.

4. Opportunity for regulation. The presence of a particu-lar enzyme and its amount is regulated by controlledgene expression and protein turnover. In addition,enzyme activity is subject to regulatory control by avariety of activators and inhibitors.

These opportunities for regulation are instrumentalin keeping complex and often competing metabolicreactions in balance.

The first step in an enzyme-catalyzed reaction is thereversible binding of a substrate molecule (S) with theenzyme (E) to form an enzyme-substrate complex (ES):

E + S � ES → E + P (8.4)

The enzyme-substrate complex then dissociates torelease the product molecule (P). The free enzyme isregenerated and is then available to react with anothermolecule of substrate.

Enzymes are proteins and the site on the proteinwhere the substrate binds and the reaction occurs iscalled the active site. Active sites are usually located in acleft or pocket in the folded protein, and contain reactiveamino acid side chains, such as carboxyl (—COO−),amino (—NH+

3 ), or sulfur (—S−) groups that positionthe substrate and participate in the catalysis. The shapeand polarity of the active site is largely responsible forthe specificity of an enzyme, since the shape and polarityof the substrate molecule must complement or ‘‘fit’’ thegeometry of the active site in order for the substrate togain access and bind to the catalytic groups. Where twoor more substrates participate in a common reaction,binding of the first substrate may induce a change inthe conformation of the protein, which then allows thesecond substrate access to the active site.

Enzymes increase the rate of a reaction because theylower the amount of energy, known as the activationbarrier, required to initiate the reaction. This effect isillustrated by the ball and hill analogy (Figure 8.18A).In order for the ball to roll down the hill, it must firstbe pushed over the lip of the depression in which itsits. This act increases the potential energy of the ball.When the ball is poised at the very top of the lip, it is ina transition state; that is, there is an equal probabilitythat it will fall back into the depression or roll forwardand down the hill.

Chemical reactions go through a similar transi-tion state (Figure 8.18B). As reacting molecules cometogether, they increasingly repel each other and thepotential energy of the system increases. If the reactantsapproach with sufficient kinetic energy, however, theywill achieve a transition state where there is an equalprobability that they will decompose back to reactants orproceed to products. In the case of an enzyme-catalyzedreaction, the enzyme-substrate complex takes a differ-ent reaction pathway—a pathway that has a transitionstate energy level substantially lower than that of theuncatalyzed reaction (Figure 8.18B).

Enzyme-catalyzed reactions exhibit reaction kinet-ics that exhibit a hyberbolic relationship between thereaction velocity, v, and the substrate concentration,[S] (Figure 8.19). Enzymes that exhibit such reactionkinetics are said to follow Michaelis-Menton kinetics.Michaelis-Menton kinetics are characterized by sub-strate saturation, which reflects the fact the enzymebecomes saturated with substrate with increasing sub-strate concentration at constant enzyme concentration.The Michaelis-Menton equation describes this relation-ship mathematically:

v = Vmax [S]/Km + [S] (8.5)

8.7 Chloroplasts of C3 Plants also Exhibit Competing Carbon Oxidation Processes 147

Transition stage

Activation barrier

Ene

rgy

leve

l

Ene

rgy

leve

l

Progress of reactionA. B.

Transition state

Progress of reaction

EnzymecatalyzedReactants

Uncatalyzed Reduction inactivation barrierby enzyme

Products

Free energychange of reaction

FIGURE 8.18 Enzymes. (A) The ‘‘ball and hill’’ analogy of chemical reactions. (B)Enzymes reduce the activation barrier, measured as transition state energy, for areaction.

where v is the initial rate of the reaction, Vmax is themaximum substrate-saturated rate of the reaction, andthe Km is the substrate concentration that provides thehalf-maximal substrate-saturated rate of the reaction.The Km is used as a measure of the affinity that anenzyme has for its substrate: a high Km value implieslow affinity of the enzyme for its substrate, whereas alow Km value implies a high affinity of the enzyme forits substrate.

Vmax

1/2 Vmax

Km

[S]

Init

ial r

ate

of r

eact

ion

(v)

FIGURE 8.19 Plot of initial reaction rate, v, versus sub-strate concentration [S] for an enzyme-catalyzed reac-tion. Vmax is the maximal rate of the reaction undersubstrate-saturated conditions. Km is the value of [S]which provides 1/2 Vmax.

It is important to note that enzymes do not alter thecourse of a reaction. They do not change the equilibriumbetween reactants and products, nor do they alter thefree energy change (�G) for the reaction. (See Chapter 5for a discussion of free energy changes.) Enzymes changeonly the rate of a reaction.

Most enzymes are identified by adding the suffix -aseto the name of the substrate, often with some indicationof the nature of the reaction. For example, α-amylasedigests amylose (starch), malate dehydrogenase oxidizes(that is, removes hydrogen from) malic acid, and phos-phoenolpyruvate carboxylase adds carbon dioxide (acarboxyl group) to a molecule of phosphoenolpyruvate.

Many enzymes do not work alone, but requirethe presence of nonprotein cofactors. Some cofactors,called coenzymes, are transiently associated with theprotein and are themselves changed in the reaction.Many electron carriers, such as NAD+ or FAD, forexample, serve as coenzymes for many dehydrogenaseenzymes (see Chapter 10). They are, in fact, cosubstratesand are reduced to NADH or FADH2 in the reaction.Prosthetic groups are nonprotein cofactors more orless permanently associated with the enzyme protein.The heme group of hemoglobin is an example of atightly bound prosthetic group. Many plant enzymesutilize ions such as iron or calcium as prosthetic groups.

Enzymes and enzyme reactions are sensitive to bothtemperature and pH. Like most chemical reactions,enzyme reactions have a Q10 of about 2, which meansthat the rate of the reaction doubles for each 10◦Crise in temperature. The rate increases with tempera-ture until an optimum is reached, beyond which therate usually declines sharply. The decline is normallycaused by thermal denaturation, or unfolding of the

148 Chapter 8 / Energy Conservation in Photosynthesis: CO2 Assimilation

enzyme protein. With most enzymes, thermal denatu-ration occurs in the range of 40 to 45◦C, although manyenzymes exhibit temperatures closer to 25 or 30◦C.Some enzymes exhibit instability at lower temperaturesas well. One example is pyruvate, pyrophosphate dik-inase (PPDK) (Chapter 15). PPDK is unstable andloses activity at temperatures below about 12 to 15◦C.Enzyme reactions are also sensitive to pH, since pHinfluences the ionization of catalytic groups at the activesite. The conformation of the protein may also bemodified by pH.

Substrate molecules as well as other relatedmetabolites may not only participate in enzyme catalysis(Equation 8.4) but may also stimulate enzyme activity.This phenomenon is called enzyme activation andoccurs as a consequence of the binding of the substrateor metabolite molecule to a site on the enzyme that isdistinct from the active site. This second alternativebinding site on the enzyme is called the allostericsite. The binding of the substrate to the allosteric siteinduces a conformational change in the active site whichenhances the rate at which the substrate (S) is convertedto product (P). Molecules capable of binding to theallosteric site are called effector molecules. Enzymeswhich exhibit such regulation are called allostericenzymes. Rubisco (8.6.2) is an example of an allostericenzyme. CO2 is not only the substrate for the reactioncatalyzed by this enzyme, but also activates Rubiscoactivity by binding to the ε-amino group located in theallosteric site of Rubisco (Figure 8.12).

Conversely, a variety of ions or molecules maycombine with an enzyme in such a way that it reducesthe catalytic activity of the enzyme. These are knownas inhibitors. Inhibition of an enzyme may be eitherirreversible or reversible. Irreversible inhibitors actby chemically modifying the active site so that thesubstrate can no longer bind, or by permanently alteringthe protein in some other way. Reversible inhibitorsoften have chemical structures that closely resemblethe natural substrate. They bind at the active site, buteither do not react or react very slowly. For example,the oxidation of succinate to fumarate by the enzymesuccinic dehydrogenase is competitively inhibited bymalonate, an analog of succinate (Figure 8.20).

Because substrate and inhibitor compete with oneanother for attachment to the active site, this form ofinhibitor is known as competitive inhibition. Anotherform of reversible inhibitor, the noncompetitiveinhibitor, does not compete with the substrate for theactive site, but binds elsewhere on the enzyme and, indoing so, restricts access of the substrate to the activesite. Alternatively, noncompetitive inhibitors may binddirectly to the enzyme-substrate complex, therebyrendering the enzyme catalytically inactive.

Enzymes play a key role in feedback inhibition,one of the most common modes for metabolic

CH2

Succinate

COO

CH2

COO

succinate dehydrogenase

COO

HOOC

C

C

H

Fumarate

Malonate

NO REACTIONsuccinate dehydrogenase

–

COO–

–

–

–

CH2

COO–

FIGURE 8.20 Malonate, a structural analog of succinate,inhibits the enzyme succinate dehydrogenase. Malonatebinds to the enzyme in place of succinate, but does notenter into a reaction.

regulation. Feedback inhibition occurs when the endproduct of a metabolic pathway controls the activityof an enzyme near the beginning of the pathway.When demand for the product is low, excessproduct inhibits the activity of a key enzyme inthe pathway, thereby reducing the synthesis ofproduct. Once cellular activities have depleted thesupply of product, the enzyme is deinhibited andthe rate of product formation increases. The enzymesubject to feedback regulation is usually the first one

A

B

C

D

E

F

G

FIGURE 8.21 Feedback inhibition. Excess product inhibitsthe enzyme that catalyzes a first committed step leadingto product formation.

Chapter Review 149

past a metabolic branch point. This is known as thecommitted step. In the example shown in Figure 8.21,reactions A→B, C→D, and C→F all represent com-mitted steps. In this example, an excess of product Gwould reduce the flow of precursor through the reactionof C → F, thereby diverting more precursor, C, to prod-uct E. Alternatively, an excess of both E and G wouldregulate the conversion of A to B. Feedback regulationis an effective way of coordinating product formationwithin complex pathways. Many of the enzymes of respi-ratory metabolism, for example, are subject to feedback

regulation, thereby balancing the flow of carbon againstthe constantly changing energy demands of the cell.

Enzymes are remarkable biological catalysts thatboth enable and control the enormous variety of bio-chemical reactions that comprise life.

FURTHER READING

Buchanan, B. B., W. Gruissem, R. L. Jones. 2000,Biochemistry and Molecular Biology of Plants. RockvilleMD: American Society of Plant Physiologists.

SUMMARY

Photosynthetic gas exchange between the leaf and theair is dependent upon diffusion and is regulated bythe opening and closing of specialized epidermal porescalled stomata. Stomatal movement is regulated byK+ levels in the guard cells. Opening and closing ofstomata are also sensitive to environmental factors suchas CO2 levels, light, temperature, and the water statusof the plant.

The photosynthetic carbon reduction (PCR)cycle occurs in the chloroplast stroma. It is thesequence of reactions all plants use to reduce carbondioxide to organic carbon. The key enzyme isribulose-1,5-bisphosphate carboxylase-oxygenase(Rubisco), which catalyzes the addition of a car-bon dioxide molecule to an acceptor molecule,ribulose-1,5-bisphosphate (RuBP). The product is twomolecules of 3-phosphoglycerate (3-PGA). Energyfrom the light-dependent reactions is required attwo stages: ATP and NADPH for the reduction of3-PGA and ATP for the regeneration of the acceptormolecule RuBP. The bulk of the cycle involves a seriesof sugar rearrangements that (1) regenerate RuBP and(2) accumulate excess carbon as 3-carbon sugars. Thisexcess carbon can be stored in the chloroplast in theform of starch or exported from the chloroplast fortransport to other parts of the plant.

Photosynthesis, like all other complex metabolicreactions, is subject to regulation. In this case, the pri-mary activator is light. Several key PCR cycle enzymes,including Rubisco, are light activated. This is one wayof integrating photosynthesis with other aspects ofmetabolism, regulating changing levels of intermedi-ates between light and dark periods and competingdemands for carbon with other cellular needs.

Plants that utilize the PCR cycle exclusively forcarbon fixation also exhibit a competing process oflight- and oxygen-dependent carbon dioxide evolution,called photorespiration. The source of carbon dioxide

is the photosynthetic carbon oxidation (PCO) cycle.The PCO cycle also begins with Rubisco, which, inthe presence of oxygen, catalyzes the oxidation, as wellas carboxylation, of RuBP. The product of RuBP oxi-dation is one molecule of 3-PGA plus one 2-carbonmolecule, phosphoglycolate. Phosphoglycolate is sub-sequently metabolized in a series of reactions thatresult in the release of carbon dioxide and recoveryof the remaining carbon by the PCR cycle. The roleof the PCO cycle is not yet clear, although it has beensuggested that it helps protect the chloroplast fromphoto-oxidative damage during periods of moisturestress, when the stomata are closed and the carbondioxide supply is cut off.

Chloroplasts also exhibit an oxidative pentosephosphate cycle (OPPC) that potentially would leadto the futile cycling of CO2. This is prevented by thedifferential light regulation of reductive and oxida-tive pentose phosphate cycles through the action ofthioredoxin.

CHAPTER REVIEW

1. Review the reactions of the photosynthetic carbonreduction cycle and show how:

(a) product is generated;(b) the carbon is recycled to regenerate the accep-

tor molecule.

2. In what chemical form(s) and where is energy putinto the photosynthetic carbon reduction (PCR)cycle? What is the source of this energy?

3. The photosynthetic carbon reduction (PCR) cycleis said to be autocatalytic. What does this meanand of what advantage is it?

4. Describe the photorespiratory pathway. What isthe relationship between photorespiration andphotosynthesis?

150 Chapter 8 / Energy Conservation in Photosynthesis: CO2 Assimilation

5. Debate the position that the oxygenase functionof Rubisco is an evolutionary ‘‘hangover.’’

6. How do plants overcome the potential forfutile cycling of CO2 in the chloroplast?

FURTHER READING

Blankenship, R. E. 2002. Molecular Mechanisms of Photosynthe-sis. London: Blackwell Science.

Buchanan, B. B., Y. Balmer. 2005. Redox regulation: Abroadening horizon. Annual Review of Plant Biology 56:187–220.

Buchanan, B. B., W. Gruissem, R. L. Jones. 2000. Biochem-istry and Molecular Biology of Plants. Rockville MD: Amer-ican Society of Plant Physiologists.

Leegood, R. C., T. D. Sharkey, S. von Caemmerer. 2000.Photosynthesis: Physiology and Metabolism. Advances in Pho-tosynthesis, Vol. 9. Dordrecht: Kluwer.

Spreitzer, R. J., M. E. Salvucci. 2002. Rubisco: Structure, reg-ulatory interactions and possibilities for a better enzyme.Annual Review of Plant Biology 53: 449–475.

H+Apoplast

Cell membrane

Symplast

H+ Sucrose

H+ATP ADP + Pi

Sucrose

9Allocation, Translocation, and Partitioning

of Photoassimilates

The previous two chapters showed how energy wasconserved in the form of carbon compounds, or pho-toassimilates. The primary function of photosynthesisis to provide energy and carbon sufficient to supportmaintenance and growth not only of the photosynthetictissues but of the plant as a whole. During daylighthours, photoassimilate generated by the PCR cycle istemporarily accumulated in the leaf as either sucrosein the mesophyll vacuole or starch in the chloroplaststroma. The conversion of photoassimilates to eithersucrose or starch is called carbon allocation. Althougha portion of the carbon assimilated on a daily basis isretained by the leaf to support its continued growthand metabolism, the majority is exported out of theleaf to nonphotosynthetic organs and tissues. There, itis either metabolized directly or placed in storage forretrieval and metabolism at a later time. The trans-port of photoassimilates over long distances is knownas translocation. Translocation occurs in the vascu-lar tissue called phloem. Phloem translocation is ahighly significant process that functions to ensure anefficient distribution of photosynthetic energy and car-bon between organs throughout the organism. Thisis called carbon partitioning. Phloem translocation isalso important from an agricultural perspective becauseit plays a significant role in determining productivity,

crop yield, and the effectiveness of applied herbicidesand other xenobiotic chemicals.

This chapter is focused on the biosynthesis ofthe photosynthetic end-products, starch and sucrose,and the structure of the phloem and its function inthe translocation and distribution of these importantphotoassimilates. The principal topics to be coveredinclude

• the biosynthesis of starch and sucrose,• the allocation of fixed carbon between the starch

and sucrose biosynthetic pathways,• the basis for identifying phloem as the route for

translocation of photoassimilates and the nature ofsubstances translocated in the phloem,

• the structure of the phloem tissue, especially theseveral unique aspects of sieve tube structure andcomposition,

• the source-sink concept and the significance ofsources and sinks to the translocation process,

• the pressure-flow hypothesis for phloem transloca-tion and the processes by which photoassimilatesgain entry into the phloem in the leaf and are sub-sequently removed from the translocation stream atthe target organ,

151

152 Chapter 9 / Allocation, Translocation, and Partitioning of Photoassimilates

• factors that regulate the distribution of photoassim-ilates between competing sinks, and

• the loading and translocation of xenobiotic agro-chemicals.

9.1 STARCH AND SUCROSEARE BIOSYNTHESIZEDIN TWO DIFFERENTCOMPARTMENTS

Many plants, such as soybean, spinach, and tobacco,store excess photoassimilate as starch in the chloroplast,while others, such as wheat, barley, and oats, accumu-late little starch but temporarily hold large amounts ofsucrose in the vacuole. The appropriation of carbonfixed by the PCR cycle into either starch or sucrosebiosynthesis is called carbon allocation. This will bediscussed further in relation to source sink relation-ships (see below). The starch and sucrose will later bemobilized to support respiration and other metabolicneeds at night or during periods of limited photosyn-thetic output. Sucrose exported from the leaf cell tononphotosynthetic tissues may be metabolized imme-diately, stored temporarily as sucrose in the vacuoles,or converted to starch for longer-term storage in thechloroplasts.

9.1.1 STARCH IS BIOSYNTHESIZEDIN THE STROMA

The dominant storage carbohydrate in higher plantsis the polysaccharide starch, which exists in two forms(Figure 9.1). Amylose is a linear polymer of glucosecreated by linking adjacent glucose residues betweenthe first and fourth carbons. Amylose is consequentlyknown as an α-(1,4)-glucan. Amylopectin is similar toamylose except that occasional α-(1,6) linkages, aboutevery 24 to 30 glucose residues, create a branchedmolecule. Amylopectin is very similar to glycogen, theprincipal storage carbohydrate in animals. Glycogen ismore highly branched, with one α-(1,6) linkage forevery 10 glucose residues compared with one in 30 foramylopectin.

The site of starch synthesis in leaves is thechloroplast. Large deposits of starch are clearly evi-dent in electron micrographs of chloroplasts fromC3 plants. In addition, the two principal enzymes in-volved—ADPglucose pyrophosphorylase and starchsynthase—are found localized in the chloroplaststroma. Starch synthesis in the chloroplast begins withthe hexose phosphate pool generated by the PCRcycle (Figure 9.2). Fructose-6-phosphate (F-6-P), onecomponent of the stromal hexose phosphate pool, isconverted to glucose-1-phosphate, another componentof the stromal hexose phosphate pool, by the two

OO

CH2OHO

H

HOH

OH

HH

HCH2OH

O

H

HOH

OH

HH

H

OO

CH2OHO

H

HOH

OH

HH

HCH2OH

O

H

HOH

OH

HH

H

OO

CH2OHO

H

HOH

OH

HH

H

O

CH2

O

H

HOH

OH

HH

H

... ...

n

�(1 →

O

O

CH2OHO

H

HOH

OH

HH

H

O

CH2OHO

H

HOH

OH

HH

H

...Branch

Mainchain

Glucose Glucose

-Amylose�

6) Branch point

Amylopectin

FIGURE 9.1 The chemical structures of the two forms of starch: amylose and amy-lopectin. Amylose is a long chain of α(1→4)-linked glucose residues. Amylopectinis a multibranched polymer of α(1→4)-linked glucose containing α(1→6) branchpoints.

9.1 Starch and Sucrose Are Biosynthesized in Two Different Compartments 153

Sucrose

CH2OHCH2OH

CH2OH

OHHO

HO

HHO

HO

H

H

H

HHO

OH

H

O

CO2

CO2

Triose-PTranslocatorTriose-P

Hexose-Ppool

Starch

Chloroplast

Pi

Triose-P

Cytosol

Hexose-Ppool

Sucrose Translocation

Pi

Pi

PiPi

Light

MESOPHYLL CELL

A.

B.

FIGURE 9.2 (A) Allocation of fixed carbon between the chloroplast and the cytosol.(B) The structure of sucrose.

chloroplast enzymes, hexose-phosphate isomerase(Equation 9.1) and phosphoglucomutase (Equation 9.2).The glucose-1-P subsequently reacts with ATP to formADP-glucose (Equation 9.3). Reaction 9.3 is cataly-zed by the enzyme ADP-glucose phosphorylase.ADP-glucose is an activated form of glucose and servesas the immediate precursor for starch synthesis. Starchdeposits within the chloroplast stroma are evident asinsoluble starch grains. As a consequence, this form ofstored carbon is osmotically inactive (Chapter 1) whichallows plants to store large amounts of fixed carbonin chloroplasts with minimal influence on the osmoticpressure of the stroma. This prevents the chloroplastmembrane from bursting upon the accumulation andstorage of fixed carbon as starch.

fructose-6-P ↔ glucose-6-P (9.1)

glucose-6-P ↔ glucose-1-P (9.2)

ATP + glucose-1-P ↔ ADP-glucose + H2O + PPi(9.3)

PPi + H2O ↔ 2Pi (9.4)

Finally, the enzyme starch synthase catalyzes formationof a new α-(1,4) link, adding one more glucose to theelongating chain (Equation 9.5).

ADP-glucose + α-(1 → 4)-glucan ↔ ADP+ α-(1 → 4)-glucosyl-glucan (9.5)

Formation of the α-(1,6) branching linkages, giving riseto amylopectin, is catalyzed by the branching enzyme,also known as the Q-enzyme.

9.1.2 SUCROSE IS BIOSYNTHESIZEDIN THE CYTOSOL

Sucrose is a soluble disaccharide containing a glucoseand a fructose residue (Figure 9.2B). It is one of the moreabundant natural products that not only plays a vital rolein plant life but is also a leading commercial commodity.Sucrose may function as a storage product as it does insugarbeets or sugarcane, where it is stored in the vacuolesof specialized storage cells. Alternatively, sucrose maybe translocated to other, nonphotosynthetic tissues inthe plant for direct metabolic use or for conversion tostarch. Sucrose is by far the most common form of sugarfound in the translocation stream.

The site of sucrose synthesis in the cell was thesubject of debate for some time. On the basis ofcell fractionation and enzyme localization studies ithas now been clearly established that sucrose synthe-sis occurs exclusively in the cytosol of photosyntheticcells (Figure 9.2). Earlier reports of sucrose synthesisin isolated chloroplasts appear attributable to contam-ination of the chloroplast preparation with cytosolicenzymes. Moreover, the inner membrane of the chloro-plast envelope is impermeable to sucrose, so that ifsucrose were synthesized inside the chloroplast it wouldbe unable to exit the chloroplast and enter the translo-cation stream.

Two routes of sucrose synthesis are possible. Theprincipal pathway for sucrose synthesis in photosyn-thetic cells is provided by the enzymes sucrose phos-phate synthase (Equation 9.6) and sucrose phosphatephosphatase (Equation 9.7).

154 Chapter 9 / Allocation, Translocation, and Partitioning of Photoassimilates

UDP-glucose + fructose-6-P ↔ sucrose-6-P + UDP(9.6)

sucrose-6-P + H2O ↔ sucrose + Pi (9.7)

Energy provided by the hydrolysis of sucrose-6-phosphate (about 12.5 kJ mol−1) may play a role in theaccumulation of high sucrose concentrations typical ofsugarcane and other sucrose-storing plants.

Another cytoplasmic enzyme capable of synthesiz-ing sucrose is sucrose synthase (SS) (Equation 9.8):

UDP-glucose + fructose ↔ sucrose + UDP (9.8)

With a free energy change of approximately +14 kJmol−1, this reaction is not spontaneous. Most of theevidence indicates that under normal conditions SSoperates in the reverse direction to break down sucrose(see Equation 9.11).

Note that, in contrast with starch biosynthesis,sucrose biosynthesis by either pathway requires acti-vation of glucose with the nucleotide uridine triphos-phate (UTP) rather than ATP:

UTP + glucose-1-P ↔ UDP-glucose + PPi (9.9)

PPi + H2O ↔ 2Pi (9.10)

Although sucrose phosphate synthase in some tissues canuse ADP-glucose, UDP-glucose is clearly predominant.

Carbon for cytoplasmic sucrose biosynthesisis exported from the chloroplast through a specialorthophosphate (Pi)-dependent transporter locatedin the chloroplast envelope membranes (Figure 9.2).This Pi/triose phosphate transporter exchangesPi and triose phosphate—probably as dihydroxyace-tone phosphate (DHAP)—on a one-for-one basis.Once in the cytoplasm, two molecules of triosephosphates (glyceraldehyde-3-phosphate and DHAP)are condensed to form fructose-1,6-bisphosphate.Subsequently, the fructose-1,6-bisphosphate entersthe cytosolic hexose phosphate pool where it isconverted to glucose-1-phosphate as it is in thechloroplast, employing cytoplasmic counterparts of thechloroplastic enzymes. Some of the orthophosphategenerated in sucrose synthesis is used to regenerateUTP while the rest can reenter the chloroplast inexchange for triose-P.

Sucrose translocated from the leaf to storage organssuch as roots, tuber tissue, and developing seeds is mostcommonly stored as starch. The conversion of sucroseto starch is generally thought to involve a reversal of thesucrose synthase reaction:

Sucrose + UDP → fructose + UDP − glucose (9.11)

Because ADP-glucose is preferred for starch biosynthe-sis, UDP-glucose is converted to ADP-glucose as shownin (Equation 9.12) and (Equation 9.13):

UDP-glucose + PPi ↔ UTP + glucose-1-P (9.12)

ATP + glucose-1-P ↔ ADP-glucose + H2O + PPi(9.13)

The resulting ADP-glucose is then converted to starchby starch synthase.

9.2 STARCH AND SUCROSEBIOSYNTHESIS ARECOMPETITIVE PROCESSES

It has traditionally been held that carbohydratemetabolism is to a large extent governed by source-sinkrelationships. The photosynthetically active leaf, forexample, would be a source, providing assimilatedcarbon that is available for transport to the sink,a storage organ or developing flower or fruit, forexample, which utilizes that assimilate. With respect torelationships between sucrose and starch, it was oftenobserved that removal of a sink, thus reducing demandfor photoassimilate, resulted in accumulation of starchin the leaves. This led to the assumption that starchrepresented little more than excess carbon. There isnow good evidence that this assumption is false.

In soybean plants (Glycine max), starch accumulationis not related to the length of the photosynthetic period.Plants maintained on a 7-hour light period put a largerproportion of their daily photoassimilate into starchthan those maintained on a 14-hour light period, eventhough the assimilation period is only half as long. Thusit appears that foliar starch accumulation is more closelyrelated to the energy needs of the daily dark period thanphotosynthetic input. Just how these needs are antici-pated by the plant is unknown. However, many speciesare now known to distribute different proportions ofcarbon between starch and sucrose in ways apparentlyunrelated to sink capacity or inherent capacities of iso-lated chloroplasts to form starch. Carbon distributionthus appears to be a programmed process, implyingsome measure of control beyond a simple source-sinkrelationship. Moreover, it is essential that sucrose syn-thesis be controlled in order to maintain an efficientoperation of photosynthesis itself. If the rate of sucrosesynthesis should exceed the rate of carbon assimilation,demand for triose-P in the cytoplasm could deplete thepool of PCR cycle intermediates, thereby decreasingthe capacity of Calvin cycle enzymes for regeneration ofRuBP and seriously inhibiting photosynthesis.

While the enzyme sucrose phosphate synthase(SPS) determines the maximum capacity for sucrosesynthesis, it appears that cytosolic fructose-1,6-bisphos-phate phosphatase (FBPase) plays the more importantrole in balancing the allocation of carbon betweensucrose and starch synthesis. The highly exergonicreaction (fructose-1,6-bisphosphate → fructose-6-phos-

9.2 Starch and Sucrose Biosynthesized Are Competitive Processes 155

phate + Pi) occupies a strategic site in the sucrosesynthetic pathway—it is the first irreversible reactionin the conversion of triose-P to sucrose. Consequentlythe flow of carbon into sucrose can easily be con-trolled by regulating the activity of FBPase—similarto regulating the flow of water by opening or closinga valve.

Unlike the chloroplastic FBPase, which is lightregulated by thioredoxin, the cytosolic FBPase is notregulated by thioredoxin, but rather is quite sensitive toinhibition by fructose-2,6-bisphosphate (F-2,6-BP)(Figure 9.3). F-2,6-BP, an analog of the natural sub-strate fructose-1,6-bisphosphate, is considered a reg-ulator metabolite because it functions as a regulatorrather than a substrate (Figure 9.4). F-2,6-BP levelsare, in turn, sensitive to a number of interacting factorsincluding the concentration of F-6-P of the cytosolichexose phosphate pool and the cytosolic triose-P/Piratio (Figure 9.4).

Control of sucrose synthesis by F-2,6-BP is essentialto ensure a balance between rates of CO2 assimila-tion and the allocation of fixed carbon. For example,sucrose export from the cell slows in the light, lead-ing to an accumulation of intermediates such as F-6-Pin the cytosolic hexose phosphate pool and triose-P.This causes a shift in allocation in favor of starch.When the consumption of sucrose decreases, sucroseand its precursors (e.g., F-6-P) will accumulate in theleaf cytosolic hexose pool (Figure 9.2). Since F-6-P isalso the precursor for F-2,6-BP, levels of the inhibitorwill increase as well—leading to an inhibition of FBPaseand an accumulation of triose-P. The accumulation ofphosphorylated intermediates probably also leads to adecrease in the concentration of Pi. The combinedaccumulation of triose-P and decrease of Pi will in turndecrease the rate at which triose-P can be exportedfrom the chloroplast through the transporter. Theconsequent accumulation of triose-P and decrease oforthophosphate in the chloroplast in turn stimulate thesynthesis of starch (Figure 9.2). The decrease in stromalPi leads to a reduction in ATP synthesis (ADP + Pi →

O OH

OH

OPHO

H

H HH

POH C2 6

CH2

Fructose-1,6-bisphosphate

Fructose-2,6-bisphosphate

MetaboliteRegulatormetabolite

O O P

OH

OHHO

H

H

POH C2 6

CH2

2

11

2

FIGURE 9.3 Comparison of the structures of F-1,6-BPand F-2,6-BP.

Triose-P

F-1,6-BP

PP-PFK

FBPase(−)

(+)

Hexose-Ppool

Sucrose

F-6-P

F-2,6-BP

Pi

Pi

2

1

ATP

ADP

CYTOSOL

FIGURE 9.4 The synthesis of sucrose is regulated byF-6-P triose-P/Pi ratio. The accumulation of F-6-Pin the cytosolic hexose-P pool favors the synthe-sis of fructose-2,6-bisphosphate (F-2,6-BP) fromF-6-P. F-6-P in the cytosolic hexose-P pool is phos-phorylated by the enzyme, F-6-P-2-kinase (reac-tion 1) and converted to F-2,6-BP. Accumulationof this regulatory metabolite inhibits the activity ofthe enzyme, FBPase, but stimulates the enzyme,pyrophosphate-dependent phosphofructokinase(PPi-PFK), which catalyzes the reverse reaction. Thenet result is a decreased rate of entry of carbon intothe cytosolic hexose-P pool and hence a decreased rateof sucrose synthesis. Low levels of F-6-P favors thebreakdown of F-2,6-BP to F-6-P by the enzymefructose- 2,6-bisphosphatase (reaction 2). This releasescytosolic FBPase from inhibition, and favors thesynthesis of F-6-P which, in turn, favors sucrosesynthesis.

ATP). This causes a buildup of the transthylakoid �pH,which in turn inhibits the rate of photosynthetic elec-tron transport through what is called photosyntheticcontrol. Photosynthetic control is defined as the regu-lation of the rate of photosynthetic electron transport bythe transthylakoid �pH. Hence, this causes a reductionin the rate of photosynthetic O2 evolution and ultimatelythe rate of CO2 assimilation. Plants exhibiting such aninhibition of photosynthesis are said to be feedbacklimited.

Many of the details remain to be worked out, but itis clear that starch synthesis in the chloroplast, triose-Pexport, and sucrose synthesis in the cytosol are in

156 Chapter 9 / Allocation, Translocation, and Partitioning of Photoassimilates

delicate balance. The balance is modulated by verysubtle changes in the level of triose-P and Pi as well asprecise regulation of a number of enzymes and requiresintimate communication between the two cellular com-partments. Thus, the allocation of fixed carbon to eitherstarch or sucrose biosynthesis in leaf mesophyll cellsillustrates that these metabolic pathways actually havea dual role: (1) to provide energy and carbon for growthand the maintenance of homeostasis and (2) to provideinformation with respect to the metabolic status in twodifferent compartments.

9.3 FRUCTAN BIOSYNTHESIS ISAN ALTERNATIVE PATHWAYFOR CARBON ALLOCATION

In addition to carbon allocation to sucrose and starch,about 10 percent of terrestrial plant species exhibit thecapacity to allocate carbon to water-soluble fructosepolymers called fructans, which are biosynthesizedin the vacuole. Plants capable of forming vacuolarfructans include agronomically important crops suchas cereals (wheat, barley, rye), in addition to onion,garlic, leek, Jerusalem artichoke, and chicory. The mostcommon form of fructan in these plants is based on thesequential enzymatic addition of fructose from a donorsucrose molecule to a sucrose acceptor molecule bythe enzyme sucrose:sucrose fructosyl transferase (SST).This results in the formation of the trisaccharide,1-kestose (Equation 9.14), which is composed one ofglucosyl unit linked two fructosyl

Sucrose + sucrose → 1-kestose + glucose (9.14)

units. This trisaccharide is extended by the action of anadditional vacuolar enzyme, fructan:fructan fructosyltransferase (FFT), which results in the formationof a polymer of fructose in the form of glucosyl-1,2-fructosyl-1,2-fructrosy-(fructosyl)N linked in the1,2-β orientation and where N can vary between 1(kestose) and 40 (Equation 9.15).

1-Kestose + fructan → glucosyl-1,2-fructosyl-1,

2-fructrosyl-(fructosyl)N (9.15)

Under conditions where the rate of carbon accu-mulation exceeds the rate of carbon utilization, sucroseaccumulates and the enzymes of vacuolar fructanmetabolism, SST and FFT, are induced. Thus, it is pre-sumed that increase in cytosolic sucrose concentrationstrigger the biosynthesis of fructans. The sucrose accu-mulated in the cytosol is transported to the vacuole andconverted to fructans. Since fructan accumulation canattain levels as high as 40 percent of the dry weight ofcereals, the biosynthesis of vacuolar fructans representsan important mechanism for carbon allocation.

TABLE 9.1 The effects of sucrose accumulationon starch biosynthesis in spinach and the grass,Lolium temulentum.

Leaf Sucrose Starch/SucrosePlant (μmol mg−1 Chl) Ratio

Spinach 1 0.25 0.4

10 0.6Lolium 5 0.1

33 0.150 0.1

Data from Pollack et al., 1995.

As discussed above, plant species such as spinach,in which starch is the major storage carbohydrate,the regulatory metabolite, fructose-2,6-bisphosphate,feedback inhibits the export of triose phosphate fromthe chloroplast stroma in response to increases incytosolic sucrose concentrations. This stimulatesstarch biosynthesis through the activation of theenzyme, ADP-glucose pyrophosphorylase and resultsin an increase in the starch/sucrose ratio (Table 9.1).However, this type of feedback inhibition does notappear to occur in plants such as Lolium temulentum thatconvert sucrose to fructans in the vacuole (Table 9.1).It has been proposed that the apparent insensitivityof chloroplast metabolism to cytosolic sucroseaccumulation in fructan-accumulators may representa selective advantage for grasses which evolved inenvironments where there were rapid changes in thebalance between the supply of and demand for fixedcarbon due either shading, cool temperatures, perennialgrowth habit, or perhaps herbivory. This decreasedsensitivity to feedback limited photosynthesis wouldallow fructan-accumulators to maintain higher rates ofCO2 assimilation due to the maintenance of a greaterflux of carbon through the sucrose biosynthetic pathwaythan starch accumulators.

9.4 PHOTOASSIMILATES ARETRANSLOCATED OVERLONG DISTANCES

Attempts to distinguish between the translocation ofinorganic and organic substances in plants can be tracedback to the seventeenth-century plant anatomist M.Malpighi. In his experiments, Malpighi removed a ringof bark (containing phloem) from the wood (containingxylem) of young stems by separating the two at the vas-cular cambium, a technique known as girdling. Becausethe woody xylem tissue remained intact, water and inor-ganic nutrients continued to move up to the leaves and

9.4 Photoassimilates Are Translocated Over Long Distances 157

the plant was able to survive for some time. Girdledplants, however, developed characteristic swellings ofthe bark in the region immediately above the girdle(Figure 9.5).

Over the years, this experiment has been repeatedand refined to include nonsurgical girdling such as bylocalized steam-killing or chilling. The characteristicswelling is attributed, in part, to an accumulation ofphotoassimilate flowing downward, which is blockedfrom moving further by removal or otherwise inter-fering with the activity of the phloem. As we nowknow, the downward stream also contains nitrogenousmaterial and probably hormones that help to stimulateproliferation and enlargement of cells above the block-age. Eventually, of course, the root system will starvefrom the lack of nutrients and the girdled plant will die.

An analysis of phloem exudate provides more directevidence in support of the conclusion that photoassim-ilates are translocated through the phloem. Unfortu-nately, phloem tissue does not lend itself to analysisas easily as xylem tissue does (described in Chapter 2).This is because the translocating elements in the phloemare, unlike xylem vessels and tracheids, living cells whenfunctional. These cells contain a dense, metabolicallyactive cytoplasm and, because of an inherent sealingaction of its cytoplasm, do not exude their contentsas readily as do xylem vessels. Moreover, phloem con-tains numerous parenchyma cells that, while not directlyinvolved in the transport process, do provide contami-nating cytoplasm. Cutting the stems of some herbaceous

APEX

BASE

Bark(phloem)

Wood(xylem)

A. B.

FIGURE 9.5 The results of girdling on woody stems.(A) The phloem tissue can be removed by separating thephloem (the bark) from the xylem (the wood) at the vas-cular cambium. (B) The girdle interrupts the downwardflow of nutrients and hormones, resulting in a prolifera-tion of tissue immediately above the girdle.

plants will produce an exudate of largely phloem ori-gin, but in some plants, such as some representatives ofthe family Cucurbitaceae, the exudate may quickly gelon contact with oxygen, making collection and subse-quent analysis difficult. The gelling of phloem exudateis due to the properties of a particular phloem protein,which is described more fully later in this chapter. Inspite of these difficulties, however, numerous investi-gators have successfully completed analyses of phloemexudates obtained by making incisions into the phloemtissue, assisted in part by the development of modernanalytical techniques applicable to very small samples.

One intriguing solution to the problem of obtainingthe contents of sieve tubes uncontaminated by other cellswas provided by insect physiologists studying the nutri-tion of aphids. Aphids are one of several groups of smallinsects that feed on plants by inserting a long mouthpart(the stylus) directly into individual sieve tubes. Whenfeeding aphids are anaesthetized with a stream of car-bon dioxide and the stylus carefully severed with a razorblade, phloem sap continues to exude from the cut stylusfor several days. The aphid technique works well for anumber of herbaceous plants and some woody shrubs,but it is restricted to those plants on which the aphidsnaturally feed. The principal advantage of this techniqueis that the severed aphid stylet delivers an uncontam-inated sieve tube sap. Although the volumes deliveredare relatively low, this technique has proven extremelyuseful in studies of phloem transport. The continuedexudation, incidentally, demonstrates that phloem sap isunder pressure, an important observation with respectto the proposed mechanism for phloem transport to bediscussed later.

The third line of evidence involves the use ofradioactive tracers, predominantly 14C and usually fed toa leaf. A typical example is the translocation of photoas-similate in petioles of sugarbeet (Beta vulgaris) leaves.In these experiments, attached leaves were allowedto photosynthesize in a closed chamber containing aradioactive carbon source (14CO2). After 10 minutes,the radiolabeled photoassimilate being transported outof the leaf was immobilized by freezing the petiole in liq-uid nitrogen. Cross sections of the frozen petiole wereprepared and placed in contact with X-ray film. Theresulting image on the X-ray film, or radioautograph,indicated that the radioactive photoassimilate beingtranslocated out of the leaf was localized exclusivelyin the phloem (Figure 9.6). Similar experiments havebeen conducted on a variety of herbaceous and woodyplants and with other radioactive nuclides, such as phos-phorous and sulphur, with the same conclusion— thetranslocation of photoassimilates and other organic compoundsover long distances occurs through the phloem tissue. Thereare exceptions to this rule, such as when stored sugarsare mobilized in the spring of the year and translocatedthrough the xylem to the developing buds (Chapter 2).

158 Chapter 9 / Allocation, Translocation, and Partitioning of Photoassimilates

Epidermis

Parenchyma

FibersVascularbundlePhloem

Xylem

FIGURE 9.6 Location of radioactivity (gray area) in thephloem of sugarbeet petioles after 10 minutes of photo-synthesis in the presence of 14CO2.

9.4.1 WHAT IS THE COMPOSITIONOF THE PHOTOASSIMILATETRANSLOCATED BY THEPHLOEM?

This question may be answered by analyzing the chem-ical composition of phloem exudate. Phloem sap canbe collected from aphid stylets or, alternatively, fromsome plants by simply making an incision into the bark.If done carefully, to avoid cutting into the underlyingxylem, the incision opens the sieve tubes and a relativelypure exudate can be collected in very small microcapil-lary tubes for subsequent analysis. As might be expected,the chemical composition of phloem exudate is highlyvariable. It depends on the species, age, and physiologi-cal condition of the tissue sampled. Even for a particularsample under uniform conditions, there may be widevariations in the concentrations of particular compo-nents between subsequent samples. For example, ananalysis of phloem exudate from stems of actively grow-ing castor bean (Ricinus communis) (Table 9.2) shows thatthe exudate contains sugars, protein, amino acids, theorganic acid malate, and a variety of inorganic anionsand cations. The predominant amino acids are glutamicacid and aspartic acid, which are common forms forthe translocation of assimilated nitrogen (Chapter 11).The inorganic anions include phosphate, sulphate,and chloride—nitrate is conspicuously absent—while the predominant cation is potassium. Althoughnot shown in Table 9.1, some plant hormones (auxin,cytokinin, and gibberellin) were also detected, butat very low concentrations. Of course, many of thecomponents identified in phloem exudate—inorganicions, for example—are cytoplasmic constituents of thetranslocating cells and do not necessarily representtranslocated photoassimilate. Protein found in phloemexudates includes a wide variety of enzymes as wellas one predominant protein (called P-protein) that isunique to the translocating cells. We will return to adiscussion of P-protein later in this chapter.

TABLE 9.2 The chemical composition ofphloem exudate from stems of actively growingcastor bean (Ricinus communis).

Organic mg 1−1

Sucrose 80–106Protein 1.45–2.20Amino acids 5.2Malic acid 2.0–3.2

Inorganic meq 1−1

Anions (inorganic) 20–30Cations (inorganic) 74–138Total dry matter 100–125 mg 1−1

Data from Hall and Baker, 1972.

The principal constituent of phloem exudate inmost species is sugar. In castor bean it is sucrose, whichcomprises approximately 80 percent of the dry matter(Table 9.2). Such a preponderance of sucrose in thetranslocation stream strongly suggests that it is thepredominant form of translocatable photoassimilate.This suggestion has been amply confirmed by labelingexperiments. In the example of translocation in sugar-beet petioles described earlier, more than 90 percent ofthe radioactivity, following 10 minutes of labeling with14CO2, was recovered as sucrose. There are exceptionsto this rule—one is the squash family (Cucurbitaceae),where nitrogenous compounds (principally aminoacids) are quantitatively more important—but overallsugar, particularly sucrose, accounts for the bulkof the translocated carbon. A survey of over 500species representing approximately 100 dicotyledonousfamilies confirms that sucrose is almost universal as thedominant sugar in the phloem stream.

A small number of families translocate, in additionto sucrose, oligosaccharides of the raffinose series (raffi-nose, stachyose, or verbascose) (Figure 9.7). Stachyose,for example, accounts for about 46 percent of the sug-ars in stem internodes of Cucurbita maxima. Yet otherfamilies (Oleaceae, Rosaceae) translocate some of theirphotoassimilates as the sugar alcohols mannitol or sor-bitol.

It is interesting to speculate on why sucrose isthe preferred vehicle for long-distance translocationof photoassimilate. One possibility is that sucrose, adisaccharide, and its related oligosaccharides are nonre-ducing sugars. On the other hand, all monosaccharides,including glucose and fructose, are reducing sugars.Reducing sugars have a free aldehyde or ketone groupthat is capable of reducing mild oxidizing agents. Someoligosaccharides, such as sucrose, are nonreducing sugarsbecause the acetal link between the subunits is stableand nonreactive in alkaline solution. The exclusive use

9.5 Sieve Elements Are the Principal Cellular Constituents of the Phloem 159

Sucrose

CH2OH

Galactose

O CH2

Galactose Galactose Glucose

CH2OHO

O

OH

HO

HOCH2

Fructose

RaffinoseRaffinose

Stachyose

O

OH

OH

HOO

OH

OH

HOO CH2

O

OH

OH

HOO CH2

O

OH

OHHO

VerbascoseFIGURE 9.7 Sugars of the raffinose series. Raffinose, stachyose, and verbascose consistof sucrose with 1, 2, or 3 galactose units, respectively. All sugars in the raffinoseseries, including sucrose, are nonreducing sugars.

of nonreducing sugars in the translocation of photoas-similate may be related to this greater chemical stability.Nonreducing sugars are less likely to react with othersubstances along the way. Indeed, free glucose and fruc-tose, both reducing sugars, are rarely found in phloemexudates. The occasional report of reducing sugars inphloem exudate probably indicates contamination bynonconducting phloem cells, where reducing sugarsare readily formed by hydrolysis of sucrose or otheroligosaccharides.

A second possible factor is that the β-fructoside link-age between glucose and fructose, a feature of sucroseand other members of the raffinose series, has a rela-tively high negative free energy of hydrolysis—about−27 kJ mol−1 compared with about −31 kJ mol−1 forATP. Sucrose is thus a small and highly mobile butrelatively stable packet of energy, which may accountfor its ‘‘selection’’ as the principal form of assimilate tobe translocated in most plants.

9.5 SIEVE ELEMENTS ARE THEPRINCIPAL CELLULARCONSTITUENTS OFTHE PHLOEM

The distinguishing feature of phloem tissue is the con-ducting cell called the sieve element. Also known as asieve tube, the sieve element is an elongated rank ofindividual cells, called sieve-tube members, arrangedend-to-end (Figure 9.8). Unlike xylem tracheary ele-ments, phloem sieve elements lack rigid walls andcontain living protoplasts when mature and functional.The protoplasts of contiguous sieve elements are inter-connected through specialized sieve areas in adjacentwalls. Where the pores of the sieve area are relativelylarge and are found grouped in a specific area, theyare known as sieve plates (Figure 9.8). Sieve plates aretypically found in the end walls of sieve-tube membersand provide a high degree of protoplasmic continuity

between consecutive sieve-tube members. Additionalpores are found in sieve areas located in lateral walls.These are generally smaller and are not, as a rule,grouped in distinct areas. These sieve areas nonethelessprovide cytoplasmic continuity through the lateral wallsof adjacent sieve elements.

Sieveplate

Phloemparenchymacells

Sieve-tubemembers

Phloemparenchymacell

Sieve-tubeplastids

Parenchymaplastid

Companioncell

FIGURE 9.8 Phloem tissue from the stem of tobacco.

160 Chapter 9 / Allocation, Translocation, and Partitioning of Photoassimilates

As noted earlier, mature sieve elements containactive cytoplasm. However, as the sieve elementmatures, it undergoes a series of progressive changesthat result in the breakdown and loss of the nucleus, thevacuolar membrane (or tonoplast), ribosomes, the Golgiapparatus (or dictyosomes), as well as microtubules andfilaments. At maturity, the cells retain the plasmalemma,and endoplasmic reticulum (although it is somewhatmodified), and mitochondria. Even though there is nocentral vacuole as such, the cytoplasmic componentsappear to assume a parietal position in the cell, that is,along the inner wall of the cell.

In addition to sieve elements, phloem tissue alsocontains a variety of parenchyma cells. Some of thesecells are intimately associated with the sieve-tube mem-bers and for this reason are called companion cells.Companion cells (Figure 9.8) contain a full comple-ment of cytoplasm and cellular organelles. A companioncell is derived from the same mother cell as its associ-ated sieve-tube member and shares numerous cytoplas-mic connections with it. The interdependence of thesieve-tube member and companion cell is reflected intheir lifetimes—the companion cell remains alive onlyso long as the sieve-tube member continues to function.When the sieve-tube member dies, its associated com-panion cell also dies. Companion cells are believed toprovide metabolic support for the sieve-tube memberand, perhaps, are involved in the transport of sucrose orother sugars into the sieve tube.

The rest of the phloem parenchyma cells are notalways readily distinguishable from companion cells,even at the ultrastructural level. The single exception isfound in the minor leaf veins of some plants, typically ofherbaceous dicotyledonous plants. Here certain phloemparenchyma cells develop extensive ingrowths of the cellwall. The result is a significant increase in the surfacearea of the plasma membrane. These cells are calledtransfer cells. The precise role of transfer cells is notunderstood but, as the name implies, they are thought tobe involved in collecting and passing on photoassimilatesproduced in nearby mesophyll cells. They may also beinvolved in recycling solutes that enter the apoplast fromthe transpiration stream. These proposed functions arespeculative, based largely on the assumption that thehigh protoplasmic surface area would be expected tofacilitate solute exchange between the transfer cell andthe surrounding apoplast.

9.5.1 PHLOEM EXUDATE CONTAINSA SIGNIFICANT AMOUNTOF PROTEIN

In the early stages of sieve element differentiation,phloem protein (P-protein) appears in the form of dis-crete protein bodies. As the sieve elements mature, theP-protein bodies continue to enlarge. At the time the

nucleus, vacuole, and other cellular organelles disappear,the P-protein bodies disperse in the cytoplasm. In somespecies, such as maple (Acer rubrum), the P-protein takesthe form of a loose network of filaments, ranging from2 to 20 nm in width. In others, such as tobacco (Nicotianasps.), the filaments appear tubular in cross-section. Inyet others, such as some leguminous plants, P-proteintakes the form of crystalline inclusions.

Biochemical investigations of phloem proteinsbegan in the early 1970s, principally in exudates ofCucurbita. Some caution must be exercised when inter-preting these results, however, since phloem exudatescontain proteins in addition to P-protein. Using thetechnique of sodium dodecyl sulphate polyacrylamidegel electrophoresis (SDS-PAGE) (Chapter 7), a varietyof polypeptide subunits with molecular mass valuesranging from 15 to 220 kD have been reported.Apparently phloem protein varies widely betweenspecies, with respect to both its subunit compositionand its chemical properties. One particularly interestingproperty of phloem protein is its capacity to form a gel.Gelation could be prevented by 2-mercaptoethanol,a reducing agent that prevents formation of inter-molecular disulfide (—S—S—) bonds. The effect ofreducing agent is fully reversible—removal of the2-mercaptoethanol allows gelling to proceed. Thiseffect was traced to a single basic protein in thephloem exudate. This protein probably accounts forthe propensity of certain phloem exudates, such as fromCucurbita, to gel rapidly on exposure to air.

P-protein has been the subject of considerable atten-tion over the years because of its prominence in sieveelements and its propensity to plug the pores in the sieveplates. Still, its role and that of other phloem-specificproteins is not yet clear. P-protein has been implicated invarious ways in the transport function of sieve elements.According to some theories, P-protein is consideredan active participant in the transport process. At thesame time, the presence of P-protein in sieve elementsis invoked as an argument against other theories. It isnow generally accepted that, in intact, functioning sieveelements, P-protein is located principally along the innerwall of the sieve element and does not plug the sieveplate.

The formation of plugs in the sieve plates occursonly when the sieve element is injured. This occursbecause the sieve element is normally under positivehydrostatic pressure, as evidenced by the continuedflow of exudate from aphid stylets. When the pressureis released through injury to the sieve element, thecontents, including P-protein, surge toward the site ofinjury. This results in the accumulation of P-protein,possibly assisted by its gelling properties, as ‘‘slime’’plugs on the side of the sieve plate away from the pressurerelease. Thus, it appears that at least one function ofP-protein is protective. By sealing off sieve plates in areas

9.7 Phloem Translocation Occurs by Mass Transfer 161

where the integrity of the phloem has been breached,P-protein helps to maintain the positive hydrostaticpressure in the phloem and reduce unnecessary loss oftranslocated photoassimilate.

Another prominent and somewhat controversialfeature of sieve elements is the presence of callose.Callose, a β1→3-glucan, is related to starch and cel-lulose. Small amounts of callose are deposited on thesurface of the sieve plate or line the pores through whichthe interconnecting strands of cytoplasm pass betweencontiguous cells (Figure 9.8). Controversy over the roleof callose arises from the frequent observation that cal-lose appears to accumulate in the pores to the extent thatit would appear to interfere with translocation. How-ever, it is now known that callose can be synthesizedvery rapidly (within a matter of seconds) and, similar toP-protein, will accumulate in the sieve area in responseto injury. Large amounts of callose also appear to bedeposited on the sieve plates of older, nonfunctionalsieve elements. In both cases, the function of calloseappears to be one of sealing off sieve elements that havebeen injured or are no longer functional, thus preservingthe integrity of the translocating system.

9.6 DIRECTION OFTRANSLOCATION ISDETERMINED BYSOURCE-SINKRELATIONSHIPS

Identification of an organ or tissue as a source or sinkdepends on the direction of its net assimilate transport.An organ or tissue that produces more assimilate than itrequires for its own metabolism and growth is a source. Asource is thus a net exporter or producer of photoassim-ilate; that is, it exports more assimilate than it imports.Mature leaves and other actively photosynthesizing tis-sues are the predominant sources in most plants. A sink,on the other hand, is a net importer or consumer of pho-toassimilate. Roots, stem tissues, and developing fruitsare examples of organs and tissues that normally functionas sinks. The underlying principle of phloem translocation isthat photoassimilates are translocated from a source to a sink.Sink organs may respire the photoassimilate, use it tobuild cytoplasm and cellular structure, or place it intostorage as starch or other carbohydrate.

Any organ, at one time or another in its devel-opment, will function as a sink and may undergo aconversion from sink to source. Leaves are an excel-lent example. In its early stages of development a leafwill function as a sink, drawing photoassimilates fromolder leaves to support its active metabolism and rapidenlargement. However, as a leaf approaches maximumsize and its growth rate slows, its own metabolic demands

diminish and it will gradually switch over to a netexporter. The mature leaf then serves as a source ofphotoassimilate for sinks elsewhere in the plant. Theconversion of a leaf from sink to source is a gradualprocess, paralleling the progressive maturation of leaftissue. In simple leaves, for example, the export of pho-toassimilate from mature regions of the leaf may beginwhile other regions are still developing and functioningas sinks. In compound leaves, such as ash (Fraxinuspennsylvanica) and honeylocust (Gleditsia triacanthos),the early maturing basal leaflets may export photoas-similate to the still-developing distal leaflets as well asout of the leaf.

9.7 PHLOEM TRANSLOCATIONOCCURS BY MASS TRANSFER

What is the mechanism for assimilate translocation overlong distances through the phloem? Any comprehensivetheory must take into account a number of factors. Theseinclude: (1) the structure of sieve elements, includingthe presence of active cytoplasm, P-protein, and resis-tances imposed by sieve plates; (2) observed rapid ratesof translocation (50 to 250 cm hr−1) over long distances;(3) translocation in different directions at the same time;(4) the initial transfer of assimilate from leaf meso-phyll cells into sieve elements of the leaf minor veins(called phloem loading); and (5) final transfer of assim-ilate out of the sieve elements into target cells (calledphloem unloading). Phloem loading and unloading willbe discussed in the following section.

At various times assimilate transport has beenexplained in terms of simple diffusion, cytoplasmicstreaming, ion pumps operating across the sieve plate,and contractile elements in the transcellular protoplas-mic strands. All of these proposals have been largelyrejected on both theoretical and experimental grounds.

The most credible and generally accepted modelfor phloem translocation is one of the earliest. Orig-inally proposed by E. Munch in 1930 but modifiedby a series of investigators since, the pressure-flowhypothesis remains the simplest model and continues toearn widespread support among plant physiologists. Thepressure-flow mechanism is based on the mass transferof solute from source to sink along a hydrostatic (turgor)pressure gradient (Figure 9.9). Translocation of solutein the phloem is closely linked to the flow of water inthe transpiration stream and a continuous recirculationof water in the plant (Chapter 2).

Assimilate translocation begins with the loadingof sugars into sieve elements at the source. Typically,loading would occur in the minor veins of a leaf, closeto a photosynthetic mesophyll or bundle-sheath cell.The increased solute concentration in the sieve elementlowers its water potential (Chapter 1) and, consequently,

162 Chapter 9 / Allocation, Translocation, and Partitioning of Photoassimilates

CO2

Sourcecell

(loading)

Sinkcell

(unloading)

Sucrose

Sucrose

Highpressure

ΔP

Lowpressure

H2O

PHLOEMSIEVE ELEMENT

XYLEMVESSEL

H2O

FIGURE 9.9 A diagram of pressure flow. The loading ofsugar into the sieve element adjacent to a source cellcauses the osmotic uptake of water from nearby xylemelements. The uptake of water increases the hydrostatic(turgor) pressure in the sieve element. The pressure islowered at the sink end when sugar is unloaded into thereceiver cell and the water returns to the xylem. Thispressure differential causes a flow of water from thesource region to the sink. Sugar is carried passively along.

is accompanied by the osmotic uptake of water fromthe nearby xylem. This establishes a higher turgor orhydrostatic pressure (Chapter 1) in the sieve element atthe source end. At the same time, sugar is unloaded atthe sink end—a root or stem storage cell, for example.The hydrostatic pressure at the sink end is lowered aswater leaves the sieve elements and returns to the xylem.So long as assimilates continue to be loaded at the sourceand unloaded at the sink, this pressure differential willbe maintained, water will continue to move in at thesource and out at the sink, and assimilate will be carriedpassively along.

According to the pressure-flow hypothesis, solutetranslocation in the phloem is fundamentally a passiveprocess; that is, translocation requires no direct inputof metabolic energy to make it function. Yet for yearsit has been observed that translocation of assimilateswas sensitive to metabolic inhibitors, temperature, andother conditions, suggesting that metabolic energy wasrequired. More recent experiments, however, have been

designed to discriminate between energy requirementsfor the actual movement of assimilate within the sieveelements and the global energy requirements for translo-cation from source to sink. The results are clear. Theeffects of low temperature and metabolic inhibitors areeither transient or cause disruption of the P-proteinand plug the sieve plates. Energy requirements fortranslocation within the sieve elements are thereforeminimal and compatible with the passive character ofthe pressure-flow hypothesis. The energy requirementindicated in earlier translocation experiments no doubtreflected the needs for loading and unloading the sieveelements.

The principle of pressure flow can be easily demon-strated in the laboratory by connecting two osmometers(Figure 9.10), but a simple physical demonstration doesnot in itself prove the hypothesis. A number of ques-tions must be answered. First, is the sieve tube underpressure? The prolonged exudation of phloem sap fromexcised aphid stylets clearly demonstrates that it is. Thetotal volume of exudate may exceed the volume of anindividual sieve tube by several thousand times. It is dif-ficult to measure the turgor pressure of individual sieve

A. B.

Water

Sucrose and dye solution

Capillary tubing

Plastic or glass tubing

Dialysis tubingcontaining sucroseand dye solution

Water Water

Dialysis tubingcontaining water

FIGURE 9.10 A physical model of the pressure-flowhypothesis for translocation in the phloem. Twoosmometers are constructed from side-arm flasks anddialysis tubing. Osmometer A (the source) initiallycontains a concentrated sucrose solution and a dye.Osmometer B (the sink) contains only water. The twoosmometers are connected by capillary tubing (the ph-loem). Water moves into osmometer A by osmosis, gen-erating a hydrostatic pressure that forces water out ofosmometer B. Water returns via the tubing (the xylem)that connects the side-arm flasks. As a consequence ofthe flow of water between the two ends of the system,the sucrose-dye solution flows through the capillary fromosmometer A to osmometer B. In the model, the systemwill come to equilibrium and flow will cease when thesucrose concentration is equal in the two osmometers.In the plant, flow is maintained because sucrose is con-tinually added to the source (A) and withdrawn at thesink (B).

9.8 Phloem Loading and Unloading Regulate Translocation and Partitioning 163

elements, although a number of attempts have beenmade over the years. Turgor pressure can be calculatedas the difference between sieve tube water potential (�)and osmotic potential (�s) (Chapter 1), or it may bemeasured directly by inserting a small pressure-sensingdevice, or micromanometer, into the phloem tissue. Forexample, the turgor pressure in willow saplings can bemeasured by sealing a closed glass capillary over a sev-ered aphid stylet. Pressure is calculated from the ratioof the compressed and uncompressed air columns inthe capillary. As might be expected, values reported inthe literature range widely, depending on the methodchosen, plant material, the time of day, and physiolog-ical status of the subject plant. Whether calculated ormeasured directly, values of 0.1 MPa to 2.5 MPa aretypical.

A second question to be addressed is whether dif-ferences in sugar concentration and the turgor pressuredrop in the sieve tube are sufficient to account for themeasured rates of transport. Sugar concentration is, ofcourse, highly variable, depending on the rate of photo-synthesis and the general physiological condition of theplant. However, most studies have confirmed that thesugar content of phloem exudate taken near the source ishigher than in exudates taken near sinks. It has been cal-culated that a pressure drop of about 0.06 MPa m−1

would be required for a 10 percent sucrose solution toflow at 100 cm hr−1 through a sieve tube with a radiusof 12 μm. In these calculations, the resistance offeredby sieve plates was taken into account by assuming that(1) the area of the pores in the sieve plate was equal toone-half the area of the sieve tube, (2) there were 60 sieveplates per cm of sieve tube, and (3) the sieve plate poreswere not blocked. Assuming that the turgor pressure ofsieve tubes in the source regions is typically in the range1.0 to 1.5 MPa, and that it is zero in the sink (which maynot be true), a pressure drop of 0.06 MPa m−1 wouldbe sufficient to push a solution through the sieve tubesover a distance of 15 to 25 m. Flow over longer distancescould be accomplished if the source sucrose concen-tration were higher and/or the flow rate were reduced.For example, assimilates can move from the source tothe sink, at a velocity of 48 cm hr−1. A pressure dropof 0.2 MPa would be required to achieve this velocity ifthe sieve plate pores were completely open. From thesucrose concentration in the source and sink, it can becalculated that the actual pressure drop was 0.44 MPa,twice that required! A pressure drop of 0.44 MPa wouldbe sufficient to accomplish a velocity of 48 cm hr−1 evenif the pores were only 70 to 75 percent open.

Another question that is frequently raised in dis-cussions of the pressure-flow hypothesis is that ofbidirectional transport. The translocation of assimi-lates simultaneously in opposite directions would at firstseem incompatible with the pressure-flow hypothesis,but it does occur. Bidirectional transport is first of all

a logical necessity. At any one time, plants will likelyhave more than one sink being served by the samesource—roots for metabolism and storage and devel-oping apical meristems or flowers, for example. It isalso easy to demonstrate experimentally the movementof radiolabeled carbon and phosphorous in oppositedirections through the same internode or petiole at thesame time. This observation might easily be explainedby movement through two separate vascular bundles oreven through different sieve tubes in the same bundle.As long as the sieve elements are connected to differentsinks, the pressure-flow hypothesis does not require thattranslocation occur in the same direction or even at thesame velocity at any one time.

Finally, it has often been argued that sieve elements,because of their structure and composition, offer a sub-stantial resistance to flow and that pressure-flow mightnot provide sufficient force to overcome this resistance.In this regard it is important to note once again thatthe sieve plates in functioning sieve tubes are not occludedby either P-protein or callose. The presence of viscouscytoplasm and sieve plates undoubtedly imposes someresistance, but a variety of experiments have indicatedthat the capacity of the phloem to translocate assimilatesis not normally a limiting factor in the growth of sinks.The phloem is a flexible system for translocation. Itis easily capable of bypassing localized regions of highresistance and the hydrostatic pressure can be adjusted inresponse to demand at either the source or sink. Thereare even developmental controls, apparently to ensurethat the phloem is adequately ‘‘sized’’ to meet antici-pated demand. In wheat, for example, both the numberand size of the vascular bundles serving a floral head cor-relates with the number of flowers. Thus, although thereis little, if any, direct proof for the pressure-flow hypoth-esis, on the balance of evidence it is strongly favored.

9.8 PHLOEM LOADING ANDUNLOADING REGULATETRANSLOCATION ANDPARTITIONING

A discussion of phloem translocation is not completewithout considering how assimilates are translocatedfrom the photosynthetic mesophyll cells into the sieveelements at the source end (phloem loading) or fromthe sieve elements into the target cells at the sink end(phloem unloading).

9.8.1 PHLOEM LOADING CANOCCUR SYMPLASTICALLYOR APOPLASTICALLY

The path traversed by assimilate from the site of pho-tosynthesis to the sieve element is not long. Most

164 Chapter 9 / Allocation, Translocation, and Partitioning of Photoassimilates

mesophyll cells are within a few tenths of a mm,at most three or four cells’ distance from a minorvein ending where loading of assimilate into the sieveelement-companion cell complex (se-cc) actuallyoccurs.1 It is generally agreed that sucrose moves fromthe mesophyll cells to the phloem, probably phloemparenchyma cells, principally by diffusion through theplasmodesmata (i.e., the symplasm). At this point, thepathway becomes less certain and the subject of somedebate. From the phloem parenchyma there are twopossible routes into the se-cc complex (Figure 9.11).Sucrose may continue through the symplasm—that is,through plasmodesmata—directly into the se-cc com-plex. This route is known as the symplastic pathway.Alternatively, the sugar may be transported across themesophyll cell membrane and released into the cell wallsolution (i.e., the apoplasm). From there it would betaken up across the membrane of the se-cc complexwhere it enters the long-distance transport stream. Thisroute is known as the apoplastic pathway.

The apoplastic model for phloem loading gainedfavor in the mid-1970s, based largely on studies oftranslocation in sugarbeet leaves. D. R. Geiger and hiscoworkers used leaves that had been abraded with car-borundum to remove the cuticle, thus improving accessto the leaf apoplast. They found that radioactive sucroseappeared in the apoplast following a period of photosyn-thesis in the presence of 14CO2. They also found thatexogenously supplied sugar was readily absorbed intothe se-cc complex when abraded leaves were bathed in asolution containing 14C-sucrose. These results indicatethat sucrose is normally found in the apoplast and can betaken into the sieve elements from the apoplast. Phloemloading in some plants is also inhibited by chemicalssuch as p-chloromercuribenzene sulfonic acid (PCMBS)when applied to abraded leaves or leaf disks. PCMBSand certain other sulfhydryl-specific reagents presum-ably interfere with carrier proteins (Chapter 3) involvedin the transport of sucrose across the plasma mem-branes. Because these reagents do not penetrate the cellmembrane, any effect they have must be localized onthe apoplastic surface of the membrane.

Sucrose and other sugars are selectively loaded intothe se-cc complex against a concentration gradient,which usually implies active transport. In addition, thereis an increasingly large body of evidence supporting theexistence, in plant cells generally and phloem loading inparticular, of a sucrose-uptake mechanism that is bothATP-dependent and linked to the uptake of protons;that is, a sugar-H+ cotransport (Figure 9.12). This con-

1It is not technically possible to discriminate between therespective roles of the companion cell and the sieve elementin phloem loading and unloading experiments. For thisreason, the sieve element and companion cells are consideredas a single se-cc complex.

Sucrose

Sucrose

SucroseSucrose

Cell wall space (apoplast)Sieve element Source cell

PlasmodesmataS

U

C

R

O

S

E

2

1

(symplast)

FIGURE 9.11 Loading and retrieval of sugars at thesource. Sucrose may be loaded into sieve elements atminor vein endings via one of two pathways. In path-way 1, the symplastic pathway, sugar moves throughplasmodesmata that connect the protoplasts of thesource cell and the sieve element. In pathway 2, theapoplastic pathway, sugar is released into the cell wall(apoplastic) space, from which it is actively transportedacross the plasma membrane of the sieve element bysugar-H+ cotransport. Alternatively, sugar may leak intothe apoplast (dashed lines) and be actively retrieved byeither the source cell or the sieve element.

clusion is supported by the observation that sugar uptakeis accompanied by an increase in pH (i.e., a depletionof protons) or polarity changes in the apoplast. Con-versely, if the pH is experimentally increased—that is,protons are removed from the apoplast by infiltrating theapoplast of abraded leaves with a basic buffer—uptakeinto the phloem cells will be inhibited. Finally, in mostcases it can be shown that only sugars taken up into

H+Apoplast

Cell membrane

Symplast

H+ Sucrose

H+ATP ADP + Pi

Sucrose

FIGURE 9.12 An illustration of sugar-H+ cotransport.The energy for sugar uptake may be provided by a plasmamembrane ATPase proton pump.

9.8 Phloem Loading and Unloading Regulate Translocation and Partitioning 165

the se-cc complex and translocated in the sieve elementswill elicit pH responses. Other sugars, which are notnormally taken up by the phloem, elicit no pH changes.In summary, much of the evidence is consistent with anapoplastic pathway for phloem loading, by which sugarsfirst pass from mesophyll or phloem parenchyma cellsinto the leaf apoplast. The sugar is subsequently takenup into the se-cc complex by sugar-H+ symport, to betranslocated out of the source region.

Although the exact nature of the sucrose-H+ sym-port carrier is not yet worked out, genes (SUT1, SUC2)coding for the carrier have been identified and clonedfrom several species. These include spinach (Spinaceaoleraceae), potato (Solanum tuberosum), Plantago major,and Arabidopsis thaliana. In several species, expressionof the SUC2 sucrose carrier gene appears limited tothe companion cells, while in potato the SUT1 geneproduct was located in the plasma membrane of thesieve elements but could not be detected within thecompanion cells. Such conflicting results might suggestthat apoplastic loading occurs differently in differentspecies or, alternatively, that different carriers are activein companion cells and sieve elements.

In spite of the strong evidence in support of anapoplastic pathway for phloem loading, there are otherdata that support transport through the symplast. Muchof the data from sucrose feeding experiments, forexample, indicate that a significant proportion—up to60 percent—of the radioactive sucrose can be detectedin leaf mesophyll cells. This leaves the experimentsopen to an alternate interpretation: that sucrose-H+cotransport exists as a mechanism for retrieving sucrosethat has leaked from the photosynthetic cells intothe apoplast. Such a retrieval mechanism would notdiscriminate between sugar that leaked from mesophyll,or possibly the se-cc complex, and sugar that wassupplied exogenously by the experimenter. If suchleakage does occur, a mechanism for retrieval wouldserve to prevent unnecessary loss of sugar from thetransport stream. Indeed it has been postulated that aleakage-retrieval cycle occurs normally along the entirelength of the translocation path.

If the uptake data do reflect retrieval by mesophyllcells rather than phloem loading, one is left with theconclusion that phloem loading occurs via the symplasticpathway. Several laboratories have presented evidencethat appears to offer further support for this hypothesis.When mesophyll cells of Ipomea tricolor are injected witha fluorescent dye, the dye moves readily into neighbor-ing mesophyll cells and appears in the minor veins within25 minutes. Since the dye is water-soluble and unable tocross membranes, it is assumed that the dye traveled intothe minor veins via the symplastic connection betweencells. It is also of interest that much of the data sup-porting apoplastic loading have come from experimentswith one species: sugarbeet. In a recent survey, plants

were selected on the basis of whether they had abun-dant symplastic connections between the se-cc complexand adjacent cells of the minor veins, or whether thesecells were symplastically isolated, that is, had no symplas-tic connections. Those plants whose se-cc complexeswere symplastically isolated exhibited characteristics ofapoplastic loading, while those with abundant symplas-tic connections exhibited characteristics of symplasticloading.

The concept of symplastic loading does, however,raise some questions. For example, if sugars diffusefreely from the mesophyll into the se-cc complex, theyshould be equally free to diffuse back into the mesophyllcells. How, then, is it possible for the se-cc complexto accumulate sugars by simple diffusion through theplasmodesmata? Based on studies of phloem loading inCucurbita sps., a polymer trap model to account forsymplastic loading has been proposed. Species, suchas the cucurbits, which have abundant plasmodesmataconnections with the se-cc complex and appear to loadsymplastically, also translocate oligosaccharides in theraffinose series. According to the polymer trap model,sucrose diffuses from the mesophyll or bundle-sheathcells into the companion cells through the connectingplasmodesmata. In the companion cell, the sucrose isconverted to an oligosaccharide, such as the tetrasaccha-ride stachyose, which is too large to diffuse back throughthe plasmodesmata. The polymer (i.e., stachyose) thusremains ‘‘trapped’’ in the se-cc complex, to be carriedaway by mass flow.

The symplastic model assumes that the plasmodes-mata limit the passage of large molecules, but this maynot be the case. Several recent studies of the sucrosetransporter gene have indicated that both the transporterprotein and its mRNA are able to pass through plasmod-esmata between companion cells and sieve elements. Ifmacromolecules can pass through plasmodesmata, it isdifficult to imagine why small oligosaccharides cannot.Perhaps plasmodesmata are more than simple tubesallowing solute flux between cells. This is an excitingissue that will no doubt receive considerable attentionin the future.

Why there is more than one pathway for phlo-em loading is not clear. The symplastic pathwayappears to have an energetic advantage by avoidingtwo carrier-dependent membrane transport steps.The observed energy dependence of loading andtranslocation, however, is more readily explained bythe apoplastic model. In the sympastic model, on theother hand, energy is required for the synthesis ofoligosaccharides in the companion cells. It has also beensuggested that species employing the symplastic pathwayare more ancestral or that the apoplastic pathway is anevolutionary adaptation that arose as plants spread fromtropical climates into more temperate regions. The newmolecular approaches now available will no doubt allow

166 Chapter 9 / Allocation, Translocation, and Partitioning of Photoassimilates

investigators to discriminate between available options.It may be that there is no universal pathway but that thepath of phloem loading is family- or species-specific.Given the theoretical and potential practical significanceof phloem loading in determining yields, we can expectthe investigation and debate to continue.

9.8.2 PHLOEM UNLOADING MAYOCCUR SYMPLASTICALLYOR APOPLASTICALLY

Once assimilate has reached its target sink, it must beunloaded from the se-cc complex into the cells of thesink tissue. In principle, the problem is similar to load-ing; only the direction varies. In detail there are somesignificant differences. As with phloem loading, phloemunloading may occur via symplastic or apoplastic routes(see Figure 9.13). The symplastic route (pathway 1)has been described predominantly in young, developingleaves and root tips. Sucrose flows, via interconnectingplasmodesmata, down a concentration gradient from these-cc complex to sites of metabolism in the sink. Thegradient and, consequently, flow into the sink cell ismaintained by hydrolyzing the sucrose to glucose andfructose.

There are two possible apoplastic routes, shown aspathways 2 and 3 in Figure 9.13. Pathway 2, which hasbeen studied most extensively in the storage parenchymacells of sugarcane, involves the release of sucrose fromthe se-cc complex into the apoplast. Release is insensitiveto metabolic inhibitors or PCMBS and therefore doesnot involve an energy-dependent carrier. Once in theapoplast, sucrose is hydrolyzed by the enzyme acidinvertase, which is tightly bound to the cell wall andcatalyzes the reaction:

Sucrose + H2O → glucose + fructose (9.16)

This reaction is essentially irreversible and the hydroly-sis products, glucose and fructose, are actively taken upby the sink cell. Once in the cell, they are again combinedas sucrose and actively transported into the vacuole forstorage. Hydrolysis of sucrose in the apoplast, perhapscombined with the irreversibility of the acid invertasereaction, serves to maintain the gradient and allowsthe unloading to continue. This pathway seems to beprominent in seeds of maize, sorghum, and pearl millet.

The third pathway for phloem unloading indicatesthat, at least in legumes, sucrose is unloaded into theapoplast by an energy-dependent carrier. The natureof the carrier has not been conclusively identified,but evidence to date suggests it is probably the samesucrose-H+ cotransporter described earlier. As withphloem loading, there does not appear to be a uni-versal path for phloem unloading into the developingembryo.

Sucrose

Sucrose

Glucose

Fructose

Cell wall space (apoplast)

Sieve element Sink cell

Plasmodesmata

(symplast)S

U

C

R

O

S

E

3

2

1

FIGURE 9.13 Three possible routes for sugar unloadinginto sink cells. In all three possible routes, a favorable dif-fusion gradient is maintained by metabolizing the sugaronce it enters the sink cell.

9.9 PHOTOASSIMILATE ISDISTRIBUTED BETWEENDIFFERENT METABOLICPATHWAYS AND PLANTORGANS

Some of the newly fixed carbon or photoassimilatein a source leaf is retained within the leaf, and therest is distributed to various nonphotosynthetic tissuesand organs. This raises several interesting questions.What, for example, determines how much carbon isretained and in what form? What determines how muchis exported and to where? What determines how muchassimilate, for example, is exported to the roots of awheat or corn plant and how much is translocated tofill the developing grain? Questions of this sort havebeen receiving increasing attention of late, because thepatterns of distribution, or more to the point, regulationof the distribution patterns is highly significant withrespect to productivity and yield (Chapter 12). Onemaize farmer may wish to maximize grain yield whileanother may require more of the carbon be put into pro-duction of vegetative (i.e., leafy) material. Each farmerwill assess the harvest index (the ratio of usable plantmaterial to total biomass) for the crop in a different way.

9.9 Photoassimilate Is Distributed Between Different Metabolic Pathways and Plant Organs 167

The traditional route to improving harvest indexhas been through breeding and selection. The uncul-tivated progenitors of modern-day wheat and maize,for example, produced sparse heads with small seeds.Centuries of agricultural selection and, in the last cen-tury, careful breeding, have been required to producethe high-yielding wheat and maize varieties in usetoday. However, the more we learn about the fac-tors regulating carbon distribution and utilization, thegreater the prospects for using modern genetic meth-ods to manipulate the harvest index. The distributionof photoassimilate occurs at two levels: allocation andpartitioning. Each of these will be discussed in turn.

9.9.1 PHOTOASSIMILATES MAY BEALLOCATED TO A VARIETYOF METABOLIC FUNCTIONSIN THE SOURCE OR THE SINK

Allocation refers to the metabolic fate of carbon eithernewly assimilated in the source leaf or delivered to asink. At the source, there are three principal uses forphotoassimilate: leaf metabolism and maintenance ofleaf biomass, short-term storage, or export to otherparts of the plant.

9.9.1.1 Leaf metabolism and biomass Some ofthe carbon will be allocated to the immediate metabolicneeds of the leaf itself. These needs include the main-tenance of cell structure, synthesis of additional leafbiomass, and the maintenance of the photosynthetic sys-tem itself. Most of this carbon is metabolized throughrespiration, which provides both the energy and car-bon skeletons necessary to support ongoing syntheticactivities.

9.9.1.2 Storage Under normal light–dark regimes,plants face a dilemma—photosynthesis is restricted tothe daylight hours, but a supply of photoassimilate forgrowth must be maintained over the entire 24 hours. Apartial solution to this dilemma is to allocate a portionof the newly fixed carbon for storage in the leaves. Mostplants, especially dicots, store the bulk of their carbon asstarch, with a smaller amount stored as sucrose. Some,such as barley (Hordeum vulgare), sugarcane (Saccharumspontaneum), and sugarbeet (Beta vulgaris), accumulatelittle if any starch but store carbon primarily as sucrosein the vacuoles of leaf, stem, or root cells, respectively.Many grasses accumulate fructose polymers called fruc-tans. Carbon stored in the leaves serves primarily asa buffer against fluctuations in metabolite levels and isavailable for reallocation to metabolism when required.

Alternatively, most plants appear to be programmedto maintain a fairly constant rate of translocation andsupply to sink tissues. Leaf reserves are therefore avail-able for reallocation to export at night or during periods

of stress when photosynthesis is very low. In plants thatstore both starch and sucrose, there are generally twopools of sucrose, one in the cytoplasm and one in thevacuole. The vacuolar pool, which is larger and turnsover more slowly than the cytoplasmic pool, is the firstsource of sucrose for export at night. Only when thevacuolar pool is depleted will the starch, stored in thechloroplast, be mobilized for export.

9.9.1.3 Export from the leaf Normally about halfthe newly assimilated carbon is allocated for immediateexport from the leaf via the phloem. In many plants, aportion of this exported carbon may be stored along thetranslocation path. As in the leaf, this stored carbohy-drate helps to buffer the carbon supply at times whenthe rate of translocation through the phloem mightotherwise be reduced.

Regulating the allocation of photoassimilate is acomplex process, involving the interactions of a numberof metabolic pathways. Allocation within a source leafis to a large extent genetically programmed but there isa strong developmental component. Young leaves, forexample, retain a large proportion of their newly fixedcarbon for growth, but as leaves mature the proportionallocated for export increases. In soybean leaves thereare corresponding changes in the activities of enzymessuch as acid invertase (Chapter 10) and sucrose synthase(Equation 9.11). The activities of these two degradativeenzymes are highest in young, rapidly expanding leaves,which no doubt reflects the need to metabolize sucrosein the early stages of leaf development when the leaf isfunctioning primarily as a sink.

As a leaf matures and becomes photosyntheticallyself-sufficient, both its need and capacity to importassimilate decline and the metabolism of the leaf switchesover to the synthesis of sucrose for export. There is acorresponding decline in the activities of acid invertaseand sucrose synthase and a steady increase in the activityof sucrose phosphate synthase (SPS), a key enzyme inthe synthesis of sucrose (Equation 9.6). Because sucroseis the predominant form of translocated carbohydrateand SPS activity is closely correlated with sucrose pro-duction, the increase in SPS activity may be a criticalfactor in determining the transition of the leaf from asink to a source.

The allocation of photoassimilate between storageand export has been extensively described, but there arefew answers to the question of how this allocation isregulated. In most plants, the level of starch fluctuateson a daily basis—increasing during the light period anddeclining at night. The rate of sucrose export exhibitssimilar, but less extreme, diurnal fluctuation. The dis-tribution of carbon between starch and sucrose dependsprimarily on the allocation of triose phosphate betweenstarch synthesis in the chloroplast and sucrose synthesisin the cytoplasm.

168 Chapter 9 / Allocation, Translocation, and Partitioning of Photoassimilates

Since metabolic regulation of starch and sucrosebiosynthesis involves the two key enzymes, fructose-1,6-bisphosphatase (FBPase) and SPS, it is reasonable toexpect that factors that influence allocation do so at leastin part by influencing the activities of these two enzymes.There are some data to bear out these expectations. Incotton leaves, for example, there is a strong correla-tion between SPS activity, sucrose content, and exportof carbon. All three increase more or less in concertduring the photoperiod and drop precipitously at thebeginning of the dark period. During the dark period,sucrose content and SPS activity remain low, but thedrop in export activity is only transient. The pattern ofexport recovery during the dark period corresponds veryclosely with the pattern of starch mobilization. Althoughthere is considerable variation in timing and magni-tude, similar diurnal fluctuations in carbon metabolitesand enzymes have been found in other species. It thusappears that during periods of active photosynthesis,carbon allocation is largely determined by the activityof SPS. At night, the determining factor appears to bethe breakdown of reserve starch.

The single most consistent aspect of source leaf allo-cation, however, is the generally steady rate of export.Except for transient increases at ‘‘dawn’’ or ‘‘dusk,’’diurnal fluctuations in export are small or nonexis-tent. Apparently plants are programmed to maintain asteady rate of assimilate translocation over the entire24-hour period. Whether this program is imposed byphotoperiod or some other factor is not known. Anunderstanding of how allocation is regulated in sourceleaves awaits further investigation.

9.9.2 DISTRIBUTION OFPHOTOASSIMILATESBETWEEN COMPETING SINKSIS DETERMINED BY SINKSTRENGTH

The distribution of assimilate between sinks is referredto as partitioning. In a vegetative plant, the principalsinks are the meristem and developing leaves at the shootapex, roots, and nonphotosynthetic stem tissues. Withthe onset of reproductive growth, the development offlowers, fruits, and seeds creates additional sinks. Ingeneral, sinks are competitive and the photoassimilateis partitioned to all active sinks. If the number of sinksis reduced, a correspondingly higher proportion of thephotoassimilate is directed to each of the remainingsinks. This is the basis for the common practice ofpruning fruit trees to ensure a smaller number of fruitper tree. Partitioning the assimilate among a smallernumber of fruit encourages the development of larger,more marketable fruit.

Partitioning of assimilate between competing sinksdepends primarily on three factors: the nature of vascular

connections between source and sinks, the proximity ofthe sink to the source, and sink strength. Translocation isclearly facilitated by direct vascular connections betweenthe source leaf and the sink. Each leaf is connected tothe main vascular system of the stem by a vascular trace,which diverts from the vascular tissue of the stem into thepetiole. Experiments have shown that photoassimilatewill move preferentially toward sink leaves above and inline (that is, in the same rank) with the source leaf. Thesesink leaves are most directly connected with the sourceleaf. Sink leaves not in the same rank, such as those onthe opposite side of the stem, are less directly linked; theassimilate must make its way through extensive radialconnections between sieve elements.

One of the more significant factors in determiningthe direction of translocation is sink strength. Sinkstrength is a measure of the capacity of a sink to accu-mulate metabolites. It is given as the product of sink sizeand sink activity:

Sink strength = sink size × sink activity (9.17)

Sink size is the total mass of the sink (usually as dryweight). Sink activity is the rate of uptake, or assim-ilate intake per unit dry weight of sink per unit time.Differences in sink strength can be measured experimen-tally, although it is not known exactly what determinessink strength or what causes sink strength to changewith time. The rate of phloem unloading is surely afactor, as well as the rate of assimilate uptake by thesink and allocation to metabolism and storage withinthe sink. Environmental factors (e.g., temperature) andhormones will also have an impact to the extent thatthey influence the growth and differentiation of the sinktissue.

Photoassimilate from most source leaves is read-ily translocated in either vertical direction—upwardtoward the apex or downward toward the roots. All elsebeing equal, however, there is a marked bias in favor oftranslocation toward the closest sink. In the vegetativeplant, photoassimilate from young source leaves near thetop of the plant is preferentially translocated toward thestem apex, while older, nonsenescent leaves near the baseof the plant preferentially supply the roots. Intermediateleaves may translocate photoassimilate equally in bothdirections. The direction of translocation is probablyrelated to the magnitude of the hydrostatic pressuregradient in the sieve elements. Given two equivalentsinks at different distances, the sink closest to the sourcewill be served by the steeper pressure gradient. The biasin favor of the shorter translocation distance is sufficientto overcome even sink size.

Because sink strength is closely related to produc-tivity and yield, most studies have been conducted withcrop species—in particular the filling of grain in cere-als such as wheat (Triticum aestivum) and maize (Zeamays). Developing grain is a particularly active sink and

9.9 Photoassimilate Is Distributed Between Different Metabolic Pathways and Plant Organs 169

has a major impact on translocation patterns. From thetime of anthesis, when the floral parts open to receivepollen, the developing grain becomes the dominant sink.The influence of developing grain on translocation pat-terns is illustrated by the results shown in Table 9.3.In this experiment the supply of photoassimilate wasaltered by reducing the supply of carbon dioxide and thedry-weight increase of various plant parts was monitoredover the grain-filling period. Reducing photoassimilatesupply had virtually no effect on grain weight, whichmeans that a higher proportion of the carbon wastranslocated to the grain. The difference was made upby an equivalent decrease in the proportion of carbondirected to the roots. Roots and the developing grain arecompeting sinks. When the supply of photoassimilate islimited, it is preferentially directed toward the sink withthe greater strength. The dominant role of developinggrain as a sink is also shown by experiments with wheat.When photosynthesis was limited by lowering the lightlevel, the proportion of 14C-photoassimilate from theflag leaf (the leaf directly below the floral head) increasedfrom 49 percent to 71 percent. In this case, however,the difference was made up by an equivalent reductionin the proportion translocated in the lower stem.

The above discussion indicates that sink strengthis a significant factor in determining the pattern oftranslocation, but to suggest that sink strength alone isresponsible for the partitioning of assimilate would beto grossly oversimplify the problem. At the very least,assimilate partitioning is a highly integrated system,depending upon interactions between the source leaf,the actively growing sinks, and the translocation pathitself. We intuitively expect that such an integrated sys-tem will be subject to regulation at one or more points.However, beyond the observation that transport rategenerally responds to sink demand—sudden changes insink activity will cause corresponding changes in trans-port rate to that sink—relatively little is known about

TABLE 9.3 Patterns of photoassimilatedistribution in Sorghum plants subjected to high(400 μl 1−1) and low (250 μl 1−1) concentrationsof carbon dioxide. Values are percentage of totaldry-weight gain during the grain-filling period.Final grain weight was the same under the twoconditions.

Carbon Dioxide Level

High Low

Grain 71.5 87Roots 18 4Other 10.5 9

Based on the data of K. Fischer and G. Wilson, 1975, AustralianJournal of Agricultural Research 26:11–23.

regulation of sink strength and interactions betweensink strength and translocation rate.

Two factors that have been implicated in influenc-ing sink strength are cell turgor and hormones. Whileinvestigating phloem exudate of castor bean (Ricinus com-munis), it was noted that the act of collecting exudate bymaking bark incisions, which causes a sudden reductionin the turgor pressure in the sieve elements, gave riseto a marked increase in sucrose loading at the source.Subsequently, through a series of experiments involv-ing artificial manipulation of turgor, it was concludedthat phloem loading is dependent on turgor pressure inthe sieve elements. Turgor-dependent phloem loadingnow forms the basis for a relatively simple hypothe-sis to explain the regulation of transport rate by sinkdemand. When the se-cc complex is rapidly unloadedat the sink, the reduction in solute concentration causesa corresponding reduction in the hydrostatic pressure,or turgor, at the sink end of the sieve elements (referto Figure 9.9). This reduced hydrostatic pressure willbe transmitted throughout the interconnected systemof sieve elements, quickly stimulating increased phloemloading at the source. The resulting increase in soluteconcentration at the source end of the system wouldserve to counter the drop in hydrostatic pressure, thusmaintaining the pressure gradient and, in accordancewith the pressure-flow mechanism, stimulating the flowof assimilate toward the sink. A reduction in sink demandwould have the opposite result, leading to a lower rateof solute withdrawal and a higher turgor in the sieveelements. Loading at the source and the hydrostaticpressure gradient would be reduced, thereby lower-ing the rate of translocation. According to this model,changes in sieve-element turgor would be an importantmessage in the long-distance communication betweensinks and sources.

It is not known how the se-cc complex or themesophyll cells sense changes in turgor. The mechanismby which pressure changes can be translated into changesin sucrose loading is also unknown. However, someexperiments have demonstrated that sucrose transportacross cell membranes of beet root tissue is turgorregulated, possibly by controlling the activity of anATPase proton pump located in the plasma membrane.

Plant hormones (see Chapters 18–21) have beenimplicated in directing long-distance translocation, par-ticularly with regard to redirection of assimilates tonew sinks. Hormone-directed transport, however, maybe simply an indirect consequence of hormone action.We know that hormones are one of several intrinsicfactors involved in regulating the growth and develop-ment of organs. Through their influence on the sizeand metabolic activity of sink organs, hormones willundoubtedly influence sink strength and, as a result,translocation rates. The role of hormones is compli-cated by the fact that they may, at least in part, be

170 Chapter 9 / Allocation, Translocation, and Partitioning of Photoassimilates

delivered to new sink organs by the phloem. As well,new sinks often themselves become sources of hormonesthat may act locally or be translocated to other regionsof the plant.

While a role for hormone-directed transport overlong distance may be uncertain, there is an accumulat-ing body of evidence that seems to indicate a moredirect involvement of hormones in the transfer ofsolute over short distances. For example, there are anumber of reported correlations between the concen-tration of abscisic acid (ABA; see Chapter 21) and thegrowth rate of developing fruits. ABA also stimulatesthe translocation of sugar into the roots of intact beanplants, the uptake of sucrose by sugarbeet root tis-sue, and the unloading of sucrose into the apoplast ofsoybean seed-coats and its subsequent uptake into theembryo. There have been conflicting reports on whetherABA stimulates the translocation of 14C-photoassimilateinto filling wheat ears. The hormone auxin (IAA; seeChapter 18), on the other hand, inhibits sucrose uptakeby sugarbeet roots but stimulates loading in bean leaves.These and other results suggest that loading and unload-ing may be susceptible to control by hormones.

Although it appears that sink strength is a majorfactor in determining assimilate distribution, the pro-cess of assimilate partitioning remains a complex, highlyintegrated, and poorly understood phenomenon. Inves-tigators have only begun to address the respective rolesof turgor and hormones, while genetic questions andother potential means of regulation have yet to beaddressed in any serious way. The regulation of load-ing, unloading, and source-sink communication shouldcontinue to be active and productive areas of research inthe future.

9.10 XENOBIOTICAGROCHEMICALSARE TRANSLOCATEDIN THE PHLOEM

Phloem mobility is of particular interest to the agro-chemical industry in producing xenobiotic chemicals.The term xenobiotic refers to biologically activemolecules that are foreign to an organism. The rate ofabsorption and translocation of xenobiotic chemicalsoften determines their effectiveness as herbicides,growth regulators, fungicides, or insecticides. Oneexcellent example is the broad-spectrum herbicideN-(phosphonomethyl)glycine, or glyphosate. Glypho-sate acts by preventing the synthesis of aromatic aminoacids, which in turn blocks the synthesis of protein,auxin hormones, and other important metabolites.Because it is highly mobile in the phloem, glyphosateapplied to leaves is rapidly translocated to meristematic

areas or to underground rhizomes for effective controlof perennial weeds.

The principal problem with xenobiotics appears tobe in gaining entry into the phloem at the minor veinendings in the leaf, that is, phloem loading. Althougha few theories have been advanced to explain phloemmobility of xenobiotics, there are relatively few consis-tent chemical and physical characteristics that describethese molecules. Because xenobiotics are not normallyencountered by plants, there are no carriers to mediatetheir uptake by the cell. Entry is probably by pas-sive diffusion. One consistent characteristic of mobilexenobiotics is their relative level of lipid solubility, orlipophilicity, a factor that helps to predict their abilityto diffuse through cell membranes.

Efforts to further understand factors controllingthe entry of xenobiotic chemicals into plants and theirsystemic mobility may ultimately lead to advances in ourunderstanding of phloem translocation generally.

SUMMARY

In many plants, the products of photosynthesis may bestored as starch in the chloroplast or exported fromthe chloroplast to the cytosol where they are con-verted to sucrose. Storage as starch or export to thecytoplasm are competing processes subject to regula-tion by subtle changes in the level of triose phosphateand inorganic phosphate (Pi) as well as the regula-tor metabolite, fructose-2,6-bisphosphate (F-2,6-BP).Other plants such as cereals, store carbon primarilyas fructans in the vacuole and exhibit and insensitiv-ity to feedback inhibition of carbon metabolism underconditions where sucrose accumulates in the cytosol.

The long-distance translocation of photoassimilateand other small organic molecules occurs in the phloemtissue. The distinguishing feature of phloem tissueis the conducting tissue called the sieve element orsieve tube. Filled with modified, but active, protoplasmat maturity, sieve tubes are interconnected throughperforated end walls called sieve plates.

The direction of long-distance translocationin the phloem is determined largely by source-sinkrelationships. An organ or tissue that produces moreassimilate than it requires for its own metabolism isa source, while a sink is a net importer of assimilate.Sinks include meristems and developing leaves atthe apex, nonphotosynthetic stem tissues, roots, andstorage organs. Organs such as leaves are commonlysinks in their early stages, but become sources as theymature.

Sugars are translocated in the phloem by masstransfer along a hydrostatic pressure gradient betweenthe source and sink. Loading of sugars into the sieveelement–companion cell complex (se-cc) in minor

Further Reading 171

veins of the source is followed by the osmotic uptake ofwater. The resulting hydrostatic pressure is transmittedthroughout the system of sieve elements. Unloadingof sugars from the minor veins in the sink maintainsthe pressure differential that causes mass flow. Phloemloading and unloading may occur through the symplast(plasmodesmata) directly into the se-cc complex. Alter-natively, sucrose may be transported across the meso-phyll cell membrane into the apoplastic space. Fromthere it would be taken across the membrane of these-cc complex and enter the long-distance transportstream. There is evidence to support both path-ways, but there are a number of issues yet to beresolved.

The distribution of photoassimilate betweenmetabolic pathways and plant organs occurs at twolevels: allocation and partitioning. Allocation refers tothe immediate metabolic fate of assimilate. It may beallocated to the immediate metabolic needs of the leafitself and maintenance of leaf biomass, it may be storedfor use during nonphotosynthetic periods, or it may beexported from the leaf. Once exported, assimilate willbe partitioned between competing sinks. Partitioningis determined by sink strength, which is a combinationof sink size and metabolic activity.

CHAPTER REVIEW

1. What factors determine whether the product ofthe PCR cycle (triose phosphate) will be con-verted to starch in the chloroplast or sucrose in thecytosol?

2. Distinguish between the roles of F-1,6-BPand F-2,6-BP in the synthesis of sucrose.

3. What is the general structure of a fructan andwhere does it accumulate?

4. What tissues are removed when a tree is girdled?What causes hypertrophic growth above a girdlewound?

5. Describe the structure of mature phloemtissue. What are its unique features? What

kinds of problems do these features raisewith respect to phloem translocation?

6. Describe the source-sink concept. To what extentare source-sink relationships involved in determin-ing the direction and rate of translocation in thephloem?

7. Describe the Munch pressure-flow hypothesis andshow how it operates to drive translocation in thephloem.

8. How are sugars loaded into the phloem sievetubes at the source and removed at the sink?

9. Distinguish between allocation and partitioning.What factors determine allocation of carbonwithin a source leaf? What factors determine par-titioning between more than one potential sink?

FURTHER READING

Buchanan, B. B., W. Gruissem, R. L. Jones. 2000. Biochem-istry and Molecular Biology of Plants. Rockville MD: Amer-ican Society of Plant Physiologists.

Cairns, A. J., C. J. Pollock, J. A. Gallagher, J. Harrison. 2000.Fructans: synthesis and regulation. In: R. C. Leegood,T. D. Sharkey, S. von Caemmerer, Advances in Photosyn-thesis, Vol. 9, pp. 301–320. Dordrecht: Kluwer.

Foyer, C. H., S. Ferrario-Mery, S. C. Huber. 2000. Reg-ulation of carbon fluxes in the cytosol: Coordinationof sucrose synthesis, nitrate reduction and organic acidand amino acid biosynthesis. In: R. C. Leegood, T. D.Sharkey, S. von Caemmerer, Advances in Photosynthesis,Vol. 9, pp. 177–203. Dordrecht: Kluwer.

Lough, T. J., W. J. Lucas. 2006. Integrative plant biology:Role of phloem long-distance molecular trafficking.Annual Review of Plant Biology 57:203–232.

Trethewey, R. N., A. M. Smith. 2000. Starch metabolismin leaves. In: R. C. Leegood, T. D. Sharkey, S.von Caemmerer, Advances in Photosynthesis, Vol. 9,pp. 205–231 Dordrecht: Kluwer.

Turgeon, R. 1996. Phloem loading and plasmodesmata.Trends in Plant Science 1: 418–422.

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FAD

Intermembranespace

Matrix

FMN

Complex I

NADH + H+

NAD+

Complex III

Complex II

3H+

Complex IV

2H+

Succinate Fumarate

Cyt b

Cyt c1

Cyt c Cyt c

H2O F1

1/2 O2 + 2H+

H+

ADP + Pi ATP

F0

ATP Synthase

Cyt a/a3

UQpool

4H+

Innermembrane

10Cellular Respiration: Unlocking the Energy

Stored in Photoassimilates

The previous four chapters have been devoted to theconservation of light energy as compounds of carbon,or photoassimilates, and factors directing the distribu-tion of those carbon compounds into different plantorgans and tissues. Sugars and other photoassimilatesrepresent two important acquisitions by the plant. Theyrepresent, first, a highly mobile form of stored photosyn-thetic energy, and second, a source of carbon skeletons.Through respiration, the plant is able to retrieve theenergy in a more useful form, and in the process thesugars are modified to form the carbon skeletons thatmake up the basic building blocks of cell structure.

This chapter is divided into three principal parts.The first part is devoted to the biochemistry and phys-iology of cellular respiration. After presenting a briefoverview of respiration, the following topics will bediscussed:

• pathways and enzymes involved in the degradationof sucrose and starch to hexose sugars,

• the conversion of hexose to pyruvate via the gly-colytic pathway and the alternate oxidative pentosephosphate pathways,

• the structure and organization of the mitochon-drion, which is the site of oxidative respiratorymetabolism,

• the pathway for the complete oxidation of pyruvateto CO2, known as the citric acid cycle (CAC),the passage of electrons to molecular oxygen viathe mitochondrial electron transport chain, and theconservation of energy as reducing potential andATP,

• several alternative pathways for electron transportthat are unique to plants and their possible physio-logical consequences, and

• the respiration of oils in seeds by first convertingfatty acids to hexose sugars via a process known asgluconeogenesis.

In the second part of this chapter, respiration inintact plants and tissues is discussed, showing how envi-ronmental factors such as light, temperature, and oxygenavailability influence respiration. Finally, the role ofrespiration in the accumulation of biomass and plantproductivity will be briefly examined.

Certain principles introduced earlier (Chapter 5)apply equally well to the present discussion. Theseinclude bioenergetics, oxidation and reduction reac-tions, proton gradients, and the synthesis of adenosinetriphosphate (ATP). You may find it helpful at this timeto review the appropriate sections of Chapter 5.

173

174 Chapter 10 / Cellular Respiration: Unlocking the Energy Stored in Photoassimilates

10.1 CELLULAR RESPIRATIONCONSISTS OF A SERIES OFPATHWAYS BY WHICHPHOTOASSIMILATES AREOXIDIZED

Higher plants are aerobic organisms, which meansthey require the presence of molecular oxygen (O2)for normal metabolism. They obtain both the energyand carbon required for maintenance and growth byoxidizing photoassimilates according to the followingoverall equation:

C6H12O6 + 6O2 + 6H2O → 6CO2 + 12H2O�G◦′ = −2869 kJ mol−1 (10.1)

Note that this equation is written as a reversal of theequation for photosynthesis (Chapter 7, Equation 7.1).The photosynthetic equation is written as the reduc-tion of carbon dioxide to hexose sugar, with water asthe source of electrons. The equation for respiration,on the other hand, is written as the oxidation of hexoseto carbon dioxide, with water as a product. Respirationis accompanied by the release of an amount of freeenergy equivalent to that consumed in the synthesis ofthe same carbon compounds by photosynthesis. Herethe similarity basically ends. Although the two processesoverall share the same reactants and products and theirenergetics are similar, the complex of enzymes involvedand the metabolic routes taken are fundamentally dif-ferent, and they occur in different locations in the cell.Moreover, respiration is a process shared by all livingcells in the plant, while photosynthesis is restricted tothose cells containing chloroplasts.

Equation 10.1 is written as the direct oxidation ofhexose by molecular oxygen, with the consequent releaseof all of the free energy as heat. Cells do not, of course,oxidize sugars in this way. The release of such a largequantity of energy all at once would literally consume thecells. Instead, the overall process of respiration occursin three separate but interdependent stages—calledglycolysis, the citric acid cycle (CAC), and the respi-ratory electron transport chain—comprised of some50 or more individual reactions in total. The transfer ofelectrons to oxygen is but the final step in this long andcomplex process. From the energetic perspective, thefunction of such a complex process is clear: by breakingthe oxidation of hexose down into a series of small, dis-crete steps, the release of free energy is also controlledso that it can be conserved in metabolically useful forms.Equally important to the cell, as we noted earlier, isthe fact that respiration also serves to produce a varietyof carbon skeletons that are then used to build othermolecules required by the cell. We will return to thispoint later in the chapter.

The equation for respiration (Equation 10.1) iscommonly written with hexose (in particular, glucose)as the initial substrate. In practice, a variety of substratesmay serve as the initial substrate. Glucose is itselfderived from storage polymers such as starch (apolymer of glucose), fructans (a polymer of fructose),or the disaccharide, sucrose. Other sugars may alsobe metabolized, as well as lipids, organic acids, andto a lesser extent, protein. The actual substrate beingrespired will depend on the species or organ, stage ofdevelopment, or physiological state.

The type of substrate being respired may onoccasion be indicated by measuring the relativeamounts of O2 consumed and CO2 evolved. From thesemeasurements the respiratory quotient (RQ) can becalculated:

RQ = moles CO2 evolvedmoles O2 consumed

(10.2)

The value of the respiratory quotient is a function of theoxidation state of the substrate being respired. Note thatwhen carbohydrate is being respired (Equation 10.1),the theoretical value of RQ is 6CO2/6O2 = 1.0. Exper-imental values actually tend to vary in the range 0.97to 1.17. Because lipids and proteins are more highlyreduced than carbohydrate, more oxygen is required tocomplete their oxidation and the RQ value may be aslow as 0.7. On the other hand, organic acids, such ascitrate or malate, are more highly oxidized than carbo-hydrate, less oxygen is required for complete oxidation,and RQ values when organic acids are being respiredare typically about 1.3.

While RQ values may provide some useful infor-mation, care must be taken when interpreting them.For example, should more than one type of substratebe respired at any one time, the measured RQ will bean average value. Should fermentation be occurring (seebelow), little or no oxygen will be consumed and anabnormally high RQ may result. Or should either CO2or O2 be trapped in the tissue for any reason, results willbe correspondingly misleading. Still, respiratory quo-tients less than 1 are typical of plants under starvationconditions as lipids and possibly proteins replace carbo-hydrate as the principal respiratory substrate. Anotherexample of the use of RQ is in germinating seeds. Duringgermination, seeds that store large quantities of lipidswill initially exhibit RQ values less than 1. Values willgradually approach 1 as the seedlings consume the lipidreserves and switch over to carbohydrate as the principalrespiratory substrate.

The dependence of plant respiration on photo-synthesis is illustrated in Figure 10.1. The reductionof CO2 in the chloroplast leads to the production offixed carbon in the form of triose phosphates (triose-P),which are represented by the three-carbon phospho-rylated intermediates, dihydroxyacetone phosphate andglyceraldehyde-3-phosphate (see Chapter 8). Triose-P

10.2 Starch Mobilization 175

Triose-P Triose-P Pyruvate

Ethanol

AcetylCoA

CAC

Calvincycle

Light

Starch Sucrose

Export

FADH2

NADH

O2CO2

CO2

CO2CO2CO2O2

–O2

+O2

H2O

ATP

Photosynthesis Glycolysis Respiration

Chloroplast Cytosol Mitochondrion

FIGURE 10.1 The metabolic interaction between the chloroplast, cytosol, and mito-chondrion in a leaf mesophyll cell. Photosynthesis oxidizes water to O2 and reducesCO2 to triose phosphates (triose-P), which represent the fixed carbon substrate formetabolic pathways in the cytosol and the mitochondrion. Triose-P is either storedas starch or exported to the cytosol. In the cytosol, triose-P is either converted tosucrose, which is exported from the mesophyll cell, or is oxidized to pyruvate by theglycolytic pathway. Pyruvate is either oxidized to ethanol in the absence of O2 (fer-mentation) or imported to the mitochondrion and completely oxidized to CO2. Thus,the light energy initially stored as fixed carbon by photosynthesis is eventually con-verted in the mitochondrion to ATP and reducing power (NADH, FADH2), which isused for growth, development, and the maintenance of cellular homeostasis.

is the carbon intermediate that connects anabolic pho-tosynthetic carbon reduction with respiratory oxidativecarbon catabolism. Triose-P generated either directlyby the reduction of CO2 or by the breakdown of starchis exported from the chloroplast to the cytosol, whereit is oxidized by the glycolytic pathway to pyruvate.In the presence of molecular oxygen, pyruvate is com-pletely oxidized through mitochondrial respiration toCO2 with the generation of chemical energy in the formof ATP and NADH used for growth, development, andthe maintenance of homeostasis. The metabolic connec-tion between photosynthesis, glycolysis, and respirationnot only represents the mechanism by which energy, inthe form of fixed carbon, is transferred between threedifferent compartments within a single plant cell, butalso represents an ‘‘information pathway’’ connectingthe chloroplast with the mitochondrion. This can beillustrated by the fact that inhibition of respiratory ATPsynthesis causes PSII reaction centers to become muchmore sensitive to photoinhibition (Chapter 13). Thisability of the chloroplast to respond to events in themitochondrion is the result of metabolic feedback loopssimilar to that discussed in Chapters 8 and 9. Clearly, thechloroplast ‘‘knows’’ what is going on in the mitochon-drion! Research is only beginning to unravel the precise

nature of these metabolic feedback loops between themitochondrion, the chloroplast, and the cytosol.

10.2 STARCH MOBILIZATION

There are two distinct pathways for the breakdown, thatis, the mobilization of starch. The hydrolytic path-way results in the production of glucose whereas thephosphorolytic pathway results in the accumulationof hexose phosphates. These pathways will now be dis-cussed separately in more detail.

10.2.1 THE HYDROLYTICDEGRADATION OF STARCHPRODUCES GLUCOSE

Because most plants store their carbohydrate as starchor sucrose (Chapter 9), the breakdown of these car-bohydrates is an appropriate point at which to beginthe path of respiratory carbon. Starch normally con-sists of a mixture of two polysaccharides: amylose andamylopectin. Amylose, which probably represents nomore than one-third of the starch present in most

176 Chapter 10 / Cellular Respiration: Unlocking the Energy Stored in Photoassimilates

R

R = (1 → 4)-linked α-D-glucose residues

β-Amylase

α-Amylase

FIGURE 10.2 A schematic representation of starch (amylopectin) degradation by α-and β-amylases. Circles indicate (1→4)-linked α-D-glucose residues. Filled circlesindicate the reducing end of the chain.

higher plants, consists of very long, straight chains of(1→4)-linked α-D-glucose units. Amylopectin, on theother hand, is a highly branched molecule in whichrelatively short (1→4)-linked α-D-glucose chains areconnected by (1→6) links (Figure 10.2). Starch is nor-mally deposited in plastids as water-insoluble granulesor grains. The complete breakdown of starch to itscomponent glucose residues requires the participationof several hydrolytic enzymes.

10.2.2 α-AMYLASE PRODUCESMALTOSE AND LIMIT DEXTRINS

α-Amylase randomly cleaves α-(1→4) glucosyl bondsin both amylose and amylopectin (Figure 10.2).α-Amylase, however, does not readily attack terminalα-(1→4) bonds. In the case of amylopectin, α-amylasewill not cleave the α-(1→6) glucosyl bonds, northose α-(1→4) bonds in the immediate vicinity ofthe branch points. Consequently, about 90 percentof the sugar released on hydrolysis of amylose andamylopectin by α-amylase consists of the disaccharidemaltose ((1→4)-α-D-glucosylglucose). The balanceconsists of a small amount of glucose and, in thecase of amylopectin, limit dextrins. Limit dextrinsare comprised of a small number of glucose residues,perhaps 4 to 10, and contain the original branchpoints. α-Amylase is not restricted to plants butcan be found widely in nature, including bacteria

and mammals (including human saliva). Indeed, thisenzyme can be expected in any tissue that rapidlymetabolizes starch. A unique and important propertyof α-amylase is its ability to use starch grains as asubstrate. α-Amylase plays an important role in theearly stages of seed germination, where it is regulatedby the plant hormones gibberellin and abscisic acid(Chapter 19, 21).

10.2.3 β-AMYLASE PRODUCESMALTOSE

β-Amylase degrades amylose by selectively hydrolyzingevery second bond, beginning at the nonreducing end ofthe chain. β-Amylase thus produces exclusively maltose.β-Amylase will degrade the short chains in amylopectinmolecules as well. However, because the enzyme canwork only from the nonreducing end and cannot cleavethe (1→6) branch points, β-amylase will degrade onlythe short, outer chains and will leave the interior of thebranched molecule intact (Figure 10.2).

10.2.4 LIMIT DEXTRINASE IS ADEBRANCHING ENZYME

Limit dextrinase acts on limit dextrins and cleavesthe (1→6) branching bond. This allows both α- andβ-amylase to continue degrading the starch to mal-tose.

10.2 Starch Mobilization 177

10.2.5 α-GLUCOSIDASE HYDROLYZESMALTOSE

The final step is the hydrolysis of maltose to twomolecules of glucose by the enzyme α-glucosidase. All ofthe above enzymes mediate the hydrolytic breakdownof starch to free sugar; that is, the molecule is cleavedessentially by the addition of water across the bond.

10.2.6 STARCH PHOSPHORYLASECATALYZES THEPHOSPHOROLYTICDEGRADATION OF STARCH

When the inorganic phosphate level is high (greaterthan 1 mM), the breakdown of starch is accompanied byan accumulation of phosphorylated sugars. This is due tothe action of the enzyme starch phosphorylase, whichcatalyzes the phosphorolytic degradation of starch:

starch + nPi → n(glucose − 1 − phosphate) (10.3)We will return to this point later, but because the endproduct is glucose-1-phosphate rather than free glucose,the action of phosphorylase offers a slight energeticadvantage. Phosphorylase cannot operate alone—it isunable to degrade starch grains and, like β-amylase,its action is confined to the outer chains of amy-lopectin molecules. Phosphorylase thus can work onlyin conjunction with α-amylase, which initiates degrada-tion of the insoluble grains, and debranching enzymes,

which render the interior glucose chains accessible tothe phosphorylase enzyme. The relative importanceof phosphorylase in vivo is not known, but in lab-oratory experiments phosphorylase accounts for lessthan half of the degradation of potato starch. Thebalance is degraded via α- and β-amylase. Starch isstored and degraded inside plastids (either chloroplastsor amyloplasts—see Chapter 5, Box 5.1), but the initialstages of cellular respiration occur in the cytosol. Theproducts of starch degradation must therefore maketheir way across the plastid envelope in order to gainaccess to the respiratory machinery. This is accom-plished by two transporter systems located in the mem-branes of the plastid envelope (Figure 10.3). The productof phosphorolytic breakdown, glucose-1-phosphate, isa component of the hexose phosphate pool and sub-sequently is converted to triose-P in the chloroplast.Triose-P exits the plastid via the Pi-triose phosphatetransporter described earlier in Chapter 8. Free glucoseis able to exit the plastid via a separate hexose transporter,a protein complex present in the inner envelope mem-brane of the chloroplast, which specifically moves glu-cose from the stroma to the cytosol. For a more detaileddiscussion of membrane transport review Chapter 3.

Sucrose synthesis has been described in Chapter 9.Two enzymes are responsible for its breakdown:sucrose synthase and invertase. Invertase occurs intwo forms, alkaline invertase, with a pH optimumnear 7.5, and acid invertase, with a pH optimum near 5.

Phosphorolytic

Stromal hexose-Ppool

Cytosolic hexose-Ppool

GlycolysisSucrose

Glucose

Glucose

Hydrolytic

Triose-P

Triose-P

Pi

PiCytosol

Stroma

Starch

Fructans

Chloroplastenvelope

FIGURE 10.3 Mobilization of starch in thestroma of the chloroplast and sucrose in thecytosol. Starch is broken down either byhydrolytic or phosphorolytic enzymes. Glu-cose is transported through the chloroplastenvelope membrane by a specific glucosetransporter to the cytosol. In the cytosol, theexported glucose enters the cytosolic hexosephosphate (hexose-P) pool. The breakdownof sucrose also feeds hexose phosphates intothe cytosolic hexose-P pool. The degradationof fructans in the vacuole also feeds hexosephosphates into the cytosolic hexose-P pool.Components of the cytosolic hexose-P poolare converted to cytosolic triose-P. The phos-phorolytic pathway feeds hexose phosphatesinto the stromal hexose-P pool. Hexose-P isconverted to stromal triose-P which is trans-ported out of the chloroplast to the cytosolby the specific Pi-translocator, which importsone Pi molecule for each triose-P exported tothe cytosol. Cytosolic triose-P is then oxidizedthrough glycolysis and respiration. Note thatthe two hexose-P pools are present in differentcompartments that are connected metaboli-cally.

178 Chapter 10 / Cellular Respiration: Unlocking the Energy Stored in Photoassimilates

Sucrose synthase and alkaline invertase appear to belocalized in the cytosol, while acid invertase is foundassociated with cell walls and vacuoles. Clearly therelative contributions of these three enzymes willdepend to some extent on the cellular location of thesucrose being metabolized. Acid invertase, for example,would be important to the mobilization of sucrose insugarcane (Saccharum spontaneum), which stores excesscarbohydrate primarily as sucrose in the vacuoles ofstem cells.

10.3 FRUCTAN MOBILIZATIONIS CONSTITUTIVE

Fructans are soluble polymeric forms of fructose biosyn-thesized in the vacuole as major storage carbohydratesof certain plants species such as grasses (see Chapter 9).Although the enzymes involved on the biosynthesis offructans are inducible through an increase in cytosolicsucrose concentrations, it appears that enzymes involvedin the hydrolysis of fructans are consitutively expressedirrespective of sucrose concentrations. Thus, the netaccumulation of fructans in the vacuole is the resultof the differential, regulated rate of biosynthesis versusthe unregulated rate of hydrolysis. The major enzymeinvolved in fructan hydrolysis in the vacuole is fructanexohydrolase (FEH). This enzyme is an exohydrolasewhich hydrolyzes one terminal fructosyl unit at a timefrom the fructan polymer (Equation 10.4).

Glucosyl-1,2-fructosyl-1,2-fructosyl-(fructosyl)N →glucosyl-1,2-fructosyl-1,2-(fructosyl)N−1 + fructose

(10.4)

The hydrolysis of this polymer is completed by theaction of the vacuolar invertase (Equation 10.5) whichbreaks down the initial sucrose acceptor molecule usedto synthesize the fructan (Chapter 9) into glucose andfructose.

Sucrose → glucose + fructose (10.5)

The free hexoses are then transported from the vacuoleto the cytosol where they are phosphorylated by cytoso-lic hexokinase and enter the cytosolic hexose phosphatepool (Figure 10.4).

10.4 GLYCOLYSIS CONVERTSSUGARS TO PYRUVIC ACID

The first stage of respiratory carbon metabolism is agroup of reactions by which hexose sugars undergoa partial oxidation to the three-carbon acid pyruvicacid or pyruvate. These reactions, collectively knownas glycolysis, are catalyzed by enzymes located in the

cytosol of the cell. Parallel reactions occur indepen-dently in plastids, in particular amyloplasts and somechloroplasts. Thus, unlike animal cells, glycolysis inplants is not restricted to the cytosol. The reaction ofglycolysis, which literally means the lysis or breakdownof sugar, was originally worked out by Meyerhof andothers in Germany during the early part of the twenti-eth century, in order to explain fermentation in yeastsand the breakdown of glycogen in animal muscle tissue.Like much of respiratory metabolism, glycolysis is nowknown to occur universally in all organisms. It is alsobelieved to represent the most primitive form of carboncatabolism since it can lead to fermentation productssuch as alcohol and lactic acid in the absence of molec-ular oxygen. Although the energy yield of glycolysisis low, it can be used to support growth in anaerobicorganisms or in some aerobic organisms or tissues underanaerobic conditions. Under normal aerobic conditions,however, the pyruvate formed by glycolysis will be fur-ther metabolized by the mitochondria to extract yetmore energy.

Glycolysis is conveniently considered in two parts.The first is a set of reactions by which the severalforms of glucose and fructose derived from storagecarbohydrate are converted to the common intermedi-ate triose-phosphate (triose-P) via the hexose-phosphatepool (Figure 10.4). Triose-P is then converted to pyru-vate, the end product of glycolysis (Figure 10.5).

10.4.1 HEXOSES MUST BEPHOSPHORYLATED TOENTER GLYCOLYSIS

In order for carbon from storage carbohydrate toenter glycolysis, the glucose and fructose derived fromhydrolysis of starch, sucrose, or fructans must first beconverted to hexose phosphates. The cytosolic hex-ose phosphate pool consists of glucose-1-phosphate,glucose-6-phosphate, and fructose-6-phosphate. Thesephosphorylated intermediates are subsequently con-verted to fructose-1,6-bisphosphate (FBP) (Figure 10.4).Note that the conversion of glucose and fructose to FBPrequires an initial expenditure of energy in the form ofATP. Two molecules of ATP are consumed for eachmolecule of sucrose that enters glycolysis (reactions 5and 6). This is analogous to priming a pump; glucose isa relatively stable molecule and the initial phosphory-lations, first to glucose-6-P and then to fructose-1,6-P,are a form of activation energy (see Chapter 8, Box 8.1).These two ATP molecules will be recovered duringglycolysis.

It is here the phosphorolytic breakdown of starchin the chloroplast offers a slight energetic advantageover hydrolytic degradation. Because the product isglucose-1-P, for each molecule of hexose entering viathe phosphorolytic route the initial expenditure of ATP

10.4 Glycolysis Converts Sugars to Pyruvic Acid 179

1

2

3

4

5

6

9

10

Phosphorolytic

Starch

Glucose-1-P

Glucose-6-P

Fructose-6-P

Glucose-1-P

Cytosolic hexose-Ppool

Stromal hexose-Ppool

Glucose-6-P

Glucose + Fructose

Fructose-6-P

Sucrose

Fructans

Starch

Glucose

Glucose

Hydrolytic

Triose-P

Fructose-1,6-BP (FBP)

Fructose-1,6-BP (FBP)

ADP

Triose-P Glycolysis

Pi

Pi

Pi

ATP

ADP

ATP

ADPATPAD

PAT

P

Cytosol

Stroma

Vacuole

7

8

EnvelopeMembrane

FIGURE 10.4 The conversion of storage carbohydrate to triose phosphate in thechloroplast and the cytosol. In the chloroplast, starch is either broken down hydrolyt-ically to glucose, which is exported to the cytosol, or broken down phosphorolyti-cally to intermediates of the stromal hexose-P pool (glucose-1-P, glucose-6-P, andfructose-6-P). In the cytosol, sucrose is hydrolyzed to glucose plus fructose. Thehydrolysis of fructans in the vacuole also supply glucose and fructose to the cytosol.Glucose and fructose are converted to intermediates of the cytosolic hexose-P poolby the enzymes hexokinase (5) and fructokinase (6), respectively. The intermedi-ates of the stromal and cytosolic hexose-P pools are interconverted by chloroplasticand cytosolic isoforms of the enzymes phosphoglucomutase (1 and 8, respectively)and hexosephosphate isomerase (2 and 7, respectively). Carbon exits the stromal andcytosolic hexose-P pools through the conversion of fructose-6-P to fructose-1,6-BP(FBP) by the ATP-dependent phosphofructokinase present in the stroma (3) andin the cytosol (9). Fructose-1,6-BP is converted to triose phosphate (triose-P) bychloroplastic and cytosolic isoforms of the enzyme aldolase (4 and 10, respectively).Triose-P is exported from the stroma to cytosol by the Pi-transporter.

180 Chapter 10 / Cellular Respiration: Unlocking the Energy Stored in Photoassimilates

Triose - P

Cytosol

1,3 - Bisphosphoglycerate

Dihydroxyacetone − P Glyceraldehyde − 3 − P

CH3

C = O

ATP

ADP3

2

Pi

NAD+

COOH

CH2

C − O − P

P

COOH

H2C − O −

HCOH

COOH

PH2C − O −

P

HCOH

C

O O −

P− = phosphate group = − PO3H−

3 - Phosphoglycerate

Phosphoenolpyruvate

6

PYRUVATE

4

2 - Phosphoglycerate

5

1

NADH

ADP

ATP

FIGURE 10.5 The conversion of triose-P to pyruvatevia glycolysis. Enzymes are (1) triosephosphate iso-merase, (2) glyceraldehydephosphate dehydrogenase,(3) phosphoglycerate kinase, (4) phosphoglyceratemutase, (5) enolase, (6) pyruvate kinase.

is reduced. Note the overall similarities in the path-ways for the breakdown of starch to triose-P in thestroma with the breakdown of sucrose in the cytosol(Equation 10.6):

storage carbohydrate → hexose-P pool→ FBP → triose-P (10.6)

10.4.2 TRIOSE PHOSPHATES AREOXIDIZED TO PYRUVATE

The reactions for the further conversion of triose-Pto pyruvate are summarized in Figure 10.5. Thetriose phosphates, dihydroxyacetone phosphate andglyceraldehyde-3-phosphate, are readily interconvert-ible (reaction 1), which means that all of the carbonin the original hexose molecule will eventually beconverted to pyruvate. In other words, one molecule ofhexose phosphate will yield two molecules of pyruvate.Thus, in order to account for the hexose moleculeoriginally entering the pathway, everything from thispoint on must be multiplied by 2.

A principal function of glycolysis is energy conser-vation, which occurs in two ways. The first is throughthe production of reducing potential in the form ofNADH. In reaction 2, two molecules of NADH (onefor each triose phosphate) are produced as glyceralde-hyde is oxidized to 1,3-bisphosphoglycerate. Since thispartial oxidation does not require molecular oxygen anddoes not result in the release of any CO2, the glycolyticoxidation of carbohydrate is an anaerobic process. TheNADH produced may be used as reducing potential bythe cell for synthesis of other molecules or, if oxygen ispresent, can be metabolized by plant mitochondria toproduce ATP (see Figure 10.9).

The second way that energy is conserved isthrough the production of ATP via reactions 3 and 6(Figure 10.5). For each molecule of hexose enteringinto glycolysis, four ATP are formed (two for eachtriose phosphate). Note that formation of ATP at thispoint does not involve a proton gradient and cannotbe explained by Mitchell’s chemiosmotic hypothesis. Itis instead linked directly to conversion of substrate inthe pathway. This form of ATP production is called asubstrate-level phosphorylation. Depending on whe-ther the storage carbohydrates were initially degradedby the hydrolytic or the phosphorolytic pathways, thisrepresents a net gain of either two or three ATP.

10.5 THE OXIDATIVE PENTOSEPHOSPHATE PATHWAY ISAN ALTERNATIVE ROUTEFOR GLUCOSE METABOLISM

Most organisms, including both plants and animals,contain an alternative route for glucose metabolismcalled the oxidative pentose phosphate pathway(Figure 10.6). Although this oxidative pathway is res-tricted to the cytosol in animals, this pathway is presentin both the chloroplast (Chapter 8) as well as thecytosol in plants. The oxidative pentose phosphatepathway shares several intermediates with glycolysisand is closely integrated with it. The first step in theoxidative pentose phosphate pathway is the oxidation ofglucose-6-P to 6-phosphogluconate (Figure 10.6). Thisinitial step, which is sensitive to the level of NADP+, isapparently the rate-determining step for the oxidativepentose phosphate pathway. This is the reaction thatdetermines the balance between glycolysis and theoxidative pentose phosphate pathway. The secondstep is another oxidation accompanied by the removalof a CO2 group to form ribulose-5-P. The electronacceptor in both reactions is NADP+, rather thanNAD+. Subsequent reactions in the pathway result inthe formation of glyceraldehyde-3-P and fructose-6-P,both of which are then further metabolized viaglycolysis.

10.6 The Fate of Pyruvate Depends on the Availability of Molecular Oxygen 181

6 − Phosphogluconate

CO2

4

1

3

P PFructose − 6 −

Glucose − 6 −

(To cytosolic Hexose-P pool)

From cytosolicHexose-P pool

PGlyceraldehyde − 3 − PSedoheptulose − 7 −

PRibose − 5 −

PRibulose − 5 −

Cytosol

PXylulose − 5 −5

6

2

NADPH

NADP+

NADP+

NADPH

P

Erythrose − 4 −

FIGURE 10.6 The oxidative pentose phosphate path-way. A principal function of this alternative pathway isto generate reducing potential in the form of NADPHand pentose sugars for nucleic acid biosynthesis. Theorigin of glucose-6-P is the cytosolic hexose-P pool(see Figure 10.4). Glyceraldehyde-3-P and fructose-6-Pmay be returned to the glycolytic pathway for furthermetabolism. Enzymes are: (1) glucose-6-phosphatedehydrogenase, (2) 6-phosphogluconate dehydrogenase,(3) phosphoriboisomerase, (4) phosphopentoepimerase,(5) transketolase, (6) transaldolase.

The role of the oxidative pentose phosphate path-way and its contribution to carbon metabolism overallis difficult to assess because the pathway is not easilystudied in green plants. This is largely because many ofthe intermediates and enzymes of this respiratory cycleare shared by the more dominant reductive pentosephosphate pathway, or PCR cycle, in the chloroplasts(Chapter 8). From studies of animal metabolism, how-ever, it can be concluded that the oxidative pentosephosphate pathway has two significant functions. Thefirst is to generate reducing potential in the form ofNADPH. NADP+ is distinguished from NAD+ byan extra phosphoryl group. NADPH serves primarilyas an electron donor when required to drive normallyreductive biosynthetic reactions, whereas NADH is usedpredominantly to generate ATP through oxidative phos-phorylation (see below). This distinction allows the cellto maintain separate pools of NADPH and NAD+in the same compartment: a high NADPH/NADP+ratio to support reductive biosynthesis and a highNAD+/NADH ratio to support glycolysis. The oxida-tive pentose phosphate pathway is therefore thoughtto be a means to generate NADPH required to drivebiosynthetic reactions in the cytosol. In animals, forexample, the oxidative pentose phosphate pathway isextremely active in fatty tissues where NADPH isrequired for active fatty acid synthesis. The second

function for the oxidative pentose phosphate pathway isthe production of pentose phosphate, which serves asa precursor for the ribose and deoxyribose required inthe synthesis of nucleic acids. Another intermediate ofthe oxidative pentose phosphate pathway with potentialsignificance to plants is the 4-carbon erythrose-4-P, aprecursor for the biosynthesis of aromatic amino acids,lignin, and flavonoids.

10.6 THE FATE OF PYRUVATEDEPENDS ON THEAVAILABILITY OFMOLECULAR OXYGEN

The fate of pyruvate produced by glycolysis dependsprimarily on whether oxygen is present (Figure 10.7).Under normal aerobic conditions, pyruvate is trans-ported into the mitochondrion, where it is furtheroxidized to CO2 and water, transferring its electronsultimately to molecular oxygen. We will address mito-chondrial respiration further in the following section.

Although higher plants are obligate aerobes and areable to tolerate anoxia for only short periods, tissues ororgans are occasionally subjected to anaerobic condi-tions. A typical situation is that of roots when the soil issaturated with water. When there is no oxygen to serveas the terminal electron acceptor, mitochondrial respi-ration will shut down and metabolism will shift over tofermentation. Fermentation converts pyruvate eitherto ethanol through the action of the enzyme alcoholdehydrogenase (ADH) or to lactate via lactate dehy-drogenase (LDH). In most plants, the principal prod-ucts of fermentation are CO2 and ethanol (Figure 10.7,

Acetaldehyde Lactate

Cytosol

CO2 + H2O

+ O2

Mitochondrialrespiration

1

Ethanol

2NADH

NAD+

3

NADH

NAD+

PYRUVATE

Anaerobic - FermentationAerobic

CO2

FIGURE 10.7 The fate of pyruvate depends largely onavailable oxygen. Enzymes are: (1) pyruvate decarboxy-lase, (2) alcohol dehydrogenase (ADH), (3) lactate de-hydrogenase (LDH).

182 Chapter 10 / Cellular Respiration: Unlocking the Energy Stored in Photoassimilates

reactions 1, 2). Some lactate may be formed, primarilyin the early stages of anoxia. However, lactate low-ers the pH of the cytosol, which in turn activatespyruvate decarboxylase and initiates the production ofethanol. Note that either one of the fermentation reac-tions (Figure 10.7, reactions 2 and 3) consumes theNADH produced earlier in glycolysis by the oxida-tion of glyceraldehyde-3-P (Figure 10.5, reaction 2).Although this means there is no net gain of reducingpotential in fermentation, this recycling of NADH isstill important to the cell. The pool of NADH plusNAD+ in the cell is relatively small and if the NADHis not recycled, there will be no supply of NAD+ tosupport the continued oxidation of glyceraldehyde-3-P.If this were the case, glycolysis and the production ofeven the small quantities of ATP necessary to maintainthe cells under anaerobic conditions would then grindto a halt.

10.7 OXIDATIVE RESPIRATIONIS CARRIED OUT BY THEMITOCHONDRION

10.7.1 IN THE PRESENCE OFMOLECULAR OXYGEN,PYRUVATE IS COMPLETELYOXIDIZED TO CO2 AND WATERBY THE CITRIC ACID CYCLE

The second stage of respiration is the complete oxi-dation of pyruvate to CO2 and water through a seriesof reactions known as the citric acid cycle (CAC)(Figure 10.8). The citric acid cycle is also known as thetricarboxylic acid (TCA) cycle or the Krebs cycle, inhonor of Hans Krebs, whose research in the 1930s was

4

3

C COOH

CH2

CH2

COOH

COOH

CoA

Isocitricacid (6C)

α - Ketoglutaricacid (5C)

CO2

CO2

Succinyl − CoA (4C)

Succinicacid (4C)

CoA − SHCoA − SH

Malicacid (4C)

HO

Citric acid (6C)C = O

CH2

COOH

COOH

O

O

Fumaricacid (4C)

7

8

9

56

NAD+

NAD+

FAD

NAD+

ADP+ Pi

CO2

CoA − SH

CoASH

(2C) CH3 − C − S − CoA

Pyruvic acidCH3 − C − COOH(3C)

2

1

NAD+

ATP

NADH

Oxaloacetic acid (4C)

Re

ge

ne

ra

tio

n Ox

ida

tio

n

C o n d e n s a t i o n

NADH

NADH

NADH

FADH2

FIGURE 10.8 The reactions of the citric acid cycle (CAC). The citric acid cycle com-pletes the oxidation of pyruvate to carbon dioxide. Reducing potential is stored asNADH and FADH2.

10.7 Oxidative Respiration is Carried Out by the Mitochondrion 183

responsible for elucidating this central metabolic pro-cess. Krebs was awarded the Nobel Prize in medicine in1954 for his outstanding contribution.

Schemes for the citric acid cycle such as that shownin Figure 10.8 invariably begin with pyruvate. Althoughpyruvate is technically not a part of the cycle, it does pro-vide the major link between glycolysis and subsequentcarbon metabolism. Note that pyruvate is produced inthe cytosol while the enzymes of the citric acid cycleare located in the matrix space of the mitochondrion(Chapter 5). Thus in order for pyruvate to be metabo-lized by the citric acid cycle, it must first be translocatedthrough the inner membrane. This is accomplishedby a pyruvate-OH− antiport carrier—that is, pyruvateis taken up by the mitochondrion in exchange for ahydroxyl ion carried into the intermembrane space.

Once inside the matrix, pyruvate is oxidized anddecarboxylated by a large multienzyme complex pyru-vate dehydrogenase. Pyruvate dehydrogenase catalyzesa series of five linked reactions, the overall effect of whichis to oxidize one molecule of pyruvate to a two-carbonacetate group:

pyruvate + NAD+ + CoA →acetyl-CoA + NADH + H+ + CO2 (10.7)

The resulting two-carbon acetyl group is finally linkedvia a thioester bond to a sulphur-protein coenzymeA(CoA). In the process, NAD+ is reduced to NADH.The CO2 given off represents the first of three carbonatoms in the degradation of pyruvate.

The citric acid cycle proper begins with the enzymecitrate synthase, which condenses the acetyl groupfrom acetyl-CoA with the four-carbon oxaloacetate toform the six-carbon, tricarboxylic citric acid (hence thedesignation citric acid cycle, CAC). The next step isan isomerization of citrate to isocitrate (Figure 10.8,reaction 3), followed by two successive oxidative decar-boxylations (Figure 10.8, reactions 4, 5). The two CO2molecules given off effectively completes the oxidationof pyruvate which adds two more molecules of NADHto the pool of reductant in the mitochondrial matrix.

The balance of the citric acid cycle serves twofunctions. First, additional energy is conserved at threemore locations. One molecule of ATP is formed fromADP and inorganic phosphate when succinate is formedfrom succinyl-CoA (Figure 10.8, reaction 6). Becausethe ATP formation is linked directly to conversion ofsubstrate, this is another example of substrate-levelphosphorylation. Additional energy is conserved withthe oxidation of succinate to fumarate (Figure 10.8,reaction 7) and the oxidation of malate to oxaloacetate(reaction 9). Second, the cycle serves to regenerate amolecule of oxaloacetate and so prepare the cycle toaccept another molecule of acetyl-CoA. Regenerationof oxaloacetate is critical to the catalytic nature of thecycle in that it allows a single oxalacetate molecule to

mediate the oxidation of an endless number of acetylgroups.

In summary, the citric acid cycle consists of eightenzyme-catalyzed steps, beginning with the condensa-tion of a two-carbon acetyl group with the four-carbonoxaloacetate to form a molecule of the six-carbon citrate.The acetyl group is then degraded to two molecules ofCO2. The cycle includes four oxidations, which yieldNADH at three steps and FADH2 at one step. Onemolecule of ATP is formed by substrate-level phospho-rylation. Finally, the oxaloacetate is regenerated, whichallows the cycle to continue.

Before leaving the citric acid cycle for the moment,it is useful to point out that the cycle must turn twice tometabolize the equivalent of one hexose sugar.

10.7.2 ELECTRONS REMOVED FROMSUBSTRATE IN THE CITRICACID CYCLE ARE PASSED TOMOLECULAR OXYGENTHROUGH THEMITOCHONDRIAL ELECTRONTRANSPORT CHAIN

We noted earlier that one of the principal functions ofthe respiration is to retrieve, in useful form, some ofthe energy initially stored in assimilates. Our traditionalmeasure of useful energy in most processes is the numberof ATP molecules gained or consumed. By this mea-sure alone, the yield from both glycolysis and the citricacid cycle is quite low. After two complete turns of thecycle, one molecule of glucose has been completely oxi-dized to six molecules of CO2, but only four moleculesof ATP have been produced (a net of two ATP fromglycolysis plus one for each turn of the cycle). At thispoint, most of the energy associated with the glucosemolecule has been conserved in the form of electronpairs generated by the oxidation of glycolytic and citricacid cycle intermediates. For each molecule of glucose,a total of 12 electron pairs were generated; 10 as NADH(�G◦′ = − 222 kJ mol−1) and two as FADH2 (�G◦′ =− 180 kJ mol−1). Thus, the total energy that has beentrapped as reducing power through the action of gly-colysis and CAC is about 2580 kJ mol−1 [(10 × 222 kJ)+(2 × 180 kJ)]. In addition, the net production of fourATP by substrate phosphorylation in glycolysis and theCAC traps a total of about 125 kJ energy. Therefore, theaerobic oxidation of one glucose molecule traps a totalof about 2705 kJ, which represents an efficiency of about94 percent (2709/2869 × 100%) given that the �G◦′for the oxidation of glucose is about −2869 kJ mol−1.

In this section we will discuss the third stage of cel-lular respiration—the transfer of electrons from NADHand FADH2 to oxygen and the accompanying conver-sion of redox energy to ATP. The transfer of electronsfrom NADH and FADH2 to oxygen involves a sequence

184 Chapter 10 / Cellular Respiration: Unlocking the Energy Stored in Photoassimilates

of electron carriers arranged in an electron transportchain. Membrane fractionation studies have shown thatthe enzymes and electron carriers making up the elec-tron transport chain are organized predominantly intofour large multimolecular complexes (complexes I—IV)and two mobile carriers located in the inner mitochon-drial membrane (Figure 10.9). In this sense, there area great number of similarities between the mitochon-drial inner membrane and the thylakoid membranesof the chloroplast (compare Figure 10.9 with Figure7.6). This is not unexpected, since the principal func-tion of each membrane is energy transformation andmany of the same or similar components are involved.The path of electrons from NADH to oxygen can besummarized as follows. Electrons from NADH enterthe electron transport chain through Complex I, knownas NADH-ubiquinone oxidoreductase. In addition toseveral proteins, this complex also contains a tightlybound molecule of flavin mononucleotide (FMN)and several nonheme iron-sulphur centers. ComplexI conveys the electrons from NADH to ubiquinone.Ubiquinone is a benzoquinone—its structure and func-tion are similar to the plastoquinone found in thethylakoid membranes of chloroplasts (see Chapter 5).Like plastoquinone, ubiquinone is highly lipid solu-ble and diffuses freely in the plane of the membrane.It is not permanently associated with Complex I, butforms a pool of mobile electron acceptors that con-veys electrons between Complex I and Complex III.Ubiquinol (the fully reduced form of ubiquinone) isoxidized by Complex III, or cytochrome c reduc-tase. Complex III contains cytochromes b and c1 andan iron-sulphur center. Complex III in turn reduces amolecule of cytochrome c. Cytochrome c is a peripheral

protein, located on the side of the membrane facing theintermembrane space. Like plastocyanin in chloroplasts,cytochrome c is a mobile carrier and conveys electronsbetween Complex III and the terminal complex in thechain, Complex IV. Also known as cytochrome c oxi-dase, Complex IV contains cytochromes a and a3 andcopper. Electrons are passed first from cytochrome cto cytochrome a, then to cytochrome a3, and finally tomolecular oxygen.

All of the oxidative enzymes of the citric acid cycle,with one exception, are located in the matrix. Theone exception is succinic dehydrogenase (Figure 10.8,reaction 7). This enzyme is an integral protein com-plex (Complex II) that is tightly bound to the innermitochondrial membrane (Figure 10.9). In fact, succinicdehydrogenase is the preferred marker enzyme for innermembranes when doing mitochondrial fractionations.Complex II, known as succinate-ubiquinone oxidore-ductase, contains flavin adenine di-nucleotide (FAD),several nonheme iron proteins, and iron-sulphur cen-ters. Like Complex I, succinic dehydrogenase transferselectrons from succinate to a molecule of ubiquinonefrom the membrane pool. From there the electrons passthrough Complexes III and IV to molecular oxygen.

It is important to note that Figure 10.9 presents astatic, essentially linear representation of electron flowin mitochondria. In vivo, the organization is far moredynamic. In Chapter 7 we showed that the electrontransport complexes of the photosynthetic membraneswere independently distributed, rather than organized inone supermolecular complex. Similar arguments applyhere. The several complexes are not found in equal sto-ichiometry and are free to diffuse independently withinthe plane of the inner membrane. The large complexes

FAD

Intermembranespace

Matrix

FMN

Complex I

NADH + H+

NAD+

Complex III

Complex II

3H+

Complex IV

2H+

Succinate Fumarate

Cyt b

Cyt c1

Cyt c Cyt c

H2O F1

1/2 O2 + 2H+

H+

ADP + Pi ATP

F0

ATP Synthase

Cyt a/a3

UQpool

4H+

Innermembrane

FIGURE 10.9 A schematic representation of the electron transport chain and pro-ton ‘‘pumping’’ sites in the inner membrane of a plant mitochondrion. Solid arrowindicates the path of electrons from NADH or succinate to molecular oxygen.Energy conserved in the proton gradient is used to drive ATP synthesis throughthe F0 —F1-ATPase coupling factor elsewhere in the membrane.

10.8 Energy is Conserved in the Form of ATP in Accordance with Chemiosmosis 185

are functionally linked through the pools of ubiquinoneand cytochrome c, which function as mobile carriers andconvey electrons from one complex to the other largelyon the basis of random collision.

10.8 ENERGY IS CONSERVED INTHE FORM OF ATP INACCORDANCE WITHCHEMIOSMOSIS

As electrons are passed from NADH (or FADH2) tooxygen through the electron transport chain, there is asubstantial drop in free energy. The actual free energychange is quantitatively the same, but of opposite sign, tothe amount consumed when electrons are moved fromwater to NADPH in photosynthesis (Chapter 7). Thisenergy is conserved first in the form of a proton gradientand ultimately as ATP. The energetics for ATP synthe-sis is explained by Mitchell’s chemiosmotic hypothesis,described earlier in Chapter 5. Here the focus is onspecific, biochemical mechanistic details of the protongradient and ATP synthesis as it applies to mitochondria.

Studies of P/O ratios (the atoms of phosphorousesterified as ATP relative to the atoms of oxygenreduced) and various inhibitors have established thatthere are three transitions in the electron transportchain that are associated with ATP synthesis. Putanother way, when internal or matrix NADH isoxidized, the P/O ratio is approximately 3. According toMitchell’s hypothesis, then, these transitions representlocations, generally described as proton pumps, wherecontributions are made to a proton gradient across themitochondrial inner membrane. The three locations,associated with Complexes I, III, and IV, respectively,are identified in Figure 10.9. The resulting protongradient then drives ATP synthesis via a F0-F1-ATPsynthase complex located in the same membrane(see below). Because mitochondrial ATP synthesis isclosely tied to oxygen consumption, it is referred to asoxidative phosphorylation.

In the course of mitochondrial electron transport,protons are extruded from the matrix into the intermem-brane space. Proton extrusion associated with ComplexI (site 1) can be explained by the vectorial arrangementof the complex across the membrane. When a pair ofelectrons is donated to the complex by NADH, a pairof protons are picked up from the matrix. When theelectrons are subsequently passed on to ubiquinone,the protons are released into the intermembrane space.Proton extrusion associated with Complex III (site 2) isprobably due to the operation of a ‘‘Q-cycle,’’ describedin Chapter 7 (see Chapter 7, Figure 7.10). The contri-bution of cytochrome c oxidase (site 3) to the protongradient has been the subject of some discussion formany years. Experiments with isolated enzyme incor-

porated into lipid vesicles indicate that cytochrome coxidase was capable of transferring protons across mem-branes. These results are difficult to explain, becausecytochromes exchange only electrons, not protons, whenreduced and oxidized. However, the H+/electron pairratio for site 3 is about 2, which can readily be explainedby the two protons consumed from the matrix whenoxygen is reduced to water. This is similar in principleto the production of protons in the intrathylakoid space,as water is oxidized early in photosynthetic electrontransport (see Chapter 7, Figure 7.6). The stoichiome-try of proton ‘‘extrusion’’ (the term is applied whetheror not the protons are physically carried across themembrane) has been studied extensively. It appears thatapproximately nine protons are extruded for each pairof electrons conveyed from internal (or matrix) NADHto oxygen.

The link between a proton gradient and ATPsynthesis in the mitochondrion embodies the same prin-ciples previously described for ATP synthesis in thechloroplast: (1) the inner membrane is virtually imper-meable to protons; (2) a proton motive force (pmf)is established across the membrane by a combinationof membrane potential and proton disequilibrium; and(3) ATP synthesis is driven by the return of protonsto the matrix through an integral membrane proteincomplex known variously as ATP synthase, couplingfactor, or F0-F1-ATPase. According to equation 10.8,pmf consists of two principal components: a chemicalcomponent, that is, �pH, which reflects the differencein H+ concentration, and an electrical component (��),which reflects a difference in charge between the matrixand the intermembrane space.

pmf = −59VpH + � (10.8)

Due to the capacity of the cytosol to buffer changesin pH in the intermembrane space, �pH contributesminimally to the overall pmf in mitochondria. Thus, incontrast to the chloroplast, the difference in electricalcharge (��) across the inner membrane is the majorfactor contributing to the proton motive force generatedby mitochondria.

Mitochondrial ATP synthase is structurally andfunctionally similar to the chloroplast enzyme. It con-sists of a hydrophobic, channel-forming portion (F0) thatspans the membrane plus a multimeric, matrix-facingperipheral protein (F1) that couples proton transloca-tion to ATP synthesis. As in the case of chloroplasts,the H+/ATP ratio is approximately 3. Because nineprotons are extruded for each pair of electrons mov-ing through the entire chain, this means that a totalof three ATP molecules could be formed from eachNADH produced in the matrix. For electrons enteringthe chain from extramitochondrial NADH, succinate, orvia the rotenone-insensitive dehydrogenase (see below),all three of which bypass site 1, a maximum of two ATPcould be formed.

186 Chapter 10 / Cellular Respiration: Unlocking the Energy Stored in Photoassimilates

ADP3−

ATP4−

Pi−

OH−

Intermembrane space

Matrix

FIGURE 10.10 The adenine nucleotide transporter. Theone-for-one exchange of mitochondrial ATP and cyto-solic ADP across the inner membrane is driven by themembrane potential. Inorganic phosphate is returnedto the matrix in exchange for hydroxyl ions.

Unlike the chloroplast, most of the ATP synthe-sized in the mitochondrion is utilized elsewhere in thecell. This requires that the ATP be readily transportedout of the organelle. As well, a supply of ADP andinorganic phosphate is required in order to maintainmaximum rates of electron transport and ATP synthe-sis. This is accomplished by two separate translocatorproteins located in the inner membrane. An adeninenucleotide transporter located in the inner mem-brane (Figure 10.10) exchanges ATP and ADP on aone-for-one basis. An inorganic phosphate translocatorexchanges Pi for hydroxyl ions.

10.9 PLANTS CONTAIN SEVERALALTERNATIVE ELECTRONTRANSPORT PATHWAYS

The electron transport chain described above is sharedin essentially the same form by virtually all organisms:

UQpool

4H+

NADH NAD+

NAD(P)H NAD(P)+

NADHdehydrogenase

NAD(P)Hdehydrogenase

Intermembranespace

Matrix

Complex III Complex IV

Cyt b

Cyt c1

Cyt c Cyt c

H2O F1

1/2 O2 + 2H+

H+

ADP + Pi ATP

F0

ATP Synthase

Cyt a/a3

2H+4H+

Innermembrane

FIGURE 10.11 Alternative electron transport pathways in plant mitochondria. Elec-trons entering the chain through the alternative dehydrogenases will pass throughtwo phosphorylating sites rather than three.

plants, animals, and microorganisms. Plant mitochon-dria contain, in addition, several other redox enzymes,at least two of which are unique to plants (Figure 10.11).These enzymes have been discovered largely by virtue oftheir insensitivity to certain classic inhibitors of electrontransport.

10.9.1 PLANT MITOCHONDRIACONTAIN EXTERNALDEHYDROGENASES

Unlike animal mitochondria, plant mitochondria con-tain ‘‘external’’ dehydrogenases that face the intermem-brane space and are capable of oxidizing cytosolicNADH and NADPH respectively (Figure 10.11). Asa consequence, electrons from the oxidation of eithercytosolic NADH or NADPH are donated directly tothe ubiquinone pool. Because the external dehydroge-nase enzymes do not span the membrane, they will nottranslocate protons as Complex I does. Consequently,only two ATP can be formed from the transfer of eachpair of electrons to oxygen.

10.9.2 PLANTS HAVE AROTENONE-INSENSITIVENADH DEHYDROGENASE

The reduction of ubiquinone by Complex I is sensitiveto inhibition by rotenone and amytal. Plants, however,appear to have another NADH dehydrogenase that isinsensitive to both of these electron transport inhibitors.Called the rotenone-insensitive dehydrogenase, thisenzyme will oxidize only internal, or matrix, NADH(Figure 10.11). The enzyme must therefore be locatedon the inner surface of the membrane, facing the matrix.

10.9 Plants Contain Several Alternative Electron Transport Pathways 187

As with the external NADH and NADPH dehydro-genase enzymes, electrons entering the chain via therotenone-insensitive dehydrogenase can generate onlytwo ATP per electron pair.

Thus, the inner membrane of plant mitochon-dria contain four distinct NAD(P)H dehydrogenasesexhibiting different P/O ratios: (1) an internal NADHdehydrogenase; (2) a rotenone-insensitive NADH dehy-drogenase; (3) an external NADH dehydrogenase; and(4) an external NADPH dehydrogenase.

10.9.3 PLANTS EXHIBITCYANIDE-RESISTANTRESPIRATION

Cytochrome c oxidase (Complex IV) is inhibited bycyanide (CN−), carbon monoxide (CO), and azide (N−

3 ).In many animals, all three of these inhibitors completelyinhibit respiratory O2 uptake. By contrast, most plantsor plant tissues show considerable resistance to theseinhibitors. In tissues such as roots and leaves of spinach(Spinacea oleraceae) or pea (Pisum sativum), for example,cyanide-resistant respiration may account for as muchas 40 percent of total respiration. Cyanide-resistant res-piration is, however, sensitive to inhibition by hydrox-amic acid derivatives such as salicylhydroxamic acid(SHAM). This cyanide-resistant, SHAM-sensitive res-piration is attributed to a so-called alternative oxidase(Figure 10.12). The pathway is commonly referredto as the alternative respiratory pathway, or, sim-ply, the alternative pathway. Although the existenceof a cyanide-resistant alternative respiratory pathwayhas been widely accepted for more than a decade,the nature of the oxidase enzyme itself proved diffi-cult to unravel. The enzyme has been difficult to studyby conventional biochemical techniques; the proteinappears to be relatively unstable and loses its acti-vity rapidly upon isolation from the membrane. How-ever, through a molecular biological approach, a gene

encoding the oxidase protein has been cloned, first fromSauromatum guttatum, the voodoo lily, and since fromtobacco, soybean, and other plants. This has led to sig-nificant advances in our understanding of the regulationof the enzyme.

The alternative oxidase is composed of two identicalsubunits (a homodimer) that span the inner mitochon-drial membrane, with the active site facing the matrixside of the membrane. It functions as a ubiquinoneO2 oxidoreductase; that is, it accepts electrons fromthe ubiquinone pool and transfers them directly tooxygen. This is an important characteristic of the alter-native oxidase because it means that electrons processedby this enzyme bypass at least two sites for protonextrusion. Consequently, energy that would otherwisebe conserved as ATP is, in the case of the alterna-tive oxidase, converted to heat instead. Depending onwhether electrons are initially donated to ComplexI, the rotenone-insensitive NADH-dehydrogenase, orsuccinic dehydrogenase, electrons passing through thealternative oxidase will contribute to the synthesis ofone ATP or none (Figure 10.12).

The physiological role of alternative pathway respi-ration is still uncertain. One possible role is thermoge-nesis, a hypothesis based largely on events in the floraldevelopment in certain members of the family Araceae.Just prior to pollination in species such as skunk cab-bage (Symplocarpus foetidus), the tissues of the spadix(the structure that bears both male and female flow-ers) undergoes a surge in oxygen consumption, calleda respiratory crisis. The respiratory crisis is attributedalmost entirely to an increase in alternative pathway res-piration and can elevate the temperature of the spadixby as much as 10◦C above ambient. The high tempera-ture volatilizes certain odoriferous amines (hence, skunkcabbage) that attract insect pollinators. Thermogenesisdoes not, however, appear to be the function of thealternative pathway in roots and leaves. In one studyof an arctic herb, for example, the alternative pathway

FAD

Intermembranespace

Matrix

FMN

Complex I

NADH + H+

NAD+

Complex II

2H+

Succinate Fumarate

H2O1/2 O2 + 2H+

UQpool AOX

AlternativeOxidase

Innermembrane

FIGURE 10.12 The alternative respiratorypathway. Electrons intercepted by the alter-native oxidase (AOX) pass through one or nophosphorylating sites.

188 Chapter 10 / Cellular Respiration: Unlocking the Energy Stored in Photoassimilates

accounts for up to 75 percent of total respiration but, inpart because the heat is rapidly dissipated, accounts forno more than a 0.02◦C rise in leaf temperature.

A second hypothesis to explain the alternative path-way is referred to as the energy overflow hypothesis.This hypothesis is based on two general observations.First, in most tissues the alternative pathway is inoper-ative until the normal cytochrome pathway has becomesaturated. Second, the rate of the alternative pathway canbe increased by increasing the supply of carbohydrateto cells. In spinach (Spinacea oleraceae), for example, thealternative pathway is engaged only after photosynthesishas been in operation for several hours and has built upa supply of carbohydrate. In other words, the alternativepathway is generally engaged when there is an excesssupply of carbohydrate, over and above what is requiredfor metabolism or processed for storage. The functionof the alternative pathway, according to this hypothesis,would be to burn off temporary accumulations of excesscarbon that might otherwise interfere with source-sinkrelationships and inhibit translocation.

In addition, the induction of the alternative oxidaserepresents a mechanism to prevent the overreduction ofthe respiratory electron transport chain, which woulddiminish the probability of superoxide formation andoxidative stress under conditions where ATP con-sumption has been slowed by either low temperature orother stresses.

10.10 MANY SEEDS STORECARBON AS OILS THAT ARECONVERTED TO SUGAR

Although lipids are a principal constituent of membranesand are stored by many tissues, they are not frequentlyused as a source of respiratory carbon. A major excep-tion to this rule is found in germinating seeds, manyof which store large quantities of lipids, principally

triglycerides, as reserve carbon (Table 10.1). Storagelipids are deposited as oil droplets (also called oil bod-ies, oleosomes, or spherosomes), which are normallyfound in storage cells of cotyledons or endosperm.

Since fats and oils are not water soluble, plants areunable to translocate fats and oils through the phloemby pressure flow from seed storage tissues to the elon-gating roots and shoots where the energy and carbon arerequired to support growth. The fatty acids must first beconverted to a form that is more readily translocated bythe aqueous phloem. Usually this is sucrose (or some-times stachyose), which is readily translocated from thestorage cells containing the oil droplets to the embryowhere the sucrose is metabolized. Complete conver-sion of triglycerides to sucrose is a complex process,involving the interaction of the oil bodies, glyoxysomes,mitochondria, and the cytosol (Figure 10.13).

We can summarize the conversion of triglyceridesto sucrose as follows. The first step is the hydrolysisof triglycerides to free fatty acids and glycerol. Thisis accomplished through the action of lipase enzymes,which probably act at the surface of the oil droplet. Thefatty acid then enters the glyoxysome, an organellesimilar in structure to the peroxisome found in leavesbut with many different enzymes. In the glyoxysome,the fatty acid undergoes β-oxidation; the fatty acidchain is cleaved at every second carbon, resulting in theformation of acetyl-CoA.

Some of the acetyl-CoA combines with oxaloac-etate (originating in the mitochondrion) to form cit-rate (6 carbons) in what is known as the glyoxylatecycle. The citrate in turn is converted to isocitrate,which then breaks down into one molecule of suc-cinate (4 carbons) and one molecule of glyoxylate(2 carbons). The succinate returns to the mitochon-drion where it enters the citric acid cycle, regeneratingoxaloacetate, which is necessary to keep the glyoxy-late cycle turning. Glyoxylate combines with anotheracetyl-CoA to produce malate. The malate then entersthe cytosol where it is first oxidized to oxaloacetate and

TABLE 10.1 Approximate lipid content of selected seeds.

Oil ContentSpecies (% dry weight)

Macadamia nut Macadamia ternifolia 75Hazel nut Coryllus avellana 65Safflower Carthmus tinctoris 50Oil palm Elaeis guineensis 50Canola Brassica napus 45Castor bean Ricinus communis 45Sunflower Helianthus annum 40Maize Zea mays 5

Hopkins & Huner.

10.11 Respiration Provides Carbon Skeletons for Biosynthesis 189

Malate

TriglycerideOIL BODY

GLYOXYSOME MITOCHONDRION

GlyoxylateCycle

CYTOSOL

(lipase)

Fatty acid+ Glycerol

Fatty acid

Oxaloacetate Oxaloacetate

Acetyl-CoA

Citrate

Isocitrate

Glycerol

NADH

NADH

NADH

NADHNADH

NAD+

NAD+

NAD+

NAD+

NAD+

DHAPα - Glycerol - P

β - oxidation

Fructose - 1, 6 - BP

Fructose - 6 - P + Glucose - 1 - P

Sucrose

Glyoxylate Succinate Succinate

Fumarate

Malate

Reverseglycolysis

Malate

Malate Oxaloacetate

ADPATP

PEP

CO2

FIGURE 10.13 Lipid catabolism, the glyoxylate cycle, andgluconeogenesis.

decarboxylated to phosphoenolpyruvate (PEP). Theglyoxylate cycle thus involves enzymes of both the gly-oxysome and the mitochondrion. Two enzymes of thecycle are unique to plants: isocitrate lyase, which con-verts isocitrate to succinate plus glyoxylate, and malatesynthase, which condenses an acetyl group with gly-oxylate to form malate. The malate is then translocatedfrom the glyoxysome into the cytosol where it is quicklyoxidized to oxaloacetate by the enzyme malate dehy-drogenase. The overall effect of the glyoxylate cycleis to catalyze the formation of oxaloacetate from twomolecules of acetyl-CoA.

In the cytosol, oxaloacetate derived from theglyoxylate cycle is decarboxylated via the enzymephosphoenolpyruvate carboxykinase (PEPCK) toform phosphoenolpyruvate (PEP). Through a sequenceof reactions that is essentially a reversal of glycolysis,PEP is converted to glucose. The conversion of PEPto glucose by a reversal of glycolysis is known as glu-coneogenesis. Gluconeogenesis utilizes the enzymesof glycolysis, with significant differences. The gly-colytic phosphofructokinase and hexokinase reactions(Figure 10.4) are effectively irreversible—their free

energy changes are highly unfavorable in the directionof glucose synthesis. During gluconeogenesis, thesereactions are replaced by reactions that make glucosesynthesis more thermodynamically favorable. Theconversion of fructose-1,6-bisphosphate to fructose-6-Pis catalyzed by cytosolic fructose-1,6-bisphosphataseand the conversion of glucose-6-P to glucose iscatalyzed by glucose-6-phosphatase. These differencesare significant because they allow both directions tobe thermodynamically favorable, yet be independentlyregulated. One direction can be activated while theother is inhibited, thus avoiding what might otherwiseend up as a futile cycle. The glycerol resulting fromlipase action in the oil droplet also enters the cytosol,where it is first phosphorylated with ATP to formα-glycerolphosphate and then oxidized to dihydroxy-acetone phosphate (DHAP). The DHAP can also beconverted to sucrose by reversal of glycolysis. Some ofthe energy stored in triglycerides is conserved in thesucrose formed by gluconeogenesis, but β-oxidationof fatty acids in the glyoxysome also produces a largeamount of NADH. The glyoxysome is unable toreoxidize NADH directly, but it can be used to reduceoxaloacetate to malate (Figure 10.13). The malate thenmoves into the mitochondrion where it is reoxidized bymalate dehydrogenase. Malate thus serves as a shuttle,carrying reducing equivalents between the glyoxysomeand the mitochondrion. Reoxidation of malate insidethe mitochondrion yields NADH, which can then enterthe electron transport chain and drive ATP synthesis.

10.11 RESPIRATION PROVIDESCARBON SKELETONS FORBIOSYNTHESIS

Before leaving the subject of cellular respiration, it isimportant to note that production of reducing poten-tial and ATP is not the sole purpose of the respiratorypathways. In addition to energy, the synthesis of nucleicacids, protein, cellulose, and all other cellular moleculesrequires carbon skeletons as well. As noted at the begin-ning of this chapter, respiration also serves to modifythe carbon skeletons of storage compounds to formthese basic building blocks of cell structure. A few ofthe more important building blocks that can be formedfrom intermediates in glycolysis and the citric acid cycleare represented in Figure 10.14.

The withdrawal of glycolytic and citric acid cycleintermediates for the synthesis of other moleculesmeans, of course, that not all of the respiratory substratewill be fully oxidized to CO2 and water. The flowof carbon through respiration no doubt represents abalance between the metabolic demands of the cell forATP to drive various energy-consuming functions onthe one hand and demands for the reducing equivalents

190 Chapter 10 / Cellular Respiration: Unlocking the Energy Stored in Photoassimilates

Nucleic acids

ATPNAD

Cytokinins

Starch

Triose - P

Pentose - P Glucose - 6 - P Cellulose

Shikimic acid

Glycerol TriglyceridesPhospholipids

Phosphoenolpyruvate

Amino acids

Protein

Pyruvate Alanine Protein

ChlorophyllCarotenoidsGibberellinsTerpenes

AcetylCoA

Citrate

α - Ketoglutarate Porphyrins

Glutarate

OxaloacetateCitric Acid

Cycle

Isoprenoids

Fatty acids

Other amino acids

Protein

Auxin Otheraminoacids

ChlorophyllCytochromePhytochrome

Aspartate

Other amino acidsAlkaloidsProtein

FIGURE 10.14 The role of respiration in biosynthesis. Intermediates in glycolysis andthe citric acid cycle are drawn off to serve as building blocks for the synthesis of cel-lular molecules. Carbon in the cycle is maintained by the synthesis of oxaloacetatethrough anaplerotic reactions. This scheme is incomplete and is intended only togive some indication of the importance of these two schemes in biosynthesis.

and carbon skeletons required to build cell structure onthe other.

It is also important to note that during periods ofactive synthesis, diversion of carbon from the citric acidcycle for synthetic reactions will lead to a significantreduction in the level of oxaloacetate. These syntheticreactions require not only carbon, but energy in theform of reducing potential and ATP as well. Withoutsome means of compensating for this loss of oxaloac-etate, the cycle will slow down or, in the extreme case,come to a complete halt and energy production will beimpaired. This eventuality is precluded by the actionof two cytosolic enzymes: phosphoenolpyruvate (PEP)carboxylase (see Chapter 15) and malate dehydrogenase.All plants, not just those with C4 photosynthetic activity(see Chapter 15), have some level of PEP carboxylaseactivity that converts phosphoenolpyruvate (PEP) intooxaloacetate:

PEP + HCO−3 → oxaloacetate (10.9)

In this case the PEP is derived from glycolysis.Although there is some evidence that oxaloacetate

may be translocated directly into the mitochondrion, it

is more likely that oxaloacetate is quickly reduced tomalate by the action of cytosolic malate dehydrogenase:

oxaloacetate + NADH → malate + NAD+ (10.10)

The malate would then pass into the mitochondrion, viaa malate (or dicarboxylate) translocator, where it is reox-idized to oxaloacetate by the action of a mitochondrialmalate dehydrogenase:

malate + NAD+ → oxaloacetate + NADH (10.11)

The replenishment of oxaloacetate in this way is anexample of a ‘‘filling-up’’ mechanism or anapleroticpathway. Thus carbon from glycolysis is delivered to thecitric acid cycle through two separate but equally impor-tant streams: (1) to citrate via pyruvate and acetyl-CoAand (2) from PEP via oxaloacetate and malate to com-pensate for carbon ‘‘lost’’ to synthesis. Anapleroticreactions such as the latter help to ensure that diver-sion of carbon for synthesis does not adversely influencethe overall carbon balance between energy-generatingcatabolic reactions and biosynthetic anabolic reactions.

In addition to the normal enzymes of the citric acidcycle, plant mitochondria tend to have significant levels

10.12 Respiratory Rate Varies with Development and Metabolic State 191

of NAD+ -malic enzyme, which catalyzes the oxidativedecarboxylation of malate:

malate+NAD+ → pyruvate+CO2+NADH (10.12)

The pyruvate may be further metabolized by pyruvatedehydrogenase to acetyl-CoA and from there enter thecitric acid cycle. Thus the mitochondrial pool of malatemay replenish the citric acid intermediates througheither oxaloacetate or pyruvate.

In addition to serving an anaplerotic role, the uptakeand oxidation of malate by mitochondria via either malicenzyme or malate dehydrogenase also provides an alter-native pathway for metabolizing malate. This alternativepathway may be particularly significant in plants suchas those of the family Crassulaceae (see Chapter 15)and others that store significant levels of malate in theirvacuoles. Finally, it should be noted that diversion ofpyruvate through oxaloacetate and malate bypasses thepyruvate kinase step in glycolysis (Figure 10.5, reaction6) and thus reduces the yield of ATP by one. This reduc-tion is, however, offset by gains achieved by reductionof malate in the cytosol and its subsequent reoxida-tion in the mitochondrion. This sequence of reactionseffectively shuttles extramitochondrial NADH (gener-ated during glycolysis) into the mitochondrion, whereit can be used to generate three molecules of ATP.This is a gain of one ATP over the two ATP gener-ated via the NADH-reductase route described earlierfor extramitochondrial NADH.

10.12 RESPIRATORY RATE VARIESWITH DEVELOPMENT ANDMETABOLIC STATE

The study of respiration at the level of individual organsor the whole plant becomes much more difficult than itis for the study of individual cells. Whole plant respi-ration is normally studied by measuring the uptake ofoxygen or the evolution of CO2, but respiration ratesobtained this way are highly variable. The balance of O2and CO2 exchange is dependent on the substrate beingrespired and the balance of fermentation, citric acidcycle, and alternative pathway activities at any pointin time. In addition, respiration rates differ betweenorgans, change with age and developmental state, andare markedly influenced by temperature, oxygen, salts,and other environmental factors. Nevertheless, the studyof respiration at the organ and plant level is a field ofactive study. Understanding respiration at this levelhas important implications for the plant physiologistinterested in growth and development, for the physio-logical ecologist interested in plant biomass production,and the agricultural scientist because of its impact onproductivity and yield.

Rel

ativ

e re

spir

atio

n

Age

Maturation Senescence

Climacteric rise

Rapidgrowth

FIGURE 10.15 Respiratory rate as a function of age. Thistype of curve applies generally to most herbaceous plants,tissues, and organs. The magnitude of the climactericwill vary—some organs exhibit little or no climactericrise.

As a general rule, respiratory rate is a reflectionof metabolic demands. Younger plants, organs, or tis-sues respire more rapidly than older plants, organs, ortissues (Figure 10.15). The rapid rate of respirationduring early stages of growth is presumably related tosynthetic requirements of rapidly dividing and enlarg-ing cells. As the plant or organ ages and approachesmaturity, growth and its associated metabolic demandsdecline. Many organs, especially leaves and some fruits,experience a transient rise in respiration, called a cli-macteric, that marks the onset of senescence and thedegenerative changes that precede death. Typically theclimacteric rise in O2 consumption is accompanied bya decline in oxidative phosphorylation, indicating thatATP production is no longer tightly coupled to elec-tron transport. The respiration rate of woody stemsand branches, expressed on a weight or mass basis, alsodeclines as they grow. This is because as the diameterincreases, the relative proportion of nonrespiring woodytissue also increases.

Carbon lost to the plant due to respiration canrepresent a significant proportion of the availablecarbon. Actual respiration rates for plant tissues rangein the extreme between barely detectable (0.005 μmolCO2 gW−1

d h−1) in dormant seeds to 1000 μmolCO2 gW−1

d h−1 or more in the spadix of skunk cabbageduring the respiratory crisis. More typically, rates forvegetative tissues range from 10 to 200 (mol CO2gW−1

d h−1 (Table 10.2). This may represent a con-siderable fraction of the carbon assimilated byphotosynthesis during a 24-hour period. (Recall thatphotosynthesis occurs only during daylight hours, butrespiration, especially of roots and similar tissues, isongoing 24 hours a day.) On the average, 30 to 60percent of daily photoassimilate is lost as respiratoryCO2. In tropical rainforest species, probably because ofaccelerated enzymic activities at higher temperatures,this loss may exceed 70 percent. Of the total daily

192 Chapter 10 / Cellular Respiration: Unlocking the Energy Stored in Photoassimilates

TABLE 10.2 Approximate specific dark respira-tion rates at 20◦C for crop species, deciduous foli-age, and conifers.

Specific Respiration Rateμ mol CO2 evolved g−1

dry mass h−1

Crops 70–180Deciduous foliage

(sun leaves) 70–90(shade leaves) 20–45

Conifers 4–25

Hopkins & Huner.

carbon loss in any given plant, some 30 to 70 percentis accounted for by respiration in the roots alone, that‘‘hidden half’’ of the plant.

10.13 RESPIRATION RATESRESPOND TOENVIRONMENTALCONDITIONS

10.13.1 LIGHT

The effects of light on mitochondrial respiration havebeen the subject of considerable debate for some time.Traditionally, photosynthesis investigators and cropphysiologists have tacitly assumed that respiration con-tinues in the light at a rate comparable to that inthe dark. The true rate of photosynthesis is thereforetaken as equal to the apparent rate (measured as CO2uptake) plus the rate of respiration (CO2 evolved) in thedark. However, attempts to study respiration in greenleaves have led to alternative and conflicting conclusions.These range from complete inhibition of mitochondrialactivities, to partial operation of the citric acid cycle, orto stimulation of respiration by light. The problem liesin the difficulty of measuring respiration during a periodwhen gas exchange is dominated by the overwhelmingflux of CO2 and O2 due to photosynthesis, the recyclingof CO2 within the leaf, and the exchange of metabolitesby chloroplasts and mitochondria.

Light effects on respiration during a subsequentdark period have been demonstrated. For example, darkrespiratory rates in leaves adapted to full sun (sun leaves)are generally higher than those of leaves of the samespecies adapted to shade (shade leaves) (Table 10.2). Aswell, the rate is consistently higher in mature leaves ofshade-intolerant species than in shade-tolerant species.Indeed, a reduced respiratory rate appears to be a fairlyconsistent response to low irradiance. This is prob-ably related to lower growth rates also observed inshade-grown plants, but it is not known which is cause

and which is effect. It is projected that low respiratoryand growth rates may confer a survival advantage underconditions of deep shade. The basis for light regula-tion of respiratory rate is unknown, although some havesuggested that low respiratory rates under conditions oflow irradiance may reflect availability of substrate. Forexample, the respiration rate 1 to 2 hours following aperiod of active photosynthesis is higher than after along dark period.

Other experiments, however, have shown that darkrespiratory rates do not correlate with CO2 supplyduring the previous light period. This seems to sug-gest a more direct effect on respiration. The pyruvatedehydrogenase complex (PDH) of the mitochondrioncan exist either in an active, nonphosphorylated formor an inactive, phosphorylated form (Figure 10.16). Ithas been shown that the photorespiratory-generatedNH+

4 (Chapter 8) stimulates the phosphorylation ofthe mitochondrial pyruvate dehydrogenase complex,thereby inhibiting the rate of respiration in the lightby decreasing the rate at which acetyl-CoA is generatedfor the CAC. Light regulation of respiration remains acontroversial issue in plant physiology.

10.13.2 TEMPERATURE

One of the most commonly applied quantitative mea-sures used to describe the effect of temperature on aprocess is the temperature coefficient, or Q10, givenby the expression:

Q10 = rate at(t + 10)◦Crate at t◦C

(10.13)

At temperatures between 5◦C and about 25◦C or 30◦C,respiration rises exponentially with temperature and the

+

PDH Kinase

PDH(inactive)

PDH(active) Phospho-PDH

Phosphatase

Pi

ADP

P

NH4+

ATP

FIGURE 10.16 Regulation of mitochondrial pyruvatedehydrogenase complex. The pyruvate dehydrogenasecomplex (PDH) exists in two forms: an active, nonphos-phorylated form and an inactive, phosphorylated form.The reversible interconversion of the active and inac-tive forms of this enzyme is the result of the activity ofthe PDH kinase, which consumes ATP to phosphorylatePDH, and the activity of the phospho-PDH phosphatase,which dephosphorylates PDH. Ammonium ion (NH+

4 )stimulates the PDH kinase and hence inactivates thePDH complex.

Summary 193

Q10 value is approximately 2.0 in many but not all plants(see Chapter 14). Within this temperature range, a dou-bling of rate for every 10◦C rise in temperature is typicalof enzymic reactions. At temperatures above 30◦C, theQ10 in most plants begins to fall off as substrate avail-ability becomes limiting. In particular, the solubility ofO2 declines as temperatures increase and the diffusionrate (with a Q10 close to 1) does not increase suffi-ciently to compensate. As temperatures approach 50◦Cto 60◦C, thermal denaturation of respiratory enzymesand damage to membranes bring respiration to a halt.

Some investigators have observed differences inthe rate of respiration in tropical, temperate, and arc-tic species at different temperatures. For example, therespiration rate of leaves of tropical plants at 30◦Cis about the same as that of arctic species at 10◦C.The temperature coefficient (Q10) for respiration isthe same in both cases and there is no evidence ofintrinsic differences in the biochemistry of respiration.It is likely that the differences reflect differences intemperature optima for growth of arctic and tropi-cal species—optima determined by factors other thanrespiration—and the consequent metabolic demand forATP. A detailed discussion of the effects of temperaturestress and acclimation on respiration will be focused inChapters 13 and 14.

10.13.3 OXYGEN AVAILABILITY

As the terminal electron acceptor, oxygen availabilityis obviously an important factor in determining respi-ration rate. The oxygen content of the atmosphere isrelatively stable at about 21 percent O2. The equilib-rium concentration of oxygen in air-saturated water,including the cytosol, is approximately 250 μM. How-ever, cytochrome c oxidase has a very high affinity foroxygen with a Km (see Chapter 8, Box 8.1) less than1 μM. Under normal circumstances, oxygen is rarely alimiting factor.

There are some situations, however, where oxygenavailability may become a significant factor. One is inbulky tissues with low surface-to-volume ratios, suchas potato tubers and similar storage tissues, where thediffusion of oxygen may be slow enough to restrictrespiration. This may not be a serious problem, however.A significant volume—as much as 40 percent—of rootsand similar tissues may be occupied by intercellular airspaces that aid in the rapid distribution of O2 absorbedfrom the soil or, in some cases, from the aerial portionsof the plant. Plants are most likely to experience oxygendeficits during periods of flooding, when air in the largepore spaces of the soil is displaced by water, therebydecreasing the oxygen supply to the roots. For similarreasons, plants grown in hydroponic culture must beaerated to maintain adequate oxygen levels in the vicinityof the roots (Chapter 3).

SUMMARY

Cellular respiration consists of a series of interde-pendent pathways by which carbohydrate and othermolecules are oxidized for the purpose of retrievingthe energy stored in photosynthesis and to obtain thecarbon skeletons that serve as precursors for othermolecules used in the growth and maintenance ofthe cell. Plants store excess photosynthate either asstarch, a long linear or branched polymer of glucose,in the chloroplast stroma or as fructans, a polymer offructose, in the vacuole. Storage carbohydrates suchas starch and fructans are enzymatically degraded toglucose or fructose, which then enter the cytosolichexose phosphate pool as either glucose-1-phosphate,glucose-6-phosphate, or fructose-6-phosphate. Hexosephosphates exit the hexose-P pool by conversion tofructose-1,6-bisphosphate (FBP). FBP is subsequentlyconverted to triose-P, which is the starting point forglycolysis, a series of reactions that ultimately producepyruvate. In the process, a small amount of ATP andreducing potential is generated. The intermediatesin glycolysis are three-carbon sugars, many of whichare precursors to triglycerides and amino acids. Pre-cursors with four and five carbons are produced byan alternative route for glucose metabolism called theoxidative pentose phosphate pathway. The oxidativepentose phosphate pathway also produces NADPH(as opposed to NADH), which provides reducingpotential when required for biosynthetic reactions inplants.

The fate of pyruvate depends on the availabilityof oxygen. In an anaerobic environment, pyruvate isreduced (usually to ethanol), while in the presence ofoxygen pyruvate is first oxidized to acetyl-CoA andcarbon dioxide. The acetate group is then furtheroxidized to carbon dioxide and water through the cit-ric acid cycle (CAC). The CAC enzymes are locatedpredominantly in the matrix of the mitochondrion.Altogether, eight enzyme-catalyzed steps degrade theacetate group to carbon dioxide and water. The cycleincludes four oxidations that yield NADH at threesteps and FADH2 at another. One molecule of ATPis generated in a substrate-level phosphorylation andthe original acetate acceptor, oxaloacetate, is regener-ated, which allows the cycle to continue. The NADHand FADH2 produced in the CAC are oxidized via anelectron transport chain found in the mitochondrialinner membrane. The chain consists of four multi-protein complexes linked by mobile electron carriers.The final complex, cytochrome oxidase, transfers theelectrons to molecular oxygen, forming water. At threepoints in the chain, the free energy drop associatedwith electron transport is used to establish a protongradient across the membrane. As in the chloroplasts,this proton gradient is used to drive ATP synthesis.

194 Chapter 10 / Cellular Respiration: Unlocking the Energy Stored in Photoassimilates

The CAC and electron transport chain are virtuallyidentical in all organisms. Plants, however, have analternative oxidase that intercepts electrons early in thechain, thus bypassing two of the three proton-pumpingsites. When using this route, at least two-thirds lessATP is formed and much of the electron energy isconverted to heat. The alternative oxidase, at least insome plants, has been associated with thermogenesis,particularly in certain members of the Araceae wherethe higher temperatures volatilize amines that appearto attract insect pollinators. The alternative oxidasemay also serve to ‘‘burn off’’ excess carbohydrate aswell as protect against oxidative stress by preventingthe overreduction of the respiratory electron transportchain.

Many seeds store carbon as fats and oils, whichmust first be converted to sugar in order to be respired.After the fatty acids are broken down into acetyl-CoAunits, a complex series of reactions involving enzymesof the mitochondrion, the glyoxysome, and the cytosolconvert the acetate units to phosphoenolpyruvate. Thepyruvate is then converted to glucose by gluconeogen-esis, a process that is essentially a reversal of glycolysis.

The respiratory rate of whole plants and organsvaries widely with age, metabolic state, and environ-mental conditions.

CHAPTER REVIEW

1. Compare respiration and fermentation. Aer-obic organisms are generally much largerthan anaerobic organisms. Can you suggesthow this may be related to respiration?

2. Phosphorous plays an important role in respira-tion, photosynthesis, and metabolism generally.What is this role?

3. Oils are a common storage form in seeds, particu-larly small seeds. What advantage does this offerthe seed?

4. There are a number of similarities betweenchloroplasts and mitochondria. Comparethese two organelles from the perspective of:

(a) ultrastructure and biochemical compartmenta-tion;

(b) organization of the electron transport chain;(c) proton motive force (pmf) and ATP synthesis;(d) alternative oxidases.

5. How does cyanide-resistant respiration differfrom normal respiration? Of what value mightcyanide-resistant respiration be to plants?

6. Discussions of respiration most often emphasizeenergy retrieval and ATP production. What othervery important metabolic role(s) does respirationfulfill?

7. Give an example of an anaplerotic path-way. What is the role of this pathway?

FURTHER READING

Buchanan, B. B., W. Gruissem, R. L. Jones. 2000.Biochemistry and Molecular Biology of Plants. Rockville,MD: American Society of Plant Physiologists.

Foyer, C. H., S. Ferrario-Mery, S. C. Huber. 2000. Reg-ulation of carbon fluxes in the cytosol: Coordinationof sucrose synthesis, nitrate reduction and organic andamino acid biosynthesis. In: Photosynthesis: Physiology andMetabolism R. C. Leegood, T. D. Sharkey, S. von Caem-merer, (eds.), Advances in Photosynthesis and Respiration,Vol. 9, pp. 177–203. Dordrecht: Kluwer.

Gardestrom, P., A. U. Igamberdiev, A. S. Raghavendra. 2002.Mitochondrial functions in the light and significance tocarbon-nitrogen interactions. In Photosynthesis: Physiologyand Metabolism R. C. Leegood, T. D. Sharkey, S. vonCaemmerer (eds.), Advances in Photosynthesis and Respira-tion, Vol. 9, pp. 151–172. Dordrecht: Kluwer.

Vanlerberghe, G. C., L. McIntosh. 1997. Alternative oxidase:From gene to function. Annual Review of Plant Physiologyand Plant Molecular Biology 48: 703–734.

Vanlerberghe, G. C., S. H. Ordog. 2002. Alternative oxidase:Integrating carbon metabolism and electron transport inplant respiration. Photosynthesis: Physiology and MetabolismIn: R. C. Leegood, T. D. Sharkey, S. von Caemmerer(eds.), Advances in Photosynthesis and Respiration, Vol. 9,pp. 173–191. Dordrecht: Kluwer.

fdred

fdox

N2 + 8H+

2NH3 + H2

Feox

Fered

Fe protein

MoFered

MoFeox

MoFe protein

MgADP + PiMgATP2-

11Nitrogen Assimilation

On a dry-weight basis, nitrogen is the fourth mostabundant nutrient element in plants. It is an essen-tial constituent of proteins, nucleic acids, hormones,chlorophyll, and a variety of other important primaryand secondary plant constituents. Most plants obtain thebulk of their nitrogen from the soil in the form of eithernitrate (NO−

3 ) or ammonium (NH+4 ), but the supply

of nitrogen in the soil pool is limited and plants mustcompete with a variety of soil microorganisms for whatnitrogen is available. As a result, nitrogen is often a lim-iting nutrient for plants, in both natural and agriculturalecosystems.

The bulk of the atmosphere, 78 percent by volume,consists of molecular nitrogen (N2, or dinitrogen),an odorless, colorless gas. In spite of its abundance,however, higher plants are unable to convert dinitro-gen into a biologically useful form. The two nitrogenatoms in dinitrogen are joined by an exceptionally sta-ble bond (N ≡ N) and plants do not have the enzymethat will reduce this triple covalent bond. Only certainprokaryote species are able to carry out this impor-tant reaction. This situation presents plants with aunique problem with respect to the uptake and assim-ilation of nitrogen; plants must depend on prokaryoteorganisms to convert atmospheric dinitrogen into ausable form. The nature of this problem and thesolutions that have evolved are the subject of thischapter.

The principal topics discussed in this chapterinclude:

• a review of the nitrogen cycle: the flow of nitrogenbetween three major global nitrogen pools,

• the biology and biochemistry of biologicalnitrogen-fixing systems, and

• pathways for assimilation of ammonium and nitratenitrogen by plants.

11.1 THE NITROGEN CYCLE:A COMPLEX PATTERNOF EXCHANGE

The global nitrogen supply is generally distributedbetween three major pools: the atmospheric pool, thesoil (and associated groundwater) pool, and nitrogencontained within the biomass. Central to the idea ofa nitrogen cycle (Figure 11.1) is the pool of nitrogenfound in the soil. Nitrogen from the soil pool enters thebiomass principally in the form of nitrate (NO−

3 ) takenup by plants and microorganisms. Once assimilated,nitrate nitrogen is converted to organic nitrogen in theform of amino acids and other nitrogenous buildingblocks of proteins and other macromolecules. Nitrogenmoves further up the food chain when animals consume

195

196 Chapter 11 / Nitrogen Assimilation

Atmospheric N2

BiologicalN2 fixation

IndustrialN2 fixation

ElectricalN2 fixation

Denitrification

NH3 NO2– –NO3

(Ammonification) (Uptake)

Plant biomassDecaying biomass

Animal biomass

Soil N Pool

FIGURE 11.1 The nitrogen cycle, illustrating relation-ships between the three principal nitrogen pools: atmo-spheric, soil, and biomass.

plants. Nitrogen is returned to the soil through animalwastes or the death and subsequent decomposition of allorganisms.

11.1.1 AMMONIFICATION,NITRIFICATION,AND DENITRIFICATION AREESSENTIAL PROCESSESIN THE NITROGEN CYCLE

In the process of decomposition, organic nitrogen isconverted to ammonia by a variety of microorgan-isms including the fungi. This process is known asammonification (Figure 11.1). Some of the ammoniamay volatilize and reenter the atmosphere, but most ofit is recycled to nitrate by soil bacteria. The first step inthe formation of nitrate is the oxidization of ammoniato nitrite (NO−

2 ) by bacteria of the genera Nitrosomonasor Nitrococcus. Nitrite is further oxidized to nitrate bymembers of the genus Nitrobacter. These two groupsare known as nitrifying bacteria and the result of theiractivities is called nitrification. Nitrifying bacteria arechemoautotrophs; that is, the energy obtained by oxi-dizing inorganic substances such as ammonium or nitriteis used to convert carbon dioxide to organic carbon.

In taking up nitrate from the soil, plants mustcompete with bacteria known as denitrifiers (e.g.,Thiobacillus denitrificans). By the process of denitrifica-tion, these bacteria reduce nitrate to dinitrogen, whichis then returned to the atmosphere. Estimates for theamount of nitrogen lost to the atmosphere by denitrifi-cation range from 93 million to 190 million metric tonsannually.

11.2 BIOLOGICAL NITROGENFIXATION IS EXCLUSIVELYPROKARYOTIC

The loss of nitrogen from the soil pool through denitri-fication is largely offset by additions to the pool throughconversion of atmospheric dinitrogen to a combinedor fixed form. The process of reducing dinitrogen toammonia is known as nitrogen fixation or dinitrogenfixation. Just how much dinitrogen is fixed on a globalbasis is difficult to determine with any accuracy. Figureson the order of 200 to 250 million metric tons annuallyhave been suggested. Approximately 10 percent of thedinitrogen fixed annually is accounted for by nitrogenoxides in the atmosphere. Lightning strikes and ultravi-olet radiation each provide sufficient energy to convertdinitrogen to atmospheric nitrogen oxides (NO, N2O)(Figure 11.1). Another 30 percent of the total dinitro-gen fixed, 80 million metric tons, is accounted for byindustrial nitrogen fixation for the production of agri-cultural fertilizers through the Haber-Bosch process(Figure 11.1). However, by far, the bulk of the nitrogenfixed on a global scale, about 60 percent or 150 to 190million metric tons annually, is accounted for by thereduction of dinitrogen to ammonia by living organisms(Figure 11.1).

Plants are eukaryotic organisms, distinguished bythe presence of a membrane-limited nuclear compart-ment. Eukaryotic organisms are unable to fix dinitrogenbecause they do not have the appropriate biochemicalmachinery. Bacteria and cyanobacteria are prokaryoticorganisms; the genetic material is not contained within amembrane-limited organelle. Nitrogen fixation is a pro-karyote domain, because only prokaryote organisms havethe enzyme complex, called dinitrogenase, that cata-lyzes the reduction of dinitrogen to ammonia. Simpleas this may seem, biological nitrogen fixation turns outto be a complex biochemical and physiological process.

Prokaryotes that fix nitrogen, called nitrogen-fixers, include both free-living organisms and thosethat form symbiotic associations with other organisms.

11.2.1 SOME NITROGEN-FIXINGBACTERIA ARE FREE-LIVINGORGANISMS

Free-living, nitrogen-fixing bacteria are widespread.Their habitats include marine and freshwater sedi-ments, soils, leaf, and bark surfaces, and the intestinaltracts of various animals. Although some species areaerobic (e.g., Azotobacter, Beijerinckia), most will fix dini-trogen only under anaerobic conditions or in the pres-ence of very-low-oxygen partial pressures (a conditionknown as microaerobic). These include both nonpho-tosynthetic genera (Clostridium, Bacillus, Klebsiella) and

11.3 Legumes Exhibit Symbiotic Nitrogen Fixation 197

photosynthetic genera (Chromatium, Rhodospirillum) ofbacteria. In addition to the bacteria, several genera ofcyanobacteria (principally Anabaena, Nostoc, Lyngbia, andCalothrix) are represented by nitrogen-fixing species.

Although free-living nitrogen-fixing organisms arewidespread, most grow slowly and, except for the photo-synthetic species, tend to be confined to habitats rich inorganic carbon. Because a high proportion of their res-piratory energy is required to fix dinitrogen, less energyis therefore available for growth.

11.2.2 SYMBIOTIC NITROGENFIXATION INVOLVES SPECIFICASSOCIATIONS BETWEENBACTERIA AND PLANTS

Several types of symbiotic nitrogen-fixing associationsare known, including the well-known associationbetween various species of bacteria and leguminousplants. Some of the more important associations arelisted in Table 11.1. In symbiotic associations the plant isidentified as the host and the microbial partner is knownas the microsymbiont. The most common form of sym-biotic association results in the formation of enlarged,multicellular structures, called nodules, on the root(or occasionally the stem) of the host plant (Figure 11.2).In the case of legumes,1 the microsymbiont is abacterium of one of three genera: Rhizobium, Bradyrhi-zobium, or Azorhizobium. Collectively, these organisms

TABLE 11.1 Some examples of specificity inrhizobia-legume symbiosis.

Bacterium Host

Azorhizobium SesbaniaBradyrhizobium japonicum Glycine (soybean)Rhizobium meliloti Medicago (alfalfa),

Melilotus (sweet clover)Rhizobium leguminosarum

biovar viciae Lathyrus (sweet pea),Lens (lentil),Pisum (garden pea),Vicia (vetch, broad bean)

biovar trifolii Trifolium (clover)biovar phaseoli Phaseolus (bean)

Rhizobium loti Lotus (bird’s-foot trefoil)

1The legumes are a heterogeneous group traditionallyassigned to the family Leguminosae. Modern treatments splitthe group into three families: Mimosaceae, Caesalpiniaceae,and Fabaceae (S. B. Jones, A. E. Luchsinger, Plant Systematics,New York: McGraw-Hill, 1986). Most of the economicallyimportant, nitrogen-fixing legumes are assigned to theFabaceae.

FIGURE 11.2 Nitrogen-fixing nodules on roots of soybean(Glycine max).

are referred to as rhizobia. Curiously, only one non-leguminous genus, Parasponia (of the family Ulmaceae),is known to form root nodules with a rhizobia symbiont.

The rhizobia are further divided into species andsubgroups called biovars (a biological variety) accordingto their host range (Table 11.1). Most rhizobia arerestricted to nodulation with a limited number of hostplants while others are highly specific, infecting onlyone host species.

Nodules are also found in certain nonleguminousplants such as alder (Alnus), bayberry (Myrica), Australianpine (Casuarina), some members of the family Rosaceae,and certain tropical grasses. However, the microsym-biont in these nonleguminous nodules is a filamentousbacterium (Frankia) of the group actinomycetes. BothRhizobium and Frankia live freely in the soil but fixdinitrogen only when in symbiotic association with anappropriate host plant.

A limited number of non-nodule-forming associa-tions have been studied, such as that between Azolla andthe cyanobacterium Anabaena. Azolla is a small aquaticfern that harbors Anabaena in pockets within its leaves.In southeast Asia, Azolla has proven useful as greenmanure in the rice paddy fields where it is either appliedas a manure or co-cultivated along with the rice plants.Because more than 75 percent of the rice acreage consistsof flooded fields, free-living cyanobacteria and anaero-bic bacteria may also make a significant contribution.These practices have allowed Asian rice farmers to main-tain high productivity for centuries without resorting toadded chemical fertilizers.

11.3 LEGUMES EXHIBITSYMBIOTIC NITROGENFIXATION

It is generally agreed that symbiotic nitrogen fixers,particularly legumes, contribute substantially morenitrogen to the soil pool than do free-living bacteria.

198 Chapter 11 / Nitrogen Assimilation

Typically a hectare of legume-Rhizobium associationwill fix 25 to 60 kg of dinitrogen annually, whilenonsymbiotic organisms fix less than 5 kg ha−1. Thereare over 17,000 species of legumes. Even though only20 percent have been examined for nodulation, 90percent of those examined do form nodules. Given theobvious significance of the legume symbiosis to thenitrogen cycle, it is worth examining in some detail.

11.3.1 RHIZOBIA INFECT THE HOSTROOTS, WHICH INDUCESNODULE DEVELOPMENT

The sequence of events beginning with bacterial infec-tion of the root and ending with formation of mature,nitrogen-fixing nodules has been studied extensivelyin the legumes, historically from the morphologicalperspective and more recently from the biochemical/molecular genetic perspective. Overall the processinvolves a sequence of multiple interactions between thebacteria and the host roots. In effect, the rhizobia andthe roots of the prospective host plant establish a dia-logue in the form of chemical messages passed betweenthe two partners. Based on studies carried out primarilywith Glycine, Trifolium, and Pisum, as many as nine or tenseparate developmental stages have been recognized.In order to simplify our discussion, however, we willconsider the events in four principal stages:

1. Multiplication of the rhizobia, colonization of therhizosphere, and attachment to epidermal and roothair cells.

2. Characteristic curling of the root hairs and invasionof the bacteria to form an infection thread.

3. Nodule initiation and development in the root cor-tex. This stage is concurrent with stage 1.

4. Release of the bacteria from the infection thread andtheir differentiation as specialized nitrogen-fixingcells.

The four principal stages are illustrated in Figure 11.3.In this section, we will concentrate on the physiologyand morphology of infection and nodule development.Genetic aspects will be addressed in a subsequentsection.

11.3.1.1 Early stage involves colonization andnodule initiation. Rhizobia are free-living, sapro-phytic soil bacteria. Their numbers in the soil are highlyvariable, from as few as zero or 10 to as many as 107

gram−1 of soil, depending on the structure of the soil,water content, and a variety of other factors. In thepresence of host roots, the bacteria are encouragedto multiply and colonize the rhizosphere. The initialattraction of rhizobia to host roots appears to involvepositive chemotaxis, or movement toward a chemicalstimulant. Chemotaxis is an important adaptive featurein microorganisms generally. It allows the organism

1� nodulemeristem

Rhizobia

Root hair

1� nodulemeristem

Infectionthread

Developingnodule

Root epidermis

Rootcortex

A. B. C.

FIGURE 11.3 Schematic diagram of the infection process leading to nodule forma-tion. (A) Rhizobia colonize the soil in the vicinity of the root hair in response tosignals sent out from the host root. The rhizobia in turn stimulate the root hair tocurl while, at the same time, sending mitogenic signals that stimulate cell divisionin the root cortex. (B) Rhizobia invade the root by digesting the root hair cell walland forming an infection thread. The rhizobia continue to multiply as the infectionthread elongates toward the root cortex. (C) The infection thread branches to pen-etrate numerous cortical cells as a visibly evident nodule develops on the root. Thefinal stage (not shown) is the release of rhizobia into the host cells and the activationof the nitrogen-fixing machinery.

11.3 Legumes Exhibit Symbiotic Nitrogen Fixation 199

to detect nutrients and other chemicals that are eitherbeneficial or required for their growth and reproduc-tion. A group of chemicals that have been implicated inattraction of rhizobia are the flavonoids (Figure 11.4).

Once rhizobia have colonized the rhizosphere, theybegin to synthesize morphogenic signal molecules callednodulation factors, or nod factors. Nod factors arederivatives of chitin, a β-(1 → 4)-linked polymer ofN-acetyl-D-glucosamine found in the cell walls of fungiand exoskeletons of insects. Nod factors are similarpolymers except that a fatty acid replaces the acetylgroup at one end of the molecule. Nod factors are con-sequently considered lipo-chitooligosaccharides. Nodfactors secreted into the soil solution by the rhizobiainduce several significant changes in the growth andmetabolism of the host roots as a prelude to rhizobialinvasion of the root hair and subsequent nodule develop-ment. These changes (Figure 11.3A) include increasedroot hair production and the development of shorter,thicker roots. Stimulated by the nod factors to renewtheir growth, the root hairs develop branching and curlat the tip.

Before actually invading the host, rhizobia alsorelease mitogenic signals that stimulate localized celldivisions in the root cortex. These cell divisions formthe primary nodule meristem, defining the region inwhich the nodule will eventually develop (Figure 11.3A).A second center of cell division arises in the pericycle.Eventually these two masses of dividing cells will fuse toform the complete nodule.

Rhizobia-host specificity is probably determinedwhen the rhizobia attach to the root hairs and mustinvolve some form of recognition between symbiont andhost. As a general principle, recognition between cellsinvolves chemical linkages that form between uniquemolecules on cell surfaces. In the case of rhizobia-hostinteractions, recognition appears to involve two classesof molecules: lectins and complex polysaccharides.Lectins are small, nonenzymatic proteins synthesized

OH

OH

OH

OH

O

O

LuteolinFIGURE 11.4 Structure of a common flavonoid impli-cated in rhizobia-host interactions. Leuteolin (flavone)is released by the host root. The flavonoid interacts withthe product of the bacterial nodD gene, leading to theinduction of other nodulation genes.

by the host and have the particular ability to recognizeand bind to specific complex carbohydrates.

Individual legume species each produce differentlectins with different sugar-binding specificities. Lectinsappear to recognize complex polysaccharides foundon the surface of the potential symbiont. Althoughbacterial surfaces normally contain an array of com-plex extracellular polysaccharides, the synthesis of addi-tional nodulation-specific extracellular polysaccharidesis directed by bacterial genes that are activated in thepresence of flavonoids in the host root exudate. Hostrange specificity would thus result from attachment ofthe rhizobium to the host root hair because of spe-cific lectin-surface polysaccharide interactions. Supportfor this hypothesis comes from experiments in whichthe gene for pea lectin was introduced into roots ofwhite clover. The result was that clover roots could benodulated by strains of Rhizobium leguminosarum, biovarviciae, which are usually specific for peas.

11.3.1.2 Second stage involves invasion of theroot hair and the formation of an infection thread.In the second stage of nodulation, the bacterium mustpenetrate the host cell wall in order to enter the spacebetween the wall and the plasma membrane. In pea, thepreferred attachment site is the tip of the growing roothair. The root hairs of pea grow by tip growth; that is,new wall material is laid down only at the tip of the elon-gating hair cell. Colonies of attached rhizobia becomeentrapped by the tip of the root hair as it curls around.How rhizobia actually breach the cell wall is not known,but the process almost certainly includes some degreeof wall degradation. There is some evidence that rhi-zobia release enzymes such as pectinase, hemicellulase,and cellulase, which degrade cell wall materials. Theseenzymes could result in localized interference with theassembly of the growing wall at the root tip and allowthe bacteria to breach the cell wall and gain access tothe underlying plasma membrane.

Once the rhizobia reach the outer surface of theplasma membrane, tip growth of the root hair ceasesand the cell membrane begins to invaginate. The resultis a tubular intrusion into the cell called an infec-tion thread, which contains the invading rhizobia(Figure 11.3B). The infection thread elongates by addingnew membrane material by fusion with vesicles derivedfrom the Golgi apparatus. As the thread moves throughthe root hair cell, a thin layer of cellulosic material isdeposited on the inner surface of its membrane. Becausethis new wall material is continuous with the originalcell wall, the invading bacteria never actually enter thehost cell but remain technically outside the cell.

The infection thread continues to elongate until itreaches the base of the root hair cell. Here it must againbreach the cell wall in order for the bacteria to gainaccess to the next cell in their path. This is apparently

200 Chapter 11 / Nitrogen Assimilation

accomplished by fusing the infection thread membranewith the plasma membrane. In the process, some bacteriaare released into the apoplastic space. These bacte-ria apparently degrade the walls of the next cell inline, thus allowing the infection process to continueinto successive cells in the cortex. As the infectionthread moves through the root hair into the cortex,the bacteria continue to multiply. When the threadreaches the developing nodule, it branches so that manyindividual cells in the young nodule become infected(Figure 11.3C).

11.3.1.3 Finally bacteria are released. The finalstep in the infection process occurs when the bacteria are‘‘released’’ into the host cells. Actually the membraneof the infection thread buds off to form small vesicles,each containing one or more individual bacteria. Shortlyafter release, the bacteria cease dividing, enlarge, anddifferentiate into specialized nitrogen-fixing cells calledbacteroids. The bacteroids remain surrounded by amembrane, now called the peribacteroid membrane.Differentiation into a bacteroid is marked by a numberof metabolic changes, including the synthesis of theenzymes and other factors that the organism requiresfor the principal task of nitrogen fixation.

The infection process continues throughout the lifeof the nodule. As the nodule increases in size due tothe activity of the nodule meristem, bacteria continueto invade the new cells. Also as the nodule enlarges andmatures, vascular connections are established with themain vascular system of the root (Figure 11.5). These

Medulla

Nodulemeristem

Root vascular tissue

RootCortex

Vascular strand

Senescent region

ActiveN2 -fixingregion

FIGURE 11.5 Schematic diagram of a cross-sectionthrough a mature nodule. Vascular connections withthe host plant provide for the exchange of carbon andnitrogen between the host and the microsymbiont.

vascular connections serve to import photosyntheticcarbon into the nodule and export fixed nitrogen fromthe nodule to the plant.

11.4 THE BIOCHEMISTRYOF NITROGEN FIXATION

Dinitrogen is not easily reduced because the interatomicnitrogen bond (N ≡ N) is very stable. In the industrialprocess, reduction of the dinitrogen triple bond withhydrogen can be achieved only at high temperature andpressure and at the cost of considerable energy. Biolog-ical reduction of dinitrogen is equally costly, consuminga large proportion of the photoassimilate provided bythe host plant.

11.4.1 NITROGEN FIXATION ISCATALYZED BY THE ENZYMEDINITROGENASE

Only prokaryote cells are able to fix dinitrogen princi-pally because only they have the gene coding for thisenzyme (see Chapter 8, Box 8.1). The enzyme dini-trogenase has been purified from virtually all knownnitrogen-fixing prokaryotes. It is a multimeric proteincomplex made up of two proteins of different size(Figure 11.6). The smaller protein is a dimer consistingof two identical subunit polypeptides. The molecularmass of each subunit ranges from 24 to 36 kD, depend-ing on the bacterial species. It is called the Fe proteinbecause the dimer contains a single cluster of four ironatoms bound to four sulphur groups (Fe4S4). The largerprotein in the dinitrogenase complex is called the MoFeprotein. It is a tetramer consisting of two pairs of identi-cal subunits with a total molecular mass of approximately220 kD. Each MoFe protein contains two molybdenumatoms in the form of an iron-molybdenum-sulphur co-factor. The MoFe protein also contains Fe4S4 clusters,although the exact number is uncertain. It varies as afunction of the species or its physiological condition.

fdred

fdox

N2 + 8H+

2NH3 + H2

Feox

Fered

MoFered

MoFeox

DINITROGENASE

MgADP + PiMgATP2-

Fe protein MoFe protein

FIGURE 11.6 Schematic diagram of the dinitrogenasereaction in bacteroids. Electron flow is from left to right.The principal electron donor is ferredoxin (fd), whichreceives its electron from respiratory substrate.

11.4 The Biochemistry of Nitrogen Fixation 201

The overall reaction for reduction of dinitrogen toammonia by dinitrogenase is shown in the followingequation:

8H+ + 8e− + N2 + 16 ATP → 2NH3

+ H2 + 16 ADP + 16Pi (11.1)

Note that the principal product of biological nitro-gen fixation is ammonia, but that for every dinitrogenmolecule reduced, one molecule of hydrogen is gen-erated. We will return to the problem of hydrogenevolution later. Also note that reduction of dinitrogenis a two-step process. In the first step, the Fe pro-tein is reduced by a primary electron donor, usuallyferredoxin. Ferredoxin is a small (14 to 24 kD) proteincontaining an iron-sulphur group. Electrons are carriedby the iron moiety, which can exist in either the reducedferrous (Fe2+) or the oxidized ferric (Fe3+) states. It isof interest to note that ferredoxin not only participatesin nitrogen fixation, but is an important electron carrierin photosynthesis as well (see Chapter 7).

In the second step, the reduced Fe protein passeselectrons to the MoFe protein, which catalyzes thereduction of both dinitrogen gas and hydrogen. Theprecise role of ATP in the reaction is not yet clear, but itis known to react with reduced Fe protein and to causea conformational change in this protein that alters itsredox potential. This facilitates the transfer of electronsbetween the Fe protein and the MoFe protein.

11.4.2 NITROGEN FIXATION ISENERGETICALLY COSTLY

Biological reduction of dinitrogen, as is industrialnitrogen fixation, is very costly in terms of energy. One

measure of energy cost is the number of ATP required.At least 16 ATP are required for each molecule ofdinitrogen reduced—two for each electron transferred(Equation 11.1). By comparison, only 3 ATP arerequired to fix a molecule of carbon dioxide in photo-synthesis (see Chapter 8). The total energy cost ofbiological nitrogen fixation, however, must take intoaccount the requirement for reduced ferredoxin aswell. Were this reducing potential not required forthe reduction of dinitrogen, it could have been madeavailable for the production of additional ATP or otheruses by the plant. It has been estimated that the reducingpotential used in nitrogen fixation is equivalent to atleast a further 9 ATP, bringing the total investment toa minimum of 25 ATP for each molecule of dinitro-gen fixed. A similar calculation for CO2 fixation bringsthe total to 9 ATP, about one-third the cost fornitrogen.

Another and perhaps better way to assess the costof nitrogen fixation is by measuring the amount ofcarbon utilized in the process. The ultimate source ofenergy for symbiotic nitrogen fixation is carbohydrateproduced by photosynthesis in the host plant. A por-tion of that carbohydrate is diverted from the plantto the bacteroid, where it is metabolized to producethe required reducing potential and ATP. It has beencalculated that, in soybean, approximately 12 grams ofcarbon are required to fix a gram of dinitrogen. It isclear that nitrogen fixation represents a considerabledrain on the carbon resources of the host plant. Adiagram summarizing the integration of photosynthe-sis, respiration, and nitrogen fixation is presented inFigure 11.7.

PHOTOSYNTHATE(from leaf)

Glycolysis

NAD+

H+

N2 + 8H+

2NH3+ H2

H+

H+

NAD+

8e−

NADH ATP

ATP

ADP + Pi

ADP + Pi

NADH

fdred

fdox

Aminoacids

EXPORTfrom nodule

Bacteroidmembrane

Dinitro genase

CAC

Respiratory chain

FIGURE 11.7 Summary diagram illustrating the interactions between photosynthesis,respiration, and nitrogen fixation in bacteroids.

202 Chapter 11 / Nitrogen Assimilation

11.4.3 DINITROGENASE IS SENSITIVETO OXYGEN

One of the more critical problems facing nitrogen-fixingorganisms is the sensitivity of dinitrogenase to molecularoxygen. Both the Fe protein and the MoFe protein arerapidly and irreversibly inactivated by molecular oxygen.The half-life, or time to reduce activity by one-half,of isolated Fe protein in air is 30 to 45 seconds; thehalf-life of MoFe protein is 10 minutes. This extremesensitivity of dinitrogenase to oxygen raises a problemfor nitrogen-fixing organisms. The large amounts ofenergy required (in the form of ATP and reductant) areproduced through a cellular respiratory pathway thatcan operate efficiently only when molecular oxygen ispresent (see Chapter 10). How then does the organismreconcile the conflicting demands of the respiratorypathway for oxygen and the sensitivity of dinitrogenaseto oxygen?

Several strategies for regulating oxygen level havedeveloped to resolve this conflict. First, many free-livingbacterial nitrogen fixers have retained an anaerobiclifestyle or, if facultative, fix dinitrogen only underanaerobic conditions. Production of ATP and reductantis markedly less efficient under anaerobic conditions,which may offer a partial explanation for why, in spiteof their numbers, free-living nitrogen fixers contributea relatively small proportion of the total nitrogen fixedbiologically.

Second, certain species of nitrogen-fixing cyanobac-teria have structurally isolated the nitrogen-fixingapparatus (Figure 11.8). The nitrogen-fixing cells of thecyanobacteria are specialized cells called heterocysts.Heterocysts have thickened, multilayered cell wallsthat restrict the diffusion of oxygen. They are alsocharacterized by a high respiratory activity that main-tains a low intracellular oxygen concentration. Thus,

FIGURE 11.8 Light micrograph of the cyanobacteriumAnabaena showing heterocysts. Nitrogen fixation is car-ried out in the enlarged cells or heterocysts, whosestructure and metabolism limits the concentration offree oxygen. (Copyright E. Reschke. Peter Arnold, Inc.Reprinted by permission.)

respiration has a dual role: it provides the necessaryATP needed for nitrogen fixation and it ensuresthat oxygen concentrations remain low in the cyto-plasm where dinitrogenase is localized. Furthermore,although heterocysts are photosynthetic cells, they lackphotosystem II and thus do not evolve oxygen. Theyalso do not contain Rubisco and therefore cannot fixCO2. However, heterocysts have retained photosystem Iand thus have retained the capacity to synthesize ATPthrough cyclic photophosphorylation (see Chapter 7).

Third, the oxygen supply is regulated to a largeextent by an oxygen-binding protein called leghemo-globin in legume nodules. Leghemoglobin is synthe-sized by the host plant and is located within thebacteroid-infected host cell. Leghemoglobin may com-prise as much as 30 percent of the host cell proteinand gives the nodule a distinctive pink color when a cutsurface is exposed to air. Leghemoglobin is similar instructure to the hemoglobin of mammalian blood. Itsfunction is also similar, since it apparently binds oxy-gen and controls the release of oxygen in the region ofthe bacteroid. The equilibrium concentration of oxy-gen in the bacteroid zone is thus kept at a level (about10 nM) sufficient to support bacteroid respiration—andthe production of ATP and reducing potential—whileat the same time preventing excess oxygen from inac-tivating dinitrogenase. Oxygen levels must be carefullybalanced, because too low an oxygen concentration canalso limit dinitrogenase activity in nodules. This couldbe a result of limiting ATP availability.

11.4.4 DINITROGENASE RESULTSIN THE PRODUCTIONOF HYDROGEN GAS

A final problem facing nitrogen-fixing organisms isthe evolution of hydrogen. Although the mechanismis not well understood, it appears that hydrogen pro-duction by dinitrogenase is an inescapable byproductof the nitrogen-fixation reaction. As noted earlier inequation 11.1, at least one molecule of hydrogen (H2)is evolved for every molecule of N2 reduced. This isactually a minimum value. When dinitrogenase is notoperating optimally, as might be the case when the sup-ply of reductant to dinitrogenase is suboptimal, evenmore electrons may be diverted to the production ofhydrogen. As much as 25 to 30 percent of the ATP andelectrons supplied to dinitrogenase may be consumedby hydrogen production. In 1980, it was estimated thatmore than one million tons of hydrogen were releasedto the atmosphere annually from nitrogen-fixing rootnodules.

Needless to say, H2 production along with nitrogenfixation is wasteful, consuming energy that might other-wise be used to reduce dinitrogen. However, although allnitrogen fixers produce hydrogen, not all release hydrogen

11.5 The Genetics of Nitrogen Fixation 203

into the atmosphere. Many nitrogen-fixing organismscontain an oxygen-dependent enzyme, called uptakehydrogenase, which recovers some of the energy lostto hydrogen production. This is accomplished by cou-pling H2 oxidation to ATP production. The electronsare returned to the reductant pool for dinitrogenase.

Understandably, considerable interest has focusedon the biochemistry and physiology of hydrogenaseaction and the expression of its genes (known as hupgenes). Using biotechnology to increase the number ofrhizobia strains with the capacity to recycle hydrogenhas the potential to increase the overall energy efficiencyof biological nitrogen fixation in important agriculturalcrops.

11.5 THE GENETICS OFNITROGEN FIXATION

It should be evident from the discussion up to thispoint that nitrogen fixation involves very complex rela-tionships between organisms and their environment orbetween rhizobia and host in the case of symbioticfixation. The switch to nitrogen-fixing metabolism inanaerobic environments or infection and subsequentnodule development in symbiotic relationships requiresmajor changes in the genetic programs of the organismsinvolved. The genetics of infection, nodulation, and thenitrogen-fixing machinery is currently one of the moreexciting and rapidly advancing areas in the study ofnitrogen fixation.

11.5.1 NIF GENES CODEFOR DINITROGENASE

In free-living nitrogen fixers, the principal requirementis for the synthesis of the enzyme dinitrogenase. Dini-trogenase synthesis is directed by a set of genes knownas nif genes. Best characterized is Klebsiella pneumonieae,where at least 17 nif genes have been described. The nifgenes include structural genes that encode for dinitro-genase protein as well as a number of regulatory genes.Two genes, the nif D and nif K genes, for example,encode the two different subunits of the MoFe protein.The Fe protein and ferredoxin are encoded by nif Hand nif F, respectively. Other nif genes are involved ininsertion of the FeMo cofactor and the activation andprocessing of the enzyme complex.

11.5.2 NOD GENES AND NIF GENESREGULATE NODULATION

At least three different sets of genes, including nif genes,are involved in the symbiotic process. In the early stagesof nodulation, prior to infection of the root, a set ofrhizobial nod genes is switched on by flavonoids in the

host root exudate. The nod genes are located in a largecircular piece of rhizobial DNA (or plasmid) knownas the Sym (for symbiosis) plasmid. Three nod genes(nodA, nodB, nodC) are basic nodulation genes commonto most rhizobia. They code for the chitooligosaccha-ride core of the nod factors. However, recently, genesequencing of two photosynthetic strains of Bradyrhizo-bia, named BTAi1 and ORS278, indicate that neither ofthese strains contain the expected nod genes. Althoughthese unique strains are capable of inducing stem nodulesin their hosts, Aeschynomene sensitiva and Aeschynomeneindica, purine derivatives appear to trigger this noduleformation.

The role of the nodD gene appears to be pivotal.Its expression is differentially affected by root exudatesand its product in turn activates transcription of boththe nodABC group and a series of host-specific genes(nodEFGH). These host-specific genes code for modi-fications to the nod factors that are important in deter-mining host specificity. The pivotal role of nodD hasbeen demonstrated by transferring the nodD gene from astrain of Rhizobium that infects Parasponia to a strain thatnormally infects only clover. The clover strain was thenable to nodulate Parasponia. One of the first responsesof root epidermis cells to the presence of nod-factors isthe rapid efflux of Ca2+ which occurs within seconds ofexposure to nod-factors. Recently, it has been reportedthat this Ca2+ efflux is associated with the activationof trimeric G-proteins. Thus, the Rhizobium nod-factorsignal transduction pathway appears to be mediated byplasma membrane G-proteins coupled to the activationof intracellular Ca2+ second messenger pathways.

During the latter stages of nodule development,rhizobial nif and fix genes are switched on. The dis-crimination between nif and fix genes is not alwaysclear, except that, as in the free-living forms, nif genesare involved in the synthesis and regulation of dinitro-genase. The fix genes are restricted to symbiotic nitro-gen fixers. At least one (fixX) encodes a ferredoxin andothers may be involved in the transport of electrons todinitrogenase.

Development of an active nodule requires a numberof nodule-specific proteins contributed by the host cells.These proteins, called nodulins, are encoded by nodgenes located in the host cell genome. Early nodulins areexpressed during the infection process and nodule devel-opment. Although several have been identified, their roleis not clear. Early nodulins appear to be involved withthe infection thread plasma membrane and in the forma-tion of the nodule primordia. The expression of latenodulin genes coincides more or less with the onset ofnitrogen fixation and appears to be involved in nodulefunction and maintenance. Leghemoglobin is the mostabundant late nodulin. Other late nodulins includeenzymes such as uricase and glutamine synthetaseinvolved in the metabolic processing of fixed nitrogen.

204 Chapter 11 / Nitrogen Assimilation

11.5.3 WHAT IS THE SOURCE OF HEMEFOR LEGHEMOGLOBIN?

Early experiments with a mutant of Rhizobium melilotiiindicated that the heme was supplied by the bacteroid.The mutant, unable to synthesize the heme precursorδ-aminolevulinic acid (ALA), produced white nodulesthat were unable to fix dinitrogen. These results suggestthe host plant is unable to provide sufficient heme tobuild adequate levels of leghemoglobin. However, inlater experiments with soybean, plants infected withBradyrhizobium japonicum carrying the same mutationproduced fully competent nodules. Thus, the sourceof the heme component of leghemoglobin is not yetclear.

Symbiotic nitrogen fixation clearly requires thecoordinated expression of many genes of both the hostand microsymbiont. Understanding how these genes areregulated and how the complex processes of infectionand nodule development are coordinated constitutes oneof the more challenging problems facing plant physi-ologists today. Armed with sufficient understanding ofthe process and the tools of modern molecular genet-ics, plant scientists may one day be able to extend therange of biological nitrogen fixation to other importantcrop species—thus extending the benefits of nitrogenfixation to nitrogen-poor soils and reducing the eco-nomic and environmental costs of chemical nitrogenfertilizers.

11.6 NH3 PRODUCED BYNITROGEN FIXATION ISCONVERTED TO ORGANICNITROGEN

The first stable product of nitrogen fixation is ammonia(NH3), although at physiological pH ammonia is almostcertainly protonated to form ammonium ion:

NH3 + H+ ↔ NH+4 (11.2)

Plants that cannot fix dinitrogen meet their nutritionalneeds by taking in nitrogen from the soil. While thereare exceptions, most plants are able to assimilate eitherNH+

4 or NO−3 , depending on their relative availability

in the soil. In most soils, ammonia is rapidly convertedto nitrate by the nitrifying bacteria described earlierin this chapter. Nitrifying bacteria do not grow wellunder anaerobic conditions and consequently ammoniawill accumulate in soils that are poorly drained. Nitri-fication itself is also inhibited in strongly acidic soils.Some members of the family Ericaceae, typically foundon acidic soils, have adapted by preferentially utiliz-ing ammonium as their nitrogen source. One extremeexample is the cranberry (Vaccinium macrocarpon), nativeto swamps and bogs of eastern North America, which

cannot exploit NO−3 as a nitrogen source and must take

up nitrogen in the form of ammonium ion.Regardless of the route taken, assimilation of min-

eral (inorganic) nitrogen into organic molecules is acomplex process that can be very energy intensive. It hasbeen estimated, for example, that assimilation of ammo-nium nitrogen consumes from 2 to 5 percent of theplant’s total energy production. Nitrate, on the otherhand, must first be reduced to ammonium before it canbe assimilated, at a cost of nearly 15 percent of totalenergy production. In this section, we will review theassimilation first of ammonium nitrogen and then ofnitrate nitrogen.

11.6.1 AMMONIUM IS ASSIMILATEDBY GS/GOGAT

Although NH+4 is readily available to many plants, either

as the product of nitrogen fixation or by uptake fromthe soil, it is also quite toxic to plants. In nitrogen-fixingsystems, NH+

4 will inhibit the action of dinitrogenase.Ammonium also interferes with the energy metabolismof cells, especially ATP production. Even at low con-centrations, NH+

4 has the potential to uncouple ATPformation from electron transport in both mitochondriaand chloroplasts (see Chapters 7 and 10). Consequently,it appears that plants can ill afford to accumulate excessfree NH+

4 . It is assumed that most plants avoid anytoxicity problem by rapidly incorporating the NH+

4 intoamino acids.

The general pathway for NH+4 assimilation in

nitrogen-fixing symbionts has been worked out largelyby supplying nodules with labeled dinitrogen (13N2or 15N2). These studies have indicated that the initialorganic product is the amino acid glutamine. Assim-ilation of NH+

4 into glutamine by legume nodules isaccomplished by the glutamate synthase cycle, a path-way involving the sequential action of two enzymes:glutamine synthetase (GS) and glutamate synthase(GOGAT)2 (Figure 11.9). Both GS and GOGAT arenodulin proteins that are expressed at high levels in thehost cytoplasm of infected cells, outside the peribac-teroid membrane. The NH+

4 formed in the bacteroidmust therefore diffuse across the peribacteroid mem-brane before it can be assimilated.

In the first reaction of the glutamate synthasecycle, catalyzed by GS, the addition of an NH+

4group to glutamate forms the corresponding amide,glutamine:

glutamate + NH+4 + ATP → glutamine + ADP + Pi

(11.3)

2The acronym GOGAT refers to glutamine-2-oxoglutarate-amino-transferase. 2-Oxoglutarate is an alternative name forα-ketoglutarate.

11.6 NH3 Produced by Nitrogen Fixation Is Converted to Organic Nitrogen 205

Energy to drive the amination of glutamate is providedby ATP, yet an additional cost of nitrogen fixation.Glutamine is then converted back to glutamate bythe transfer of the amide group to a molecule ofα-ketoglutarate.

glutamine + α-ketoglutarate + NADH →2glutamate + NAD+ (11.4)

Reaction 11.4 is catalyzed by GOGAT and requiresreducing potential in the form of NADH. Theα-ketoglutarate is probably derived from photosyn-thetic carbon through respiration in the host cell.α-Ketoglutarate is an intermediate in the respiratorypathway for the oxidation of glucose. Note that reactionequation 11.4 gives rise to two molecules of glutamate.Since only one molecule of glutamate is required tokeep the cycle going, the other is available for exportto the host plant (Figure 11.9). Overall, then, carbonskeletons originating with photosynthesis and nitrogenfixed by the microsymbiont are combined to formorganic nitrogen that is exported out of the nodule foruse by the host.

There is a possible alternative pathway for nitrogenassimilation, involving the direct reductive aminationof α-ketoglutarate by the enzyme glutamate dehy-drogenase (GDH). However, although GDH activityhas been detected in nodules, there is no convincingevidence that it plays a significant role in NH+

4 assimi-lation under normal circumstances. Both the quantitiesand activities of GS and GOGAT are much higher thanGDH. GS alone may account for as much as 2 percent ofthe total soluble protein outside the bacteroid. In addi-tion, GDH has a much lower affinity (see Chapter 8,Box 8.1) for NH+

4 than does GS and could hardly beexpected to compete with GS for available NH+

4 . Thecost of the glutamate synthase cycle is one ATP foreach NH+

4 assimilated, but the benefit is rapid assim-ilation. The high affinity of GS for NH+

4 , togetherwith the high concentration of the enzyme, ensures

that the free NH+4 concentration is kept below toxic

levels.Before leaving the glutamate synthase cycle, it

is important to note that GS and GOGAT are notrestricted to nodules. These enzymes are located in theroots and leaves of non-nitrogen-fixing plants wherethey also catalyze the assimilation of NH+

4 nitrogen.

11.6.2 PII PROTEINS REGULATEGS/GOGAT

As illustrated in Figure 11.9, GS/GOGAT representsa critical metabolic point of coordination betweennitrogen assimilation (NH+

4 ) and carbon metabolism(α-ketoglutarate). Cellular carbon/nitrogen statusis sensed by the signal sensing protein, PII. PIIproteins are one of the most ubiquitous and highlyconserved regulatory proteins found in nature. Theyare present in the ancient archeabacteria, eubacteria aswell as eukaryotes such as plants. This relatively smallchloroplastic protein of about 14 kDa is nuclear encodedand senses both the ATP status and α-ketoglutaratelevels by allosteric means (see Chapter 8, Box 8.1).Cellular nitrogen status is sensed through changes inglutamine availability which results in post-translationalmodification of PII by uridylylation (Figure 11.10).Like the light-regulated enzymes of the Calvin Cycle(see Chapter 8), GS/GOGAT can exist in either anactive or an inactive state. When glutamine levelsin the GS/GOGAT cycle are low, PII proteins areuridylylated using uridine triphosphate (UTP) to formPII-UMP (Figure 11.10). This modified PII proteininteracts with inactive GS/GOGAT to convert thelatter from the inactive to the active form whichstimulates glutamine synthesis and hence the stimulatesassimilation of NH+

4 . Conversely, when glutaminelevels are high, PII-UMP is converted back to PIIwhich converts the active GS/GOGAT to the inactiveform which slows the rate of NH+

4 assimilation.

NH4 + Glutamate Glutamine

ATP ADP + Pi

GS

GOGAT

Export

+

Glutamate + NAD+ �-Ketoglutarate + NADH + H+

FIGURE 11.9 Assimilation of ammonium bythe glutamate synthase cycle. The enzymeglutamate synthetase (GS) converts ammo-nium plus glutamate to glutamine. Glu-tamine undergoes a transamination reac-tion with α-ketoglutarate, which results inthe production of two molecules of gluta-mate via the enzyme glutamate synthase(GOGAT). One glutamate is exportedwhile the other is recycled through thereaction catalyzed by GS.

206 Chapter 11 / Nitrogen Assimilation

PII

PII-UMP

High GlutamineLow Glutamine

GS/GOGAT(inactive)

GS/GOGAT(active)

GS/GOGAT(inactive)

GS/GOGAT(active)

UMPUTP

PPi

FIGURE 11.10 Regulation of GS/GOGAT by Uridyly-lation of PII Proteins. Uridine triphosphate (UTP) isa phosphorylated nucleotide analogous to ATP (seeChapter 5) except that the nitrogenous base, uracil,replaces adenine. UTP, uridine triphopshate; UMP, uri-dine monophosphate; PPi, pyrophosphate.

11.6.3 FIXED NITROGEN IS EXPORTEDAS ASPARAGINE AND UREIDES

The final step in nitrogen fixation is the export of thefixed nitrogen from the nodule to other regions of thehost plant. Export of the organic nitrogen productsfrom nodules is primarily through the xylem. Conse-quently, the form in which the nitrogen is exportedhas been identified primarily by analysis of xylem sap.There are some pitfalls to such analyses, however, as

Allantoin

NH

H2N CO

CCO

HN

C

NH

O

H

Urea

H2N C NH2

O

Urate

NH

HN CC

C

HN

O

H2N C

O

HN

H

NH

C C

O

NH2

COO_

Allantoate

NH2

COO+

Citrulline

C

O

CH2

CH2

CH2

NH

CH NH3+C O

O CNH

NH

FIGURE 11.11 Structures of the principal ureides used in the transport of assimilatednitrogen in some nitrogen-fixing species. Ureides are considered derivatives of ureaand are formed principally from uric acid (urate). The N—C—N urea backbone isshown in bold print.

there is no guarantee that all of the nitrogen presentin sap represents current nodule production. Some ofthe better analyses have been conducted directly ondetached nodules or by monitoring the flow of organicnitrogen following fixation of 15N2. These studies haveshown that although glutamine is the principal organicproduct of nitrogen fixation, it rarely accounts for asignificant fraction of the nitrogen exported, at least inlegumes. In some groups of legumes, largely those oftemperate origins, such as pea and clover, the aminoacid asparagine is the predominant form translocated.Legumes of tropical origins, for example, soybean andcowpea, appear to export predominantly derivatives ofurea, known as ureides (Figure 11.11).

The biosynthetic pathway for asparagine in nodulesinvolves two transamination reactions. A transami-nation reaction is the transfer of an amino groupfrom an amino acid to the carboxyl group of a ketoacid. Transamination reactions, catalyzed by a class ofenzymes known as aminotranferases, enable nitrogeninitially fixed in glutamate to be incorporated into otheramino acids and, ultimately, into protein. Aminotrans-ferases are found throughout the plant—in the cytosol,in chloroplasts, and in microbodies—wherever pro-tein synthesis activity is high. The enzymes involvedin asparagine biosynthesis in nodules appear to be sim-ilar to those found elsewhere in the plant. The firststep is the transfer of an amino group from glutamateto oxaloacetate, catalyzed by the enzyme aspartateaminotransferase.

glutamate + oxaloacetate→ α-ketoglutarate + aspartate (11.5)

11.7 Plants Generally Take Up Nitrogen in the Form of Nitrate 207

The glutamate used in this reaction is derived fromthe GS-GOGAT reactions in the nodule. In order tocontinue the synthesis and export of asparagine, thenodule requires a continued supply of the 4-carbon acidoxaloacetate. This could be provided through the oxi-dation of carbon in the nodule; oxaloacetate is anotherintermediate in the respiratory oxidation of glucose.However, nodules from a number of species exhibithigh activities of the enzyme phosphoenolpyruvate car-boxylase (PEP carboxylase). PEP carboxylase catalyzesthe addition of a carbon dioxide (a carboxylation reac-tion) to the 3-carbon phosphoenolpyruvate (PEP) toform oxaloacetate (OAA) (Equation 11.6). PEP car-boxylase is involved in a number of important metabolicpathways in plants and animals, including respirationand photosynthesis.

PEP + CO2 → OAA (11.6)

In the second step of asparagine biosynthesis,the amide nitrogen is transferred from glutamine toaspartate.

glutamine + aspartate + ATP→ glutamate + asparagine + ADP + Pi (11.7)

The enzyme for this reaction is asparagine synthetaseand the reaction is driven by the energy of one moleculeof ATP for each asparagine synthesized.

The synthesis of ureides is more complex, bothbiochemically and with respect to the division of laborbetween the microsymbiont and tissues of the hostplant. Allantoin and allantoic acid (Figure 11.11)are formed by the oxidation of purine nucleotides,which apparently requires an active symbiosis. Urei-des apparently serve specifically for the transport ofnitrogen. They are translocated through the xylemto other regions of the plant, where they are rapidlymetabolized. In the process, NH+

4 is released, which isthen reassimilated via GS and GOGAT in the targettissue.

Although the synthesis of asparagine, and especiallythe ureides, both appear to be complex processes, thereare some advantages relating to the energy costs andefficiencies of nitrogen export. It has been estimatedthat the carbon metabolism associated with nitrogenexport may consume as much as 20 percent of the photo-synthate diverted to nitrogen fixation. One way to judgeefficiency is to consider the amount of carbon requiredfor each nitrogen exported. The ureides, with a carbonto nitrogen ratio of 1 (C:N 1), are the most economicin the use of carbon. Both asparagine and citrulline(C:N 2) require more carbon in their transport andglutamine (C:N 2.5) would be the least economic.The energy costs of ureides, asparagine, and citrulline,in terms of ATP consumed, are about the same, so theprincipal advantage to be gained by the ureide-formersappears to be a favorable carbon economy.

11.7 PLANTS GENERALLY TAKEUP NITROGEN IN THEFORM OF NITRATE

Except in extreme situations noted earlier, nitrate(NO−

3 ) is the more abundant form of nitrogen insoils and is most available to plants that do notform nitrogen-fixing associations. However, in spiteof numerous studies describing the physiology ofNO−

3 uptake, there is a great amount of uncertaintysurrounding the mechanism of NO−

3 transport intoroots. It has been shown in various studies that uptakeof NO−

3 is sensitive to (1) low temperature, (2) inhibi-tors of both respiration and protein synthesis, and(3) anaerobic conditions. All of these results supportthe hypothesis that NO−

3 transport across the root cellmembrane is an energy-dependent process mediated bya carrier protein (see Chapter 3).

In root cells that have never been exposed to nitrate,there appears to be a limited capacity for NO−

3 uptake.This suggests a small amount of carrier is present in themembrane at all times (that is, a constitutive protein).On exposure to external nitrate, the rate of uptakeincreases from two- to fivefold, but addition of inhibitorsof protein synthesis causes the rate to fall rapidly backto the constitutive level. This pronounced sensitivity ofNO−

3 uptake to inhibitors of protein synthesis suggeststhat the bulk of the carrier protein is inducible, that is,the presence of NO−

3 in the soil stimulates the synthesisof new carrier protein. Once inside the root, NO−

3 maybe stored in the vacuole, assimilated in the root cells, ortranslocated in the xylem to the leaves for assimilation.

Nitrate cannot be assimilated directly but must firstbe reduced to NH+

4 in order to be assimilated intoorganic compounds. This is a two-step process, the firstbeing the reduction of NO−

3 to nitrite (NO−2 ) by the

enzyme nitrate reductase (NR).

2H+ + NO−3 + 2e− → NO−

2 + H2O (11.8)

NR is generally assumed to be a cytosolic enzyme.The product NO−

2 then moves into plastids (in roots)or chloroplasts (in leaves) where it is quickly reducedto NH+

4 by the enzyme nitrite reductase (NiR). Inleaves, the electrons required for the reduction of NO−

2to NH+

4 are generated

8H+ + NO−2 + 6e− → NH+

4 + 2H2O (11.9)

by photosynthetic electron transport. Thus, the assimi-lation of NO−

3 competes with the assimilation of CO2(Chapter 8) for photosynthetic electrons. As a con-sequence, NO−

3 assimilation in leaves can also beconsidered a photosynthetic process similar to CO2assimilation. The interactions between carbon andnitrogen metabolism indicate the importance ofphotosynthetic electron transport in overall primaryreductive metabolism.

208 Chapter 11 / Nitrogen Assimilation

Nitrite is toxic and is rarely found at high concen-trations in plants. This is no doubt because the activityof NiR (per gram dry weight of tissue) is normally sev-eral times higher than the activity of NR. The resultingammonia is then rapidly assimilated into organic com-pounds via the GS/GOGAT system already described.In non-nitrogen-fixing systems, both GS and GOGATare commonly found in root and leaf cells. GS is foundin the cytosol of root cells and in both the cytosol andchloroplasts of leaf cells. GOGAT is a plastid enzyme,localized in the chloroplasts of leaves and in plastidsin roots. Depending on its location, GOGAT may useferredoxin, NADH, or NADPH as electron donors.

Nitrate reductase is a ubiquitous enzyme found inboth prokaryote and eukaryote cells. In prokaryotes,the principal electron donor is ferredoxin, while inhigher plants electrons are donated by the reducedforms of one of the pyrimidine nucleotides, nicotinamideadenine dinucleotide (NAD) or nicotinamide adeninedinucleotide phosphate (NADP) (Chapter 7). The en-zyme isolated from a variety of higher plants is composedof two identical subunits with a molecular mass of approx-imately 115 kD. A key constituent of NR is molybdenum;NR is the principal Mo-protein in non-nitrogen-fixingplants. One of the results of Mo deficiency is markedlyreduced levels of nitrate reductase activity and conse-quent nitrogen starvation (see Chapter 4).

NR is a highly regulated, inducible enzyme. It haslong been recognized that both substrate (NO−

3 ) andlight are required for maximum activity and that induc-tion involves an increase in the level of NR messengerRNA followed by de novo synthesis of NR protein.Treatment of cereal seedlings such as barley (Hordeumvulgare) or maize (Zea mays) with nitrate in the darkinduces relatively low levels of NR activity, but activ-ity is strongly promoted if seedlings are also exposedto light. Induction by light is eliminated by varioustreatments that interfere with chloroplast developmentor photosynthetic energy transformations, implying arequirement for photosynthetic energy. NR activity canalso be reversibly regulated by red and far-red light, indi-cating control by the phytochrome system (Chapter 22).

More recent work has established that NR activity isalso subject to post-translational regulation by a specificNR protein kinase. Protein kinases, first characterizedby Edwin Krebs and Edmund Fischer in the 1950s, area ubiquitous class of enzymes that phosphorylate otherproteins by transferring a phosphate group from adeno-sine triphosphate (ATP). The phosphate group can thenbe removed by a second enzyme called protein phos-phatase. It is increasingly evident that, by switchingenzymes and other proteins on and off, reversible pro-tein phosphorylation plays a central role in regulatingmetabolism. The fundamental role of protein kinaseswas recognized by the award of the Nobel Prize toKrebs and Fischer in 1992.

In the case of NR, the enzyme appears to be active inboth the phosphorylated and nonphosphorylated states.When NR is phosphorylated and the leaf is transferredfrom the light to dark, however, the enzyme is rapidlyinactivated by binding with a small inhibitor protein. NRactivity is slowly restored on return to light by a release ofthe inhibitor protein and subsequent phosphatase action.The question of why there exists such a complex systemfor regulation of NR activity has yet to be answered.The overall effect, however, is to coordinate nitratereduction with photosynthetic activity. It ensures thatnitrate reduction is engaged only after photosynthesis isfully active and able to provide both the energy requiredand the carbon skeletons necessary for incorporation ofammonia.

As indicated earlier, nitrate assimilation can be car-ried out in either the root or shoot tissues in most plants.Several studies have shown that the proportion of NO−

3reduced in the root or shoot depends to a large extenton the external NO−

3 concentration. At low concentra-tions, most of the NO−

3 can be reduced within the roottissues and translocated to the shoot as amino acids oramides. At higher concentrations of NO−

3 , assimilationin the roots becomes limiting and a higher proportionof the NO−

3 finds its way into the translocation stream.Thus, at higher concentrations, a higher proportion ofthe nitrogen is assimilated in the leaves.

Not all plants have the same capacity to metabolizeNO−

3 in their roots. In the extreme, NO−3 is virtually

the sole nitrogen source in the xylem sap of cocklebur(Xanthium strumarium). This is because cocklebur hasno detectable NR in its roots. On the other hand,plants such as barley (Hordeum vulgare) and sunflower(Helianthus annus) translocate roughly equal proportionsof NO−

3 and amino acid/amide nitrogen, and radish(Raphanus sativus) translocates only about 15 percent ofits nitrogen as NO−

3 .

11.8 NITROGEN CYCLING:SIMULTANEOUS IMPORTAND EXPORT

Nitrogen uptake by most plants is highest during its earlyrapid-growth phase and declines as reproductive growthbegins and the plant ages. Cereals, for example, take upas much as 90 percent of their total nitrogen requirementbefore the onset of reproductive growth. Most of thisnitrogen is directed toward young, expanding leaves,which reach their maximum nitrogen content just priorto full expansion. The leaf then begins to export nitrogen.Several studies have shown that mature leaves continueto import nitrogen, even though they have become netnitrogen exporters and the total nitrogen content of theleaf is in decline. This simultaneous import and exportof nitrogen is known as nitrogen cycling.

11.9 Agricultural and Ecosystem Productivity Is Dependent on Nitrogen Supply 209

The export of nitrogen from leaves becomes par-ticularly significant as the seed begins to develop. Thenitrogen requirement of developing seeds is sufficientlygreat that it cannot be met by uptake from the soil (inthe case of cereals, for example) or by nitrogen fixedin nodules. The additional nitrogen must come fromvegetative parts, principally leaves (Figure 11.12). Thismay have significant implications for the photosyntheticcapacity of leaves. The major leaf protein is Rubisco,the enzyme that catalyzes photosynthetic incorporationof carbon dioxide (see Chapter 8). Rubisco maycomprise from 40 to 80 percent of the total solubleprotein in leaves of soybean and cereal grains. Perhapsbecause of its abundance, Rubisco also functions as astorage protein; it may be degraded when nitrogenis required elsewhere in the plant, such as developingseeds. With the loss of Rubisco there is a concomitantdecline in photosynthetic carbon fixation. In the caseof soybean and other symbiotic legumes, this meansless energy available to the nodule to support nitrogenfixation. This competition between nitrogen and carbonsupply may be a major factor limiting seed developmentin legumes. In the case of cereals or plants growing in

20 40 60 80 100 120

0.6

1.2

1.8

2.4

3.0

3.6

Tota

l red

uced

nit

roge

n (m

mol

)

Days after sowing

Leaves

Ear

FIGURE 11.12 Nitrogen redistribution in wheat. (FromAbrol et al., in H. Lambers, J. J. Neeteson, I. Stuhlen(eds.), Fundamental, Ecological and Agricultural Aspectsof Nitrogen Metabolism in Higher Plants, Dordrecht:Martinus Nijhoff, 1986. Reprinted by permission ofKluwer Academic Publishers).

nitrogen-poor soils, mobilization of accumulated nitro-gen from the leaves represents the principal source ofnitrogen for developing fruits and seeds. In perennialplants, nitrogen from senescing leaves is mobilized andtranslocated to the roots for storage over the winter. Inthis way, the nitrogen is conserved and made available tosupport the first flush of renewed growth the followingspring.

11.9 AGRICULTURALAND ECOSYSTEMPRODUCTIVITY ISDEPENDENT ONNITROGEN SUPPLY

In terms of quantity, nitrogen is the fourth most abun-dant element in plants and is the most abundant mineralelement (see Chapter 4). On a dry-weight basis, herba-ceous plant material typically contains between 1 and 4percent nitrogen, mostly in the form of protein. At thesame time, the availability of nitrogen in the soil maybe limited by a number of environmental factors, suchas temperature, oxygen, water status, and pH, whichinfluence the activity of microorganisms responsiblefor nitrogen fixation, nitrification, and ammonification.Moreover, a substantial quantity of nitrogen is removedeach year with the harvested crop. It is not too sur-prising, then, that crop growth is most often limited bynitrogen supply.

In agricultural situations, the application of nitrogenfertilizers overcomes environmentally imposed nitrogenlimitation. Most crops respond to applied nitrogen withincreases in yield (Figure 11.13). At sufficiently highapplication rates, factors other than nitrogen becomelimiting and there is no further gain. At even higherapplication rates, yield may decline slightly, but this isprobably due to excess salt in the soil rather than someform of nitrogen toxicity. Data such as those shownin Figure 11.13 are of considerable practical value tofarmers, who want to maximize the ratio of yield to inputcosts. Throughout North America, corn is the leadingconsumer of nitrogen fertilizer and farmers typicallyapply 100 to 150 kg N ha−1 each growing season. Duringits early rapid growth phase, a well-irrigated stand ofcorn will take up as much as 4 kg of nitrogen ha−1 day−1.The use of such large amounts of nitrogen fertilizers isalso costly in terms of energy. It has been estimated, forexample, that fully one-third of the energy cost of a corncrop is accounted for by the production and distributionof nitrogen fertilizers.

Without continued application of fertilizers, yieldsof nonleguminous crops traditionally decline over aperiod of years. In a few situations where records havebeen kept, yields on plots from which nitrogen fer-tilizers have been withheld will eventually stabilize at

210 Chapter 11 / Nitrogen Assimilation

300

Yiel

d (b

ushe

ls/h

a)

200

100

50

040 80 120 160 200 220

Nitrogen input (kg/ha)

FIGURE 11.13 The effect of applied nitrogen fertilizer oncorn yield.

a lower level that can be sustained indefinitely. Sus-tained low yields are possible because the extraction ofnitrogen (and other nutrients) from the soil is balancedagainst replenishment from all sources, including rain-fall, irrigation water, dust, and weathering of parentrock.

The role of nitrogen in natural ecosystems is muchmore difficult to define, in part because the level of inputsis very low relative to the total nitrogen pool and much ofthe nitrogen is recycled. Still, it is generally agreed thatnitrogen is limiting in most natural ecosystems just asit is in agriculture. In forest ecosystems, approximatelytwo-thirds of the annual nitrogen input is contributedby nitrogen fixation, while the other third is believed tobe derived primarily from atmospheric sources: eitherthrough rainfall or dry deposition of nitrogen oxides. Arecent study has shown that, on average, close to half ofthe incoming fixed nitrogen is retained in the canopy.This should not be surprising since the photosyntheticapparatus present in the leaves of the canopy containsthe bulk of the assimilated organic nitrogen in formof proteins such as Rubisco (see Chapter 8), the mostabundant protein in nature, the major light harvestingpolypeptides present in thylakoid membranes, and ofcourse, the pigments, chlorophyll a and chlorophyll b,which are also rich in nitrogen (see Chapter 4). Thisimplies that foliar absorption of nitrogen could playa significant role in nitrogen uptake by forest species.

This seems particularly true of trees growing at highelevations, which are frequently bathed in cloud cover,or trees that grow near urban industrialized areas.

Except in mature, slowly growing forests, most ofthe nitrogen is taken up and either retained in thecanopy or held in long-term storage in the litter onthe forest floor where it is slowly recycled. The nitro-gen content of the litter is slowly leached into the soilby rain and surface water or is broken down into simplercompounds by a variety of soil bacteria, fungi, earth-worms, and other decomposing organisms. The finalstep in the breakdown is mineralization, or the for-mation of inorganic nitrogen from organic nitrogen.Mineralization is largely due to the process of ammoni-fication described earlier. Mineralization is invariablyaccompanied by immobilization, or the retention anduse of nitrogen by the decomposing organisms. Avail-ability of litter nitrogen to plants depends above all onnet mineralization, or the extent to which mineraliza-tion exceeds immobilization.

The balance between mineralization and immobi-lization and oxidation of the mineralized NH+

4 -nitrogenby nitrifying bacteria is regulated by environmentalparameters. Principal among these are temperature, pH,soil moisture, and oxygen supply. The optimum tem-perature for nitrification generally falls between 25◦Cand 35◦C, although climatic adaptations of indigenousnitrifying bacteria to more extreme temperatures havebeen demonstrated. In one study, for example, the opti-mum temperature for nitrification in soils of northernAustralia was 35◦C, in Iowa (U.S.) it was 30◦C, andin Alberta (Canada), 20◦C. As noted earlier, soil pHis a major limiting factor in the growth of nitrifyingbacteria. The growth of Nitrobacter is probably inhib-ited by ammonium toxicity at high pH (7.5 and above),while aluminum toxicity is suspected as the cause of lim-ited nitrification in acid soils. Soil moisture and oxygensupply go hand-in-hand—little nitrification occurs inwater-saturated soils because of the limited O2 supply.At the other extreme, the rate of nitrification declineswith decreasing soil water potential (�/soil) below about−0.03 to −0.04 MPa.

The relative significance of nitrification in the nitro-gen cycle of natural ecosystems is not altogether clear.Most studies indicate a relatively minor role, since littleif any surplus NO−

3 is found in the soil or streams ofmost undisturbed ecosystems. Experimental deforesta-tion, however, leads to a rapid rise (as much as 50-fold)in the levels of NO−

3 in stream water. NO−3 levels grad-

ually returned to normal only as the vegetation beganto regrow. These results suggest that nitrification is asignificant source of nitrogen, but rapid uptake of NO−

3by plants is an important factor in maintaining low levelsof NO−

3 in the soil solution.Trees and other plants also tend to conserve a

large proportion of their nitrogen, withdrawing nitrogen

Further Reading 211

from the leaves and flowers before they are shed andplacing it in storage in the roots and stem tissues.Between one-third and two-thirds of a plant’s nitrogenmay be conserved by such internal cycling. In the case ofdeciduous trees, for example, this stored nitrogen offersa degree of nutritional independence from the oftennitrogen-poor soil during the flush of growth in earlyspring.

SUMMARY

Nitrogen is often a limiting nutrient for plants,even though molecular nitrogen is readily availablein the atmosphere. Plants do not have the genecoding for dinitrogenase but must depend instead onthe nitrogen-fixing activities of certain prokaryoteorganisms to produce nitrogen in a combined form.

Nitrogen-fixing organisms may be free-living orform symbiotic associations with plants. Symbioticnitrogen fixation involves complex genetic and bio-chemical interactions between host plant roots andbacteria. The invading rhizobia induce the formationof root nodules, where the protein leghemoglobinhelps to ensure a low-oxygen environment in whichthe enzyme dinitrogenase can function. The host plantprovides energy in the form of photosynthate and, inturn, receives a supply of combined nitrogen for itsown growth and development.

The product of nitrogen fixation is ammonium,which is rapidly incorporated into amino acids throughGS/GOGAT before it is exported from the nodule.PII proteins sense cellular carbon/nitrogen balanceand regulate GS/GOGAT activity. Plants that do notform nitrogen-fixing associations generally take upnitrogen in the form of nitrate. Nitrate must first bereduced to ammonium before it can be incorporatedinto organic molecules. In leaves, this reducing power isgenerated by photosynthetic electron transport. Conse-quently, the reduction of nitrate to NH+

4 in leaves canbe considered photosynthetic since it competes withthe reduction of CO2 for photosynthetically generatedelectrons.

CHAPTER REVIEW

1. What are ammonification, nitrification, anddenitrification? What are their respec-tive contributions to the nitrogen cycle?

2. What is meant by the statement that biologi-cal nitrogen fixation is exclusively a prokaryotedomain?

3. Describe the process of rhizobial infectionand nodule development in a legume root.

4. Review the biochemistry of nitrogen fixation.How does a bacteroid differ from a bacterium?

5. What is the function of leghemoglobin in symbi-otic nitrogen fixation?

6. The product of nitrogen fixation is ammonia.Trace the path of nitrogen as the ammonia is con-verted to organic nitrogen and translocated to aleaf cell.

7. What is the role of PII proteins in the assimilationof NH+

4 ?8. While most plants take up nitrogen in the

form of nitrate ion, there are some that seemto prefer ammonium. Can you suggest apossible biochemical basis for this difference?

9. Why can nitrate conversion to NH+4 in

leaves be considered photosynthetic?10. Heavy fertilization of agricultural crops with

nitrogen is a costly process, both economicallyand energetically. Is it feasible to produce cropswithout nitrogen fertilizers? If so, what wouldbe the consequences with respect to yields?

FURTHER READING

Buchanan, B. B., W. Gruissem, R. L. Jones. 2000. Biochem-istry and Molecular Biology of Plants. Rockville MD: Amer-ican Society of Plant Physiologists.

Foyer, C. H., G. Noctor. 2002. Photosynthetic NitrogenAssimilation and Associated Carbon and RespiratoryMetabolism. Advances in Photosynthesis and Respiration,Vol. 12. Dordrecht: Kluwer Academic Publishers.

Giraud, E., et al. 2007. Legumes symbioses: Absence ofnod genes in photosynthetic Bradyrhizobia. Science316:1307–1312.

Guerts, R., T. Bisseling. 2002. Rhizobium Nod factorperception and signalling. The Plant Cell (Supplement)14:S239–S249.

Lam, H.-M., K. T. Coschigano, I. C. Oliveira,R. Melo-Oliveira, G. M. Coruzzi. 1996. The moleculargenetics of nitrogen assimilation into amino acids inhigher plants. Annual Review of Plant Physiology and PlantMolecular Biology 47:569–593.

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Respiration

Photosynthesis

Nitrogenassimilation

12Carbon and Nitrogen Assimilation

and Plant Productivity

In an earlier chapter, the theme was introduced thatphotosynthesis is the fundamental basis of competitivesuccess in green plants (Chapters 7 and 8). The sig-nificance of carbon assimilation, however, extends wellbeyond the performance of green plants. Carbon assimi-lation creates plant biomass, or dry matter, which in turnsupports humans and virtually all other heterotrophicorganisms in the biosphere. Since the beginnings of theindustrial revolution, growth of the human populationand industries has been putting increasing pressures onthe biosphere. These pressures have in turn stimulatedinterest in studies of plant productivity, which is directlyrelated to carbon gain through photosynthesis. Produc-tivity and its relationship to yield is of obvious concernto agriculture, but it also has relevance in broaderecological terms because it provides the energeticand material basis for other organisms. Knowledge ofplant productivity provides a basis level for ecosystemresearch, helping to elucidate problems of energy andnutrient flow and their relationships to the structuresof communities. Plant productivity also carries implica-tions for the upper limit of the earth’s sustainable humanpopulation. Knowing how plants function in a naturalenvironment, their potential for harvest, and how theymight respond to potentially stressful environmental

change is essential to learning how to manage worldresources in a time of burgeoning world population.

The physiology of photosynthesis—light capture,energy conversion, and partitioning of carbon—are atthe root of productivity. This chapter will address

• the concepts of carbon gain and productivity,• the interactions between respiration, photosynthesis

and nitrogen assimilation and how these relation-ships determine the overall carbon gain of a plant,

• environmental factors such as light, available car-bon dioxide, temperature, availability of soil waterand nutrients, which influence photosynthesis andproductivity, and various aspects of leaf and canopystructure and their impact on productivity.

The chapter will finish with a brief discussion ofprimary productivity on a global scale.

12.1 PRODUCTIVITY REFERS TOAN INCREASE IN BIOMASS

Although inorganic nutrients are a part of this drymatter, by far the bulk of dry matter for any organism

213

214 Chapter 12 / Carbon and Nitrogen Assimilation and Plant Productivity

consists of carbon. The basic input into the biosphereis the conversion of solar energy into organic matterby photosynthetic plants and microorganisms, known asprimary productivity (PP). Total carbon assimilation isknown as gross primary productivity (GPP). Not allof the GPP is available for increased biomass, however,since there is a respiratory cost that must be takeninto account. The principal focus of most productivitystudies is therefore net primary productivity (NPP).NPP is determined by correcting GPP for energy andcarbon loss due to respiration. NPP is a measure ofthe net increase in carbon, or carbon gain, and reflectsthe additional biomass that is available for harvest byanimals.

12.2 CARBON ECONOMY ISDEPENDENT ON THEBALANCE BETWEENPHOTOSYNTHESISAND RESPIRATION

Because photosynthesis occupies such a prominent posi-tion in the metabolism of higher plants, its rate is oftenregarded as the primary factor regulating biomass pro-duction and crop productivity. Yet it has often beenobserved that plants with similar photosynthetic ratesmay differ markedly with respect to growth rate andbiomass accumulation. Clearly, other factors such as par-titioning and allocation of carbon, translocation rates,and respiration rates must be considered when attempt-ing to understand the overall carbon budget or carboneconomy of a plant. Carbon economy is the term used todescribe the balance between carbon acquisition and itsutilization. Respiration is the principal counterbalanceto photosynthesis. Respiration consumes assimilatedcarbon in order to obtain the energy required to increaseand maintain biomass. Respiratory loss of carbon con-stitutes one of the most significant intrinsic limitationson plant productivity.

In an effort to better understand the impact ofrespiration on the carbon economy of plants, somephysiologists have sought to distinguish experimen-tally between the carbon and energy costs of growth onthe one hand and maintenance on the other. The termgrowth respiration has been coined to account for thecarbon cost of growth. Growth respiration includes thecarbon actually incorporated plus the carbon respired toproduce the energy (in the form of reducing potentialand ATP) required for biosynthesis and growth. Main-tenance respiration, on the other hand, provides theenergy for processes that do not result in a net increasein dry matter, such as turnover of organic molecules,maintenance of membrane structure, turgor, and soluteexchange. These distinctions are not easily made, but

Growthcomponent

Maintenancecomponent

Relative growth rate

Rel

ativ

e re

spir

atio

n ra

te

FIGURE 12.1 Growth and maintenance respiration. Theproportion of respiration devoted to maintenance can beestimated by extrapolation to zero growth rate.

may be estimated by relating respiration rate to the rela-tive growth rate (Figure 12.1). When respiration rate isextrapolated back to zero growth, it can be assumed thatthe residual respiration represents the carbon and energyrequirements for maintenance of the nongrowing cells.

From Figure 12.1 it can be seen that maintenancerespiration relative to total respiration will vary withthe growth rate. The proportion devoted to maintenancewill be least in a young, rapidly growing plant or organwhile it will account for the bulk of respiration ina nongrowing organ such as a mature leaf. Indeed,measuring maintenance respiration by one commonlyused experimental approach assumes that the respirationof a mature leaf is essentially 100 percent maintenance,although a small (but unknown) amount must be usedfor the translocation of solutes into and out of theleaf. Maintenance respiration also tends to be higher inroots than in shoots and other above-ground organs.This may be related to the expenditure of maintenanceenergy by roots for ion uptake to satisfy not only theirown needs but the needs of the shoot as well. It mayalso reflect the observation that the cyanide-resistantalternative pathway tends to be higher in roots.

Respiration produces the metabolic energy thatis required for various growth processes that increasebiomass and agricultural yield, but it can also consumecarbon with little or no apparent yield of useful energy.Since the latter situation represents a loss of carbon tothe plant, it has been assumed that lower respirationrates would establish a more positive carbon economythat might, in turn, result in more rapid growth andincreased productivity. Is it possible to manipulate respi-ration rates in favor of higher productivity? In a series ofstudies, genotypes of perennial rye grass (Lolium perenne)were selected for respiratory rates ranging from ‘‘slow’’(2.0 mg CO2 g−1 h−1) to ‘‘rapid’’ (3.5 mg CO2 g−1 h−1).

12.3 Productivity is Influenced by a Variety of Environmental Factors 215

Growth rate (g m−2 day−1)

Dar

k re

spir

atio

n ra

te(n

l O2 m

in−1

mg−

1 d

ryw

eigh

t)

20

22

24

26

28

30

32

8 10 12 14 16 18

FIGURE 12.2 The inverse correlation between respi-ratory rate and growth rate in genotypes of perennialrye grass (Lolium perenne). (From Wilson, D. Annals ofBotany (London) 49:303–312, 1982. With permission ofthe Annals of Botany Company).

Selection was based on the specific respiration rate ofmature leaves at 25◦C (i.e., maintenance respiration).The results (Figure 12.2) establish a negative correla-tion between respiration and growth rate; that is, thehighest growth rates were recorded for genotypes hav-ing the lowest rates of respiration. It was concluded thatthe higher growth rates resulted from a more efficientuse of carbon as a consequence of reduced respira-tory evolution of CO2 from fully grown tissues. In otherwords, with less of the carbon consumed in maintenancerespiration, a higher proportion of the carbon was avail-able for allocation to growth. Other investigators havefound evidence of similar negative correlations betweenrespiration and growth with wheat, barley, and oats.

Another method for improving respiratory effici-ency might be to reduce the contribution, if any,of cyanide-resistant alternative pathway activity(Chapter 10). The alternative pathway oxidizes carbonwith most of the energy released as heat. The overallimpact of the alternative pathway remains to be firmlyestablished, but there are some promising indications.Certain cultivars have been identified that differ withrespect to the engagement of alternative pathwayrespiration at two different CO2 levels. The cultivarswere crossed to produce progeny that either stronglyexpressed the alternative pathway or showed no expres-sion. The accumulation of biomass was greater in theprogeny from which the alternative pathway was absent.

Studies such as these suggest that respiration has asignificant impact on biomass accumulation and yield.They also indicate that improving yield by manipulatingrespiration might be feasible. Given that approximately

half of the carbon assimilated in photosynthesis is even-tually lost by respiration, reducing either the level ofmaintenance respiration or the engagement of the alter-native pathway should shift the carbon balance in favorof growth respiration and increased biomass production.It is assumed, of course, that maintenance respirationand/or the alternative pathway can be reduced withoutdetrimental effects on the plant, but this remains to beclearly established. Certainly some level of maintenancerespiration is essential to the health of the plant—thislevel is yet to be determined.

One approach to improving the efficiency of netprimary production is through breeding and selectionprograms. It is quite possible that this has already beenachieved to some extent in existing breeding programs,without consciously evaluating the role of respiration.Another approach would be to manipulate respirationthrough genetic engineering. However, respiration isa complex process, both biochemically and physio-logically. It is central to the metabolism of the cell,and involves many different enzymes and ultimatelythe coordinated activities of cellular organelles plusthe cytosol. It is difficult to know which enzymesor reactions—that is, which genes—might be prof-itably manipulated or how an altered respiratory balancewill affect other physiological processes. Although theprospects for improving productivity by manipulatingrespiration are encouraging, there is clearly a great dealof fundamental research yet to be conducted beforesignificant progress can be expected.

12.3 PRODUCTIVITY ISINFLUENCED BY A VARIETYOF ENVIRONMENTALFACTORS

The rate of photosynthesis may be limited by a hostof variables. Which of these variables have an influ-ence and the extent to which the influence is feltdepend on whether one is concerned with a singleleaf, a whole plant, or a population of plants that forma canopy. Included in these variables are both envi-ronmental factors and genetic factors. Environmentalfactors include light, availability of CO2, temperature,soil water, nutrient supply, pathological conditions, andpollutants. Major factors include leaf age, and morphol-ogy, leaf area index, leaf angle, and leaf orientation.The influence of environmental stress, acclimation andadaptation on plant productivity will be discussed inChapters 13, 14 and 15.

12.3.1 FLUENCE RATE

Typical responses of photosynthesis to fluence rate areillustrated in Figure 12.3. At very low fluence rates the

216 Chapter 12 / Carbon and Nitrogen Assimilation and Plant ProductivityR

ate

of C

O2 u

ptak

e (+

) or

evo

luti

on (

–)

Fluence rate (μmol m-2 s-1)

Light compensation point

0 500 1000 1500

0

(+)

(–)

FIGURE 12.3 A graph showing the typical light responseof photosynthesis in C3 plants.

rate of CO2 evolution due to dark respiration exceedsthe rate of photosynthetic CO2 uptake. This results ina negative CO2 uptake, or net CO2 evolution. As flu-ence rate increases, photosynthesis also increases and sodoes CO2 uptake until the rate of CO2 exchange equalszero. This is the fluence rate, known as the light com-pensation point, at which the competing processes ofphotosynthesis and respiration are balanced. The lightcompensation point for most plants falls somewhere inthe range of 10 to 40 μmol m−2 s−1, roughly equiv-alent to the light level found in a well-lighted office,laboratory, or classroom.

At fluence rates above the compensation point, therate of photosynthesis continues to increase until, atleast in C3 plants, it reaches light saturation. In most C3plants at normal atmospheric CO2 levels, photosynthesissaturates with light levels of about 500 to 1000 μmolphotons m−2 s−1, that is, about one-quarter to one-halfof full sunlight. Light saturation occurs because someother factor, usually CO2 levels, becomes limiting. Inmost cases, both the saturation rate of photosynthesisand the fluence rate at which saturation occurs can beincreased by increasing the CO2 level above ambient.

A small number of C3 plants, such as peanut (Arachishypogea), do not light saturate. It is not clear why this isthe case, but these are exceptions to the rule. Individualleaves and plants will also acclimate to the light envi-ronment in which they are grown. The light-saturatedrate of photosynthesis, for example, is lower in leavesthat have acclimated to growth at low irradiance (shadeleaves) than in those that have acclimated to higher irra-diance (sun leaves). Acclimation and adaptation to lightenvironments is discussed in more detail in Chapters 14and 15.

In a natural environment, even C3 plants rarelylight saturate and then only for relatively brief periods.Between dawn and dusk, the rate of photosynthesisgradually increases, reaching a maximum near midday,

and then declines. The photosynthetic rate generallyparallels changes in the irradiance that accompaniesthe rising and setting of the sun. Even during midday,measurable decreases in photosynthetic rate have beenobserved with passing cloud cover, suggesting that eventhen photosynthesis was barely, if at all, light saturated.In another study, annual productivity of several speciesgrowing in a European hedgerow was limited to less thanhalf their potential maximum. The failure of carbon gainto match leaf photosynthetic capacity was attributed toreduced average irradiance due to effects of dawn anddusk, short photoperiods in the spring and fall, and cloudcover. Long-term carbon gain is clearly dependent oncumulative irradiance over the growing season.

12.3.2 AVAILABLE CO2

The carbon dioxide concentration of the atmosphereis relatively low, at least over the short term, at about0.035 percent by volume or 350 μl l−1. This is belowthe CO2 saturation level for most C3 plants at normalfluence rates (Figure 12.4), which means that availabilityof CO2 is often a limiting factor in photosynthesis. InC3 plants, increased photosynthetic rates with higherCO2 levels results from two factors: increased substratefor the carboxylation reaction and, through competitionwith oxygen, reduced photorespiration. Note the inter-action between ambient CO2 levels and light. At higherfluence rates, both the maximum rate of photosynthe-sis and the CO2 saturation level increase. Furthermore,under high light, C3 plants typically exhibit a lower CO2compensation point. This represents the CO2 concen-tration where the rate of CO2 uptake by photosynthesisequals the rate of CO2 evolution due to respiration.

Assessing the impact of CO2 levels on photosyn-thesis is not quite as straightforward as it might at first

Rat

e of

CO

2 u

ptak

e

CO2 concentration (μl l-1)

C3 species (high light)

0 200

C3 species (low light)

100 300 400 500 600 700

FIGURE 12.4 A graph showing the typical response of C3species to ambient CO2 concentration. Arrow indicateCO2 compensation concentration.

12.3 Productivity is Influenced by a Variety of Environmental Factors 217

appear. The rate of photosynthesis is actually deter-mined not by the ambient CO2 concentration, as muchas by the intracellular CO2 concentration, that is,the supply of CO2 at the carboxylation site in thechloroplast. It is assumed that the intracellular CO2concentration is in equilibrium with the intercellularspaces. Since diffusion rates depend in part on con-centration gradients, the primary effect of increasingambient CO2 levels would be to increase the intercellu-lar CO2 concentration by increasing the rate of diffusioninto the leaf. Here it is assumed that water supply is ade-quate and, consequently, stomatal CO2 conductance isnot limiting.

Although it was once thought that stomatalCO2 conductance was the principal factor limitingphotosynthesis, more recent studies suggest it may bethe other way around—stomatal conductance varies inresponse to photosynthetic capacity. Photosyntheticcapacity is determined by the balance betweencarboxylation capacity and electron transport capacity(Figure 12.5). At low CO2 concentrations, the rate ofphotosynthesis is limited by available CO2 and, hence,the carboxylation capacity of the system, but is saturatedwith respect to availability of the acceptor molecule,ribulose-1,5-bisphosphate (RuBP) (see Chapter 8).However, any excess generation of RuBP, which is in

Rat

e of

CO

2 a

ssim

ilati

on

CO2 concentration (μl l-1)

Transition zone

100 200 300 400 500

Acceptor regenerationlimited

Carboxylationlimited

Ci Co

FIGURE 12.5 A model to describe limitation of pho-tosynthetic rate as a function of CO2 concentration.At low CO2 concentrations, photosynthesis is limitedby the carboxylation capacity of the enzyme Rubisco.At high concentrations of CO2, the rate is limited bythe rate of regeneration of the acceptor molecule,ribulose-1,5-bisphosphate. Stomata probably operateto keep the intercellular CO2 concentration within thetransition zone where there is neither an excess of car-boxylating capacity nor an excess of electron transport.Ci and C0 indicate the intercellular and ambient CO2

concentrations, respectively. (Redrawn from Farquhar,G., T. Sharkey. 1982. Annual Review of Plant Physiology33:317–345).

turn dependent on the electron transport reactions, overthat required to support carboxylation would representan inefficient use of resources. Conversely, at high CO2concentrations or in low light, the limiting factor wouldbe the energy-limited capacity to regenerate the accep-tor molecule, ribulose-1,5-bisphosphate. In this case, anexcess of carboxylating capacity—that is, an excess ofRubisco—would be an inefficient use of resources.

The most efficient use of resources for the plantwould be to maintain intercellular CO2 levels in thetransition zone, where there is neither an excess of elec-tron transport capacity nor an excess of carboxylatingcapacity. Because intercellular CO2 levels are at leastpartly determined by stomatal conductance, it appearsthat the principal function of the stomata might be toregulate CO2 uptake in order to keep intercellular CO2levels as much as possible within the transition range.Note that this is not the traditional view of stomatalfunction, which says that stomata operate principally toregulate water loss. Note also that CO2 enrichment athigh fluence rates leads to both higher photosyntheticmaxima and higher CO2 saturation levels (Figure 12.4).These observations suggest that plants are also ableto compensate for higher light levels by increasingtheir carboxylating capacity. Such an increase could beachieved by regulating the amount of catalytic activitiesof photosynthetic enzymes, principally Rubisco.

CO2 limitation is a particular problem in green-houses, especially in winter when the greenhouses areclosed and CO2 levels are reduced due to photosynthe-sis. Even under more normal conditions, most plantswill grow significantly faster and increase yields whenthe atmosphere is enriched with CO2. For these rea-sons, CO2 enrichment has become common practice forcommercial growers of vegetable crops such as lettuce,tomato, and cucumbers. In practice, the CO2 contentof the greenhouse atmosphere is increased up to twicepresent atmospheric levels. Much beyond 700 μl l−1

there is an increasing risk of stomatal closure as well asthe potential to induce feedback-limited photosynthesis(Chapter 9) that will cause a reduction in the rate ofphotosynthesis. Thus, there is an upper limit to whichCO2 concentrations can be increased with an increase inplant productivity. This limitation may occur when thephotosynthetic capacity of source leaves far exceeds sinkcapacity of the plant (see Chapter 9). CO2 enrichmentmight be expected to improve growth and productivityof field crops, but there are obvious technical and eco-nomic problems related to controlling the supply of gasin an open environment as well as potential physiologi-cal problems related to possible feedback inhibition ofphotosynthesis. The rate of photosynthesis is reducedwhen the rate of CO2 assimilation exceeds the capacityfor carbon utilization and export. The rate of photo-synthesis is down-regulated through complex metabolicfeedback loops.

218 Chapter 12 / Carbon and Nitrogen Assimilation and Plant Productivity

12.3.3 TEMPERATURE

Photosynthesis, like most other biological processes,is sensitive to temperature. The temperature responsefor most biological processes reflects the temperaturedependence of the enzymic and other chemical reactionsinvolved. The temperature response curve can be char-acterized by three cardinal points: the minimum andmaximum temperatures (Tmin and Tmax, respectively)at which the reaction can proceed and the optimumtemperature (Topt) (Figure 12.6). Thus there is a rangeof temperatures below the optimum over which the rateof the reaction or process is stimulated with increasingtemperature, and a range beyond the optimum overwhich the rate declines. These points are largely deter-mined by biochemical factors such as the binding ofsubstrate with active sites (Chapter 8, Box 8.1) andprotein (enzyme) stability.

The temperature response of chemical and biologi-cal reactions can generally be characterized by compar-ing the rate of the reaction at two temperatures 10◦Capart, a value known as the Q10:

Q10 = RT + 10/RT (12.1)

The value of Q10 for enzyme-catalyzed reactions is usu-ally about 2, meaning that the rate of the reaction willapproximately double for each 10◦C rise in temperature.This value for Q10 applies primarily to stimulation ofthe reaction by temperatures between Tmin and Topt.Once the optimum is reached, the reaction rate maydecline sharply due to enzyme inactivation (Figure 12.6).Because photosynthesis, respiration, and nitrogen assim-ilation are complex, multienzyme processes, it is gen-erally assumed that temperature responses will tend toreflect the average temperature characteristics for all ofthe enzymes. As we will see in Chapter 14, plants canadjust their Q10 for respiration during acclimation totemperature.

Rea

ctio

n ra

te

Temperature

Tmin

Topt

Tmax

FIGURE 12.6 Temperature dependence and cardinalpoints for a typical biological reaction.

A basic characteristic of photochemical reactionsis that they occur largely independent of temperaturein the biologically relevant range of 0◦–50◦C. Thus,the Q10 for photochemical reactions is close to 1.0.Consequently, the short-term temperature responseof photosynthesis largely reflects the effect of tem-perature on the reactions of carbon metabolism andintersystem electron transport but not the photochem-ical reactions within the reaction centers of photosys-tem II and photosystem I. However, prolonged exposureto high temperature can cause the destabilization ofphotosystem II, which may lead to a decreased capa-city for photosynthetic photochemistry and hence aninhibition in the rates of photosynthetic electron trans-port. Photosystem I is generally much more stable tochanges in environmental conditions than is photo-system II.

Measurement of photosynthetic activity in leavesand whole plants is normally based on net gas exchange;that is, it is based on apparent rates of photosynthesis(AP)—the difference between the actual rate of photo-synthetic CO2 uptake (gross photosynthesis, GP), CO2evolution due to respiration (R), and photorespiration(PR) (Figure 12.7; Equation 12.2).

AP = GP − (R + PR) (12.2)

Because gross photosynthesis (GP), respiration (R),and photorespiration (PR) respond very differently totemperature, the optimum temperature for net photo-synthesis (AP) is not the same as for gross photosynthesis(Figure 12.8). Note that the rate of respiration contin-ues to increase with temperature, reaching a maximumnear 50◦C, where it drops off sharply due to inactivationof enzymes. The temperature response of the rate ofphotorespiration is thought to follow a curve similar tothat shown for respiration.

PR

R

GP

CO2CO2

CO2

Light

FIGURE 12.7 Diagram illustrating processes giving rise toCO2 exchange in the light in a C3 leaf.

12.3 Productivity is Influenced by a Variety of Environmental Factors 219

�Temperature ( C)

Rel

ativ

e ph

otos

ynth

esis

or

resp

irat

ion

-10 0 10 20 30 40 50 60

Topt

Grossphotosynthesis

Netphotosynthesis

Tmin

Topt

Tmax

Respiration

FIGURE 12.8 Diagram illustrating the temperaturedependence of gross photosynthesis, respiration, andnet photosynthesis. Gross photosynthesis increases withthermal activation of the participating enzymes untilinhibitory factors (enzyme inactivation, stomatal clo-sure) take effect. Respiration increases more slowlywith temperature and has a higher temperature opti-mum, but declines more rapidly at high temperature.Net photosynthesis (dashed curve) is determined as thedifference between gross photosynthesis and respiration.The resulting cardinal points for net photosynthesis areindicated.

12.3.4 SOIL WATER POTENTIAL

The importance of available water in determining pro-ductivity cannot be underestimated. The rate of photo-synthesis declines under conditions of water stress, andin cases of severe water stress may cease completely.Stomatal closure and the resultant decrease in CO2 sup-ply due to water stress imposes a major limitation onphotosynthesis. When this occurs in the presence oflight for prolonged periods of time, this lack of CO2supply may lead to photoinhibition of photosynthesis(Chapters 13 and 14). Photorespiration may protect thephotosynthetic apparatus from excess light under suchconditions because the energy absorbed can be used tofix O2 when the CO2 supply is limiting due to stomatalclosure.

Low water potentials reduce turgor pressure inleaf cells, which in turn reduces leaf expansion (seeChapter 17). Under prolonged water stress this resultsin a reduced photosynthetic surface area. There maybe some compensation as stored reserves are mobilizedto offset the loss of new assimilate, but overall evenmild water stress causes a reduction in net productivity.C4 plants enjoy some advantage over C3 plants withrespect to photosynthesis and water stress because oftheir higher water use efficiency (Chapter 15).

12.3.5 NITROGEN SUPPLY LIMITSPRODUCTIVITY

The maximum possible photosynthetic rate of a leaf,known as the leaf photosynthetic capacity, is determinedas the rate of photosynthesis per unit leaf area underconditions of saturating incident light, normal CO2 andO2 concentrations, optimum temperature, and high rel-ative humidity. Although leaf photosynthetic capacitymay vary as much as a hundredfold, it is generally high-est in plants acclimated to resource-rich environments;that is, where light, water, and nutrients are abundant.Reduced photosynthesis is a consequence of deficienciesof virtually all essential elements, but photosyntheticcapacity is particularly sensitive to nitrogen supply. Asa basic constituent of chlorophyll, redox carriers in thephotosynthetic electron transport chain, and all of theenzymes involved in carbon metabolism, nitrogen playsa critical role in primary productivity.

In a C3 species, Rubisco alone will account for morethan half of the total leaf nitrogen. In one study, netphotosynthesis increased linearly with nitrogen content(Figure 12.9). In barley seedlings, a 5-fold increase innitrate supply stimulated a 25-fold increase in net photo-synthesis. One impact of nitrogen deficiency is to reducethe amount and activity of photosynthetic enzymes, butleaf expansion and other factors no doubt contributeto reductions in photosynthetic capacity as well. Thus,to maximize plant biomass requires the integration ofcomplex metabolic pathways including photosynthesis,respiration, and nitrogen assimilation (Figure 12.10).This is due to the fact that photosynthesis providesenergy in the form of fixed carbon as well as basic car-bon skeletons for all the necessary cellular constituents.The energy stored as fixed carbon through photosynthe-sis (Chapters 7 and 8) is retrieved by the cell for all other

Net

pho

tosy

nthe

sis

(mg

CO

2 d

m-2

hr-

1)

30

20

10

00 1 2

Organic N (% of dry matter)

3 4 5

FIGURE 12.9 The relationship between leaf organic nitro-gen content and net photosynthesis for the C3 species,Tall fescue. (Adapted from Plant Physiology 66:97–100.1980. Copyright American Society of Plant Physiolo-gists).

220 Chapter 12 / Carbon and Nitrogen Assimilation and Plant Productivity

Respiration

Photosynthesis

Nitrogen assimilation

FIGURE 12.10 Diagram illustrating the interdependenceof photosynthesis, respiration, and nitrogen assimilation.

processes such as growth and development throughthe process of respiration (Chapter 10). Intermediatesof the TCA cycle combine with nitrate assimilation toprovide amino acid biosynthesis (Chapters 10 and 11).Photosynthetic energy is used directly in nitrate reduc-tion (Chapter 11) and nitrogen assimilation providesamino acids for the synthesis of enzymes and proteinsinvolved in both photosynthesis and respiration. Thus,all three primary metabolic pathways are intimatelyconnected to each other through GS/GOGAT and itsregulation by PII proteins (Chapter 11).

12.3.6 LEAF FACTORS

The net carbon gain of an individual leaf depends onits photosynthetic capacity limited by environmentalparameters and balanced against its construction andmaintenance costs. For example, the net carbon gain of aleaf varies markedly during leaf development and aging.During initial development and the rapid growth phase,the photosynthetic capacity of a leaf also increases. How-ever, the developing leaf functions as a sink, utilizingcarbon assimilated locally as well as importing carbon tosupport its expansion. Leaf photosynthetic capacity thendeclines as the aging leaf undergoes senescence, a pro-gressive deterioration of the leaf characterized in partby the loss of chlorophyll and photosynthetic enzymes.Only in the period between full expansion and the onsetof senescence does a leaf produce a profit in terms ofcarbon gain.

Different types of leaves may also have differ-ent photosynthetic capacities. Evergreen leaves, forexample, have a lower photosynthetic capacity thandeciduous leaves and, because they take longer todevelop, their construction costs are also higher. Still,the evergreen leaf may be favored if the cost of main-taining the leaf over winter together with the cost of alower photosynthetic capacity are less than the cost ofproducing a new leaf.

Regardless of leaf photosynthetic capacity, unfavor-able environmental conditions will cause a reductionin long-term carbon gain (Chapter13). In any naturalenvironment, available water, the quantity of light, andtemperature will all vary widely and to some extent inde-pendently, often keeping the photosynthetic rate well

below full capacity. In addition, a prominent feature inthe environment of any leaf is the presence of otherleaves; that is, leaves are normally part of a canopy. Netprimary productivity of a stand of plants is markedlyinfluenced by canopy structure. Canopy structure is inturn determined by the age, morphology, angle, andspacing of individual leaves.

A herbaceous C3 annual plant is characterized bya gradient in leaf age and development along the stemaxis. The young, growing leaves at the top are exposedto full sunlight while older leaves further down maybe heavily shaded. Irradiance reaching shaded leavesmay be reduced to 10 percent or less, thus producinga very low net photosynthesis. Very often the fluencerate reaching leaves lowermost in a canopy may fallbelow the light compensation point for a large part ofthe day. Those leaves would not only no longer con-tribute to net photosynthesis, but would incur a negativecarbon gain through respiratory loss. Many herbaceousannuals avoid the costs of maintaining such nonpro-ductive leaves by undergoing sequential senescence;that is, the older leaves lower in the canopy senesce asnew leaves are being formed at the top of the canopy.Senescing leaves may lose as much as 50 percent oftheir dry weight, largely in the form of soluble organicnitrogen compounds, before dying and falling to theground. These compounds are exported to developingleaves and other sinks where they are reused. In thisway, limited resources are redistributed among leaves ofvarying ages in order to maximize whole plant carbongain.

Canopy architecture is important when consider-ing agricultural crops and natural ecosystems becauseit determines how efficiently light is absorbed. Highproductivity is in part dependent on the extent to whichground area is covered with photosynthetic surface.Because sunlight striking exposed soil does not con-tribute to productivity, most agricultural systems aredesigned so that the young plants fill in the canopyrapidly in order to maximize interception of availablelight. On the other hand, planting at too high a densitywill introduce mutual shading by leaves in the canopy.Shading reduces the overall efficiency of light intercep-tion and, consequently, reduces long-term carbon gain(Figure 12.11). The ratio of photosynthetic leaf area tocovered ground area is known as the leaf area index(LAI). Leaf area is usually taken as the area of a singlesurface (or projected area, where leaves are not planar).Because both leaf surface and the covered ground aremeasured as areas (m2), LAI is dimensionless. Values ofLAI in productive agricultural ecosystems typically fallin the range of 3 to 5.

The optimum LAI for a given stand of plantsdepends on the angle between the leaf and the stem.Horizontal leaves, typical of beans (Phaseolus) and sim-ilar crops, are efficient light absorbers because of the

Summary 221

Rel

ativ

e bi

omas

s

Leaf area index (LAI)

FIGURE 12.11 The relationship between biomass and leafarea index (LAI) in a crop. LAI is varied by varying thedensity of plants in the stand. Higher planting densitieslead to a decline in biomass because of mutual shadingof leaves and loss of carbon by respiration in the shaded,nonproductive leaves.

broad surface presented to the sun, but they also moreeffectively shade leaves lower down in the canopy. Erectleaves, typical of grasses like wheat (Triticum) and maize(Zea mays), produce less shading but, because of theirsteeper angle, are not as efficient at intercepting light.Experiments with closely spaced rice plants have shownthat carbon gain dropped by a third when leaves wereweighted in the horizontal position. With maize, tyingupper leaves into a more erect posture increased yieldby as much as 15 percent. In some crop plants, suchas sugarbeet (Beta vulgaris), leaf angle varies from nearvertical at the top of the plant to horizontal at thebase. This arrangement reduces light interception bythe uppermost leaves, but allows light to penetrate moredeeply into the canopy. Overall, the more uniform dis-tribution of light tends to improve the efficiency of lightinterception by the canopy and thus the efficiency ofcarbon gain.

The relationship between LAI, leaf angle, and pho-tosynthetic rate has been tested by computer models.The results show that with low values of LAI leaf anglehas little effect, but above a LAI of 3, more layers ofvertical leaves are required to maximize photosynthesis.Field studies have confirmed that canopies with pre-dominantly horizontal leaves have LAI values of 2 orless, while vertical leaf canopies support LAI values of3 to 7.

In cases where the leaves are more or less fixed inspace, the efficiency of light interception will changewith the angle of the sun. Many desert and agricul-tural species, however, have the capacity to alter theorientation of their leaves, allowing them to track thesun as it moves across the sky through the day. Called

heliotropism, or solar tracking, the leaf blades movein such a way that their surfaces remain perpendicularto the sun’s direct rays. Solar tracking would help tomaximize daily carbon gain in those plants that mustcomplete their life cycle in a brief period before theonset of unfavorable conditions such as drought or hightemperatures.

SUMMARY

Carbon assimilation by plants creates the plantbiomass that supports humans and virtually all otherheterotrophic organisms. The study of carbon gain, orproductivity, at both the organismal and populationlevel is an important component of agricultural andecosystem research. Overall carbon gain dependson net primary productivity—the balance betweencarbon uptake by photosynthesis and carbon loss torespiration. Carbon loss to respiration can be dividedinto the carbon cost of growth, or growth respiration,and the cost of simply maintaining structure andprocesses that do not result in a net increase indry matter. Several studies have shown a negativecorrelation between respiration and growth rate. Theimplication is that respiration has a significant impacton biomass accumulation and yield, and that improvingyield by manipulating respiration might be feasible.

Productivity is also influenced by a variety ofgenetic and environmental factors that influence photo-synthesis. These include light, available carbon dioxide,temperature, soil water, nutrients, and canopy struc-ture. The rate of photosynthesis increases betweenthe light compensation point and saturation. Becauseirradiance changes constantly throughout the day,long-term carbon gain depends on the cumulative irra-diance over the growing season.

Although the carbon dioxide content of the atmo-sphere is relatively constant in the short term, thereis evidence that plants use the stomata to keep theinternal carbon dioxide concentration in balance withelectron transport capacity. On the other hand, withadequate light, plants will respond to carbon dioxideenrichment by increased productivity. However, ulti-mately productivity is also dependent on source-sinkrelationships.

Photosynthesis, respiration, and photorespirationrespond differently to temperature. Thus the optimumtemperature for net photosynthesis is not the same asthe optimum temperature for gross photosynthesis.Plants also require an adequate water and nitrogensupply in order to maximize their leaf photosyntheticcapacity. Productivity in a stand depends on the patternof leaf senescence and the structure of the canopy.The ideal canopy maximizes the efficiency of lightinterception and carbon gain by balancing leaf area, leaf

222 Chapter 12 / Carbon and Nitrogen Assimilation and Plant Productivity

angle, leaf orientation, plant density, and senescence ofolder leaves.

CHAPTER REVIEW

1. Distinguish between growth respiration andmaintenance respiration. Which might best bemanipulated in order to improve productivity?

2. Describe how various environmental factors influ-ence plant productivity.

3. Why do large values for leaf area index lead toan overall decline in productivity of a stand?

4. How do plants alter leaf structure and morphologyto protect themselves from excess light?

FURTHER READING

Buchanan, B. B., W. Gruissem, R. L. Jones. 2000.Biochemistry and Molecular Biology of Plants. Rockville,MD: American Society of Plant Physiologists.

Field, T. S., D. W. Lee, N. M. Holbrook. 2001. Why leavesturn red in autumn: The role of anthocyanins in senesc-ing leaves of red-osier dogwood. Plant Physiology 127:566–574.

Foyer, C. H., G. Noctor. 2002. Photosynthetic NitrogenAssimilation and Associated Carbon and RespiratoryMetabolism. Advances in Photosynthesis and Respiration,Vol. 12. Dordrecht: Kluwer Academic Publishers.

Pathogen

Pathogen

HR

Primary infection

SAS

SASASASA

MSAMSASS

MSA

Secondary infection

13Responses of Plants to Environmental Stress

The previous chapters have focused on the underlyingprocesses by which plant roots acquire water and nutri-ents from the soil (Chapters 1 to 4), leaves harvest lightenergy and convert atmospheric CO2 and soil NO−

3 intostable chemical forms of sucrose, starch, and and aminoacids (Chapters 5, 7, 8, 11) and subsequently unlockthe stored chemical energy through the process of gly-colysis and mitochondrial respiration for growth anddevelopment (Chapter 10). By necessity, to elucidatethe biochemical and molecular mechanisms by whichthese processes occur, plants are studied under normalor ideal environmental conditions for growth and devel-opment. However, plants often encounter unusual orextreme conditions: trees and shrubs in the northerntemperate latitudes experience the extreme low temper-atures of winter; alpine plants experience cold and dryingwinds; and agricultural crops may experience periods ofextended drought as well as high and low temperatures.Extremes in environmental parameters create stress-ful conditions for plants, which may have a significantimpact on their physiology, development, and survival.

The study of plant responses to environmental stresshas long been a central theme for plant environmentalphysiologists and physiological ecologists. How plantsrespond to stress helps to explain their geographic dis-tribution and their performance along environmentalgradients. Because stress invariably leads to reducedproductivity, stress responses are also important to

agricultural scientists. Understanding stress responses isessential in attempts to breed stress-resistant cultivarsthat can withstand drought, and other yield-limitingconditions. Finally, because stressful conditions causeperturbations in the way a plant functions, they providethe plant physiologist with another very useful tool forthe study of basic physiology and biochemistry.

This chapter will examine some of the stresses thatplants encounter in their environment. The principaltopics to be addressed include

• the basic concepts of plant stress, acclimation, andadaptation,

• the light-dependent inhibition of photosynthesisthrough a process called photoinhibition,

• the effects of water deficits on stomatal conductance,• the effects of high- and low-temperature stress on

plant survival,• the challenge of freezing stress on plant survival, and• the responses of plants to biotic stress due to infes-

tations by insects and disease.

13.1 WHAT IS PLANT STRESS?

Because life is an endergonic process, that is �G > 0(Chapter 5), energy is an absolute requirement for the

223

224 Chapter 13 / Responses of Plants to Environmental Stress

maintenance of structural organization over the lifetimeof the organism. The maintenance of such complexorder over time requires a constant through put ofenergy. This means that individual organisms are notclosed systems but are open systems relative to theirsurrounding environment. This results in a constantflow of energy through all biological organisms, whichprovides the dynamic driving force for the performanceof important maintenance processes such as cellularbiosyntheses and transport to maintain its characteristicstructure and organization as well as the capacity toreplicate and grow. Such energy flow ensures that livingbiological organisms are never at equilibrium with theirenvironment, that is, �G is never equal to zero, butremain in a steady-state condition far from equilibrium(see Chapter 5). The maintenance of such a steady-stateresults in a meta-stable condition called homeostasis.As a consequence, all life forms may be consideredtransient energy storage devices with finite but varyinglifetimes (Figure 13.1).

Any change in the surrounding environment maydisrupt homeostasis. Environmental modulation ofhomeostasis may be defined as biological stress. Thus,it follows that plant stress implies some adverse effecton the physiology of a plant induced upon a suddentransition from some optimal environmental conditionwhere homeostasis is maintained to some suboptimalcondition which disrupts this initial homeostatic state(Figure 13.2). Thus, plant stress is a relative term sincethe experimental design to assess the impact of a stressalways involves the measurement of a physiologicalphenomenon in a plant species under a suboptimal,stress condition compared to the measurement of thesame physiological phenomenon in the same plantspecies under optimal conditions. Since the extent of astress can be quantified by assessing the difference inthese measurements under optimal versus suboptimalconditions, the basis of stress physiology is comparative

EA EOUTEIN

A

FIGURE 13.1 Biological life forms as energy-storingdevices. A is any biological life form. EIN (thick arrow)is the energy flowing from the surrounding environmentinto the biological organism, A. EOUT (thin arrow) is theenergy flowing out of the biological organism back intothe environment. The thicknesses of the arrows indi-cate the differences in the relative flux of energy flowingin and out of a living organism. EA is the steady-stateenergy stored or trapped by a living organism. Accordingto the First Law of thermodynamics (Chapter 5), EIN +EA + EOUT = 1. Thus, EA = EIN − EOUT.

Rat

e of

aP

hysi

olog

ical

Pro

cess

Homeostatic

State

Death

Stress

Time

FIGURE 13.2 The effects of environmental stress on planthomeostasis. Under some optimal environmental condi-tion, a plant is in homeostasis as indicated by a constantrate of some important physiological process over time.Upon the imposition of an external stress, the rate ofthis physiological process decreases rapidly. It is fatalfor some plants that are unable to adjust to an imposedstress and can not establish a new homeostatic state. Suchplants are classified as susceptible to the stress.

physiology. However, plant species are highly variablewith respect to their optimum environments and theirsusceptibility to extremes of, for example, irradiance,temperature, and water potential. Is stress a function ofthe environment or the organism? For example, are theextreme environments encountered in deserts or arctictundra stressful for plants that normally thrive there?Are these environments stressful only to some speciesbut not to others?

13.2 PLANTS RESPOND TOSTRESS IN SEVERALDIFFERENT WAYS

Plant stress can be divided into two primary categories.Abiotic stress is a physical (e.g., light, tempera,-ture) or chemical insult that the environment mayimpose on a plant. Biotic stress is a biological insult,(e.g., insects, disease) to which a plant may be exposedduring its lifetime (Figure 13.3). Some plants may beinjured by a stress, which means that they exhibitone or more metabolic dysfunctions. If the stress ismoderate and short term, the injury may be temporaryand the plant may recover when the stress is removed.If the stress is severe enough, it may prevent flowering,seed formation, and induce senescence that leads toplant death. Such plants are considered to be suscep-tible (Figure 13.3). Some plants escape the stress alto-gether, such as ephemeral, or short-lived, desert plants.

13.3 Too Much Light Inhibits Photosynthesis 225

Environmental StressAbiotic Biotic

Stress response

Resistance Susceptibility Avoidance

Death

Acclimation

Growth

Survival

Senescence Survival

FIGURE 13.3 The effect of environmental stress on plantsurvival.

Ephemeral plants germinate, grow, and flower veryquickly following seasonal rains. They thus completetheir life cycle during a period of adequate moistureand form dormant seeds before the onset of the dryseason. In a similar manner, many arctic annuals rapidlycomplete their life cycle during the short arctic summerand survive over winter in the form of seeds. Becauseephemeral plants never really experience the stress ofdrought or low temperature, these plants survive theenvironmental stress by stress avoidance (Figure 13.3).

Avoidance mechanisms reduce the impact of astress, even though the stress is present in the environ-ment. Established plants of alfalfa (Medicago sativa), forexample, survive dry habitats as adult plants by sendingdown deep root systems that penetrate the water table.Alfalfa is thereby ensured an adequate water supplyunder conditions in which more shallow-rooted plantswould experience drought. Other plants develop fleshyleaves that store water, thick cuticles or pubescence (leafhairs) to help reduce evaporation, or other modifica-tions that help to either conserve water or reduce waterloss. Cacti, with their fleshy photosynthetic stems andleaves reduced to simple thorns, are another example ofdrought avoiders. Most drought avoiders would beseverely injured should they ever actually experiencedesiccation.

Many plants have the capacity to tolerate a particu-lar stress and hence are considered to be stress resistant(Figure 13.3). Stress resistance requires that the organ-ism exhibit the capacity to adjust or to acclimate to thestress.

Finally, a controversy over terminology is con-cerned with use of the term strategy. Strategy is oftenused to describe the manner in which a plant responds

successfully to a particular stress. Some physiologistsobject to use of the term for the reason that strat-egy implies a conscious plan; that is, it is teleological.1However, strategy can validly describe a genetically pro-grammed sequence of responses that enable an organismto survive in a particular environment.

13.3 TOO MUCH LIGHTINHIBITS PHOTOSYNTHESIS

In Chapters 7 and 8, we discussed the conversion ofvisible light energy into ATP and NADPH throughphotosynthetic electron transport. In addition to form-ing Triose-P that can be converted to sucrose or starch,the ATP and NADPH are required to regenerate RuBPby the Calvin Cycle (Figure 13.4A). The continuousregeneration of RuBP is an absolute requirement forthe continuous assimilation of CO2 by Rubisco. Thisrequirement which is satisfied by the light-dependentbiosynthesis of ATP and NADPH is what makes CO2assimilation light dependent. However, although lightis required for the photosynthetic assimilation of CO2,too much light can inhibit photosynthesis.

In all plants, the light response curve for pho-tosynthesis exhibits saturation kinetics as illustratedin Figure 13.4B. At low irradiance, the rate of CO2assimilation increases linearly with an increase in irra-diance. This is to be expected since more absorbedlight means higher rates of electron transport which, inturn, means increasing levels of ATP and NADPH forthe regeneration of RuBP (Figure 13.4A). Thus, underlow, light-limiting conditions, the rate at which RuBPis regenerated through the consumption of ATP andNADPH by the Calvin Cycle limits the rate of photo-synthesis measured either as CO2 assimilation or O2evolution. The maximum initial slope of the photo-synthetic light response curve under low, light-limitingconditions provides a measure of photosynthetic effi-ciency measured either as moles of CO2 assimilated perphoton absorbed, or alternatively, moles of O2 evolvedper photon absorbed if photosynthesis is measured asthe rate of O2 evolution (Figure 13.4B).

Upon further increases in irradiance, the rate ofphotosynthesis is no longer a linear function of irra-diance but rather levels off. At these higher lightintensities, the rate of photosynthesis is said to belight saturated (Figure 13.4B, red shaded area). Thismeans that the Calvin Cycle is saturated with ATP andNADPH which, in turn, means that Rubisco is satu-rated with one of its substrates, RuBP. The maximumlight saturated rate is a measure of photosynthetic

1Teleology is the doctrine of final causes, assigning purposeto natural processes. Teleological arguments are consideredinappropriate in natural science.

226 Chapter 13 / Responses of Plants to Environmental Stress

RegenerationPS II

A.

B.

hv hv

PS I

2PGA RuBP

Triose-P

Rubisco

CalvinCycle

CO2

ATPNADPH

Rat

e of

CO

2A

ssim

ilati

on

Rat

e of

O2E

volu

tion

Irradiance (μmol m−2s−1)

LightLimited

Photoinhibited

Excess Light

O 2500

Lightsaturated

Pit cavity

FIGURE 13.4 A schematic illustration of the response of photosynthesis to increasingirradiance. (A) A model illustrating the interaction between photosynthetic linearelectron transport and the Calvin Cycle. (B) A schematic light response curve forphotosynthesis measured as either the rate of CO2 assimilation or the rate of O2

evolution. The area above the light response curve represents excess irradiance thatis not used in photosynthesis.

capacity and will have the units of either moles ofCO2 assimilated or moles of O2 evolved per leaf areaper unit time. Thus, under light-saturated conditions,the rate of regeneration of RuBP no longer limitsthe rate of CO2 assimilation but rather it is the rateat which Rubisco can consume RuBP and CO2 thatlimits the rate of photosynthesis. Consequently, underlight-saturated conditions, the rate of photosynthesisbecomes light-independent, that is, the exposure of aplant to higher irradiance no longer changes the rate ofphotosynthesis. Under light-saturated conditions, plants

become exposed to increasing levels of excess light(Figure 13.4B, yellow shaded area), that is, they becomeexposed to more light than the plant can use for photo-synthesis. If the plant continues to be exposed to higherand higher levels of excess light, the rate of photosyn-thesis begins to decrease (Figure 13.4B). This is calledphotoinhibition of photosynthesis and is defined as thelight-dependent decrease in photosynthetic rate thatmay occur whenever the irradiance is in excess of thatrequired either for the photosynthetic evolution of O2or the photosynthetic assimilation of CO2.

13.3 Too Much Light Inhibits Photosynthesis 227

There is a consensus in the literature that inmost plants, PSII is more sensitive to photoinhibitionthan PSI. The effects of exposure to photoinhibitionby shifting plants from low light to high lightcan be assessed either by monitoring changes inphotosynthetic efficiency and photosynthetic capac-ity (Figure 13.5A) or by monitoring chlorophyllfluorescence (Box 13.1) to assess changes in max-imum PSII photochemical efficiency measured asFv/Fm (Figure 13.5B). Photoinhibition that resultsin a decrease in photosynthetic efficiency as wellas photosynthetic capacity (Figure 13.5A) usually reflects

0 500

Chronic Photoinhibition

Control

Capacity

Effic

ienc

y

1000

80

60

40

20

0 10

Dark Control

10

PhotoinhibitionTime (h)

RecoveryTime (h)

Fv/F

m

55

0.8

0.6

0.4

0.2

A.

B.

FIGURE 13.5 The effects of chronic photoinhibition onphotosynthesis. (A) Schematic light response curves forcontrol plants which have not been pre-exposed to highlight and plants that have been pre-exposed to high lightto induce chronic photoinhibition prior to measuringtheir light response curve. The maximum initial slope isa measure of photosynthetic efficiency. The maximumlight saturated rate is a measure of photosynthetic capac-ity. (B) A schematic representation illustrating the effectof exposure time to high light to induce chronic pho-toinhibition on the maximum photochemical efficiencyof PSII measured as Fv/Fm (see Box 13.1). The brokenline illustrates the slow rate of recovery of Fv/Fm whenthe chronically photoinhibited plants are shifted backto low-light conditions. Control plants were kept in thedark over the course of the experiment.

chronic photoinhibition which is the result of photo-damage to PSII. Specifically, the site of damage is theD1 reaction center polypeptide of PSII which causesa decrease in the efficiency of PSII charge separation(Chapter 7). This decrease in the PSII photochemicalefficiency can be measured as Fv/Fm (Figure 13.5B)(Box 13.1). The decrease in PSII photochemical effi-ciency is usually paralleled by a decrease in photosyn-thetic efficiency of O2 evolution. A characteristic ofchronic photoinhibition is that it is only very slowlyreversible after plants are shifted from excess light tolow light (Figure 13.5B).

13.3.1 THE D1 REPAIR CYCLEOVERCOMES PHOTODAMAGETO PSII

PSII reaction centers exhibit an inherent lifetime. Thiswas first indicated by the fact that the D1 polypeptide ofPSII reaction centers exhibits the fastest turnover rateof any plant protein. The D1 polypeptide is degradedand resynthesized in the time span of approximately30 minutes. Recently, it has been proposed that PSIIactually exhibits the properties consistent with a pho-ton counter, that is, its lifetime is dependent uponthe number of photons absorbed and not the absolutetime. It has been calculated that, under normal growthconditions, each PSII reaction center is irreversibly dam-aged presumably due to photooxidative damage andspontaneously degraded after the absorption of 105 to107 photons. Thus, exposure to excess light simplyshortens the lifetime of PSII because these conditionswould enhance the probability of photooxidative dam-age and shorten the time to absorb the necessary photonsto cause the degradation of D1.

How do plants ensure a constant supply of func-tional PSII reaction centers? Plants and green algaeexhibit a single chloroplastic gene that encodes the D1polypeptide called psbA. These organisms have evolveda D1 repair cycle which repairs photodamage to PSII(Figure 13.8). When the D1 polypeptide is damaged, itis marked for degradation by protein phosphorylation.After phosphorylation of D1, PSII is partially disassem-bled and the D1 polypeptide is degraded by proteolysis.Subsequently, the psbA gene is transcribed and translatedusing the chloroplastic transcriptional and translationalmachinery with the subsequent accumulation of a newD1 polypeptide. This new D1 polypeptide is insertedinto the nonappressed, stromal thylakoids and a new,functional PSII complex is reassembled. Given the lat-eral heterogeneity present in thylakoid membranes, themechanism by which damaged PSII complexes migratelaterally from granal stacks to nonappressed, stromalthylakoids for disassembly and reassembly remains to beelucidated.

228 Chapter 13 / Responses of Plants to Environmental Stress

hνhν

Q−

e−

A+

A Q

BOX 13.1MONITORINGPLANT STRESSBY CHLOROPHYLLFLUORESCENCE

In Chapter 6 we defined fluorescence as the emissionof a photon of light by an excited molecule as itreturns to ground state from its lowest singlet excitedstate. Due to the initial thermal deactivation to thelowest singlet excited state, chlorophyll emits red lightas it fluoresces to ground state. Thus, regardless ofthe wavelength used to excite chlorophyll, it alwaysemits red light. Of the total energy absorbed by aleaf, less than 3 percent of that energy is lost due tochlorophyll fluorescence. Due to the differences in thestructure and composition of PSII and PSI, chlorophyllfluorescence measured at room-temperature emanatesprimarily from PSII (Figure 13.6). Due to thisproperty, room-temperature chlorophyll fluorescenceis exploited as a sensitive, intrinsic probe not onlyof PSII function but also of the overall functionof photosynthesis. As a consequence, chlorophyllfluorescence is undoubtedly the most widely used spec-troscopic technique in photosynthesis and plant stressresearch.

Light

PSIPSII PQ

Fluorescence

Sugars

Heat

NADP+

NADPHATPADP+Pi

KP

KD

KF

CO2

FIGURE 13.6 A model illustrating the possible fates ofabsorbed light energy in the photosynthetic apparatus.The primary fates of photosynthetically absorbed lightenergy are thought to include useful photochemistrydesignated by the rate constant, KP, nonphotochemi-cal dissipation of excess absorbed light energy as heatdesignated by the rate constant, KD, and chlorophyll flu-orescence designated by the rate constant, KF. Since lessthan 3 percent of the absorbed light energy is ever lost aschlorophyll fluorescence, this pathway is not considereda major pathway for the safe dissipation of excess lightenergy. However, chlorophyll fluorescence is a sensitive,intrinsic probe for the function of the photosyntheticapparatus and overall physiological status of the plant.

Fluo

resc

ence

yie

ld (

rela

tive

)

Actinicon

FR onSaturatingpulse

Saturatingpulses

Measuringbeam off

Measuringbeam on

FO′

FV′

FM′

FSFO

FV

FM

FIGURE 13.7 Schematic trace of a measurement ofpulse-amplitude-modulated chlorophyll fluorescence(PAM). This method allows one to analyze Chl fluores-cence quenching by the saturation pulse method. Whena sample is in the dark-adapted state, turning on thelow-intensity modulated measuring beam results in aminimal fluorescence yield called Fo. The maximal fluo-resence yield (Fm) can be estimated by the subsequentapplication of a high-intensity millisecond saturatingpulse of white light. Since variable fluorescence (Fv) isdefined as Fm − Fo, therefore,

Fm − Fo

Fm= Fv

Fm.

When the sample is illuminated with actinic light suf-ficient to induce photosynthesis, the fluorescence yield(FS) undergoes complex transitions until eventually FS

reaches a minimal, steady-state level. This is called theKautsky induction curve. PAM fluorescence exploitsthe fact that maximal fluorescence yield (F ′

m) can beassessed during active photosynthesis induced by theactinic light by repetitively applying saturating pulses oflight as indicated by the vertical spikes emanating fromthe Kautsky induction curve indicated in Figure 13.7.To a first approximation, Fm − F ′

m represents nonpho-tochemical quenching of the fluoresence yield, whereasF ′

m − Fs represents photochemical quenching.

Kautsky and Hirsch were the first to report thevariable nature of the chlorophyll fluorescence signal.Subsequent detailed studies of this complex fluorescencesignal (Figure 13.7) showed it to be rich in informationwith respect to the properties of the PSII reaction cen-ter as well as its association with overall photosyntheticelectron transport and CO2 assimilation. When a leaf oran algal suspension is dark adapted for anywhere from5 minutes to 60 minutes at room temperature dependingon the species used, all electrons are drained from thephotosynthetic electron transport chain by PSI causingall PSII reaction centers to be in the open configuration

13.4 Water Stress is a Persistent Threat to Plant Survival 229

[P680 Pheo QA] (Chapter 7). The minimal fluorescenceyield with all PSII reaction centers in the open con-figuration is called the background or Fo fluorescence(Figure 13.7). If the samples are now exposed to a lightintensity sufficiently strong to initiate photosynthesis, afluorescence signal characterized by complex transientscan be detected (Figure 13.7). This is called the Kaut-sky effect. The fast rise from Fo to an initial maximumfluorescence (Fm) (Figure 13.7) is due to the fact that thelight absorbed closes the PSII reaction centers [P680+Pheo Q−

A ] (Chapter 7) much faster through energy trans-fer and photochemistry than they can be reopened byPSI, intersystem electron transport and ultimately CO2assimilation (Figure 13.7). The subsequent decrease inthe fluorescence yield or fluorescence quenching reflectsthe reopening of PSII reaction centers. This is due tothe induction of the much slower, enzyme-catalyzedreactions involved in ATP and NADPH biosynthesis,which are in turn ultimately consumed by biochemicalreactions involved in CO2 assimilation. Thus, chloro-phyll fluorescence yield is inversely proportional to therate of photosynthesis.

It is incumbent upon all photosynthetic organismsto maintain a balance in energy budget, that is, a balancebetween the energy absorbed through photochemistryversus energy either utilized through metabolism andgrowth and/or dissipated nonphotochemically as heat.There is a consensus that most photosynthetic organ-isms possess two primary mechanisms to maintain anenergy balance. (1) Photochemical quenching, mea-sured as qP (Equation 13.1), reflects the capacity toutilize excess absorbed energy through metabolism andgrowth.

qP = Fm′ − Fs/Fm′ − Fo (13.1)

NPQ = Fm − Fm′/Fm′ (13.2)

This involves the upregulation of the expression andactivity of specific enzymes involved in stromal CO2

assimilation (Rubisco), cytosolic sucrose biosynthesis(SPS), as well as vacuolar fructan biosynthesis incertain plant species such as the cereals (Chapter 9).(2) Nonphotochemical quenching, measured asNPQ (Equation 13.2) reflects the capacity to dissipateexcess absorbed energy as heat (Figure 13.6). Thisinvolves primarily the stimulation of the thylakoid,xanthophyll cycle enzymes involved in the reversiblede-epoxidation of violaxanthin to antheraxanthin andzeaxanthin. Photochemical-quenching capacity (qP) andnonphotochemical-quenching capacity (qN, NPQ) canbe estimated in vivo using pulse-amplitude-modulatedchlorophyll fluorescence (PAM). The extent ofphotoinhibition induced by any environmental stresscan be rapidly assessed by measuring the maximumphotochemical efficiency of PSII (Fv/Fm) beforeand after a photoinhibition treatment. Fv/Fm can becalculated from the following equation (Equation 13.3).As expected, the decrease in Fv/Fm follows a similarpattern as the decrease in photosynthetic efficiency ofO2 evolution in response to time under conditions ofphotoinhibition.

Fv/Fm = (Fm − Fo)/Fm (13.3)

REFERENCES

Adams III, W. W., B. Demmig-Adams. 2004. Chlorophyllfluorescence as a tool to monitor plant response to theenvironment. Chlorophyll a Fluorescence—A Signature ofPhotosynthesis. Advances in Photosynthesis and Respiration,Vol. 19, pp. 583–604. Dordrecht: Springer.

Schreiber, U. 2004. Pulse-amplitude-modulation (PAM)fluorometry and saturation pulse method: An overview.Chlorophyll a Fluorescence—A Signature of Photosynthesis.Advances in Photosynthesis and Respiration, Vol. 19,pp. 279–319. Dordrecht: Springer.

Baker, N. R. 2008. Chlorophyll fluorescence: a probe ofphotosynthesis in vivo. Annual Review of Plant Biology 59:89–113.

13.4 WATER STRESS IS APERSISTENT THREATTO PLANT SURVIVAL

Water stress may arise through either an excess ofwater or a water deficit. An example of excess wateris flooding. Flooding stress is most commonly an oxy-gen stress, due primarily to reduced oxygen supply tothe roots. Reduced oxygen in turn limits respiration,

nutrient uptake, and other critical root functions. Stressdue to water deficit is far more common, so much sothat the correct term water deficit stress is usuallyshortened to simply water stress. We will focus onwater deficit stress in this chapter. Because water stressin natural environments usually arises due to lack ofrainfall, a condition known as drought, this stress isoften referred to as drought stress. In the laboratory,water stress can be simulated by allowing transpirationalloss from leaves (Chapter 2), a condition commonlyreferred to as desiccation stress.

230 Chapter 13 / Responses of Plants to Environmental Stress

DI Degradation

Functional PSII

PSII Disassembly

DI Phosphorylation

DI Insertion PSII Reactioncenter damage

Excess light

DI Synthesis

FIGURE 13.8 The D1 repair cycle. Although light is theultimate source of energy for photosynthesis, too muchlight can be dangerous because it may result in damageto the photosystems, especially the D1 reaction cen-ter polypeptide present in PSII. Plants and algae haveevolved an elaborate mechanism to repair photodam-age called the D1 repair cycle. Exposure to an irradiancethat exceeds the capacity of the plant either to utilizethat energy in photosynthesis or to dissipate it safelyas heat results in damage to the D1 polypeptide. Func-tional PSII reaction centers are converted to damagedPSII reaction centers. When this happens, PSII is dis-assembled and D1 is degraded by thylakoid proteolyticenzymes. Subsequently, new D1 is synthesized de novoby the chloroplast translational machinery and insertedinto the thylakoid membrane to form functional PSIIreaction centers.

13.4.1 WATER STRESS LEADSTO MEMBRANE DAMAGE

Damage resulting from water stress is related to thedetrimental effects of desiccation on protoplasm.Removal of water, for example, leads to an increasein solute concentration as the protoplast volumeshrinks, which may itself have serious structural andmetabolic consequences. The integrity of membranesand proteins is also affected by desiccation, which inturn leads to metabolic dysfunctions. Stresses mayalter the lipid bilayer and cause the displacementof membrane proteins, which, together with soluteleakage, contributes to a loss of membrane selectivity, ageneral disruption of cellular compartmentation, and aloss of activity of membrane-based enzymes.

In addition to membrane damage, numerous stud-ies have shown that cytosolic and organellar proteinsmay undergo substantial loss of activity or even com-plete denaturation when dehydrated. Loss of membraneintegrity and protein stability may both be exacer-bated by high concentrations of cellular electrolytesthat accompany dehydration of protoplasm. The con-sequence of all these events is a general disruption ofmetabolism in the cell upon rehydration.

13.4.2 PHOTOSYNTHESIS ISPARTICULARLY SENSITIVETO WATER STRESS

Photosynthesis can be affected by water stress in twoways. First, closure of the stomata normally cuts offaccess of the chloroplasts to the atmospheric supply ofcarbon dioxide. Second, there are direct effects of lowcellular water potential on the structural integrity ofthe photosynthetic machinery. The role of water stressin stomatal closure will be discussed in the followingsection.

Direct effects of low water potential on photo-synthesis have been studied extensively in chloroplastsisolated from sunflower (Helianthus annuus) leaves sub-jected to desiccation. Sunflower has proven useful forthese studies because stomatal closure has only a minoreffect on photosynthesis. This is because direct effectson the photosynthetic activity of chloroplasts decreasethe demand for CO2 and the CO2 level inside the leafremains relatively high. Both electron transport activityand photophosphorylation are reduced in chloroplastsisolated from sunflower leaves with leaf water potentialsbelow about -1.0 MPa. These effects reflect damage tothe thylakoid membranes and ATP synthase protein(CF0-CF1) complex (Chapters 5 and 7).

The direct effects of water stress on photosynthesisare exacerbated by the additional effects of light. Sincewater stress inhibits CO2 assimilation, this means thatwater stress will expose plants to excess light. Thelight absorbed by the photosynthetic pigments of theleaf continue to absorb light but this absorbed lightenergy can not be processed because photosyntheticelectron transport is inhibited. Thus, a concomitanteffect of exposure of plants to a water deficit is chronicphotoinhibition.

13.4.3 STOMATA RESPOND TOWATER DEFICIT

Plants are often subjected to acute water deficits due to arapid drop in humidity or increase in temperature whena warm, dry air mass moves into their environment.The result can be a dramatic increase in the vaporpressure gradient between the leaf and the surroundingair. Consequently, the rate of transpiration increases(Chapter 2). An increase in the vapor pressure gradientwill also enhance drying of the soil. Because evaporationoccurs at the soil surface, the arrival of a dry air masshas particular consequences for the uptake of water byshallow-rooted plants.

Plants generally respond to water stress by closingtheir stomata in order to match transpirational water lossthrough the leaf surfaces with the rate at which watercan be resupplied by the roots. It has been shown in vir-tually all plants studied thus far, including plants from

13.4 Water Stress is a Persistent Threat to Plant Survival 231

desert, temperate, and tropical habitats, that stomatalopening and closure is responsive to ambient humidity.Unlike the surrounding epidermal cells, the surfaces ofthe guard cells are not protected with a heavy cuti-cle. Consequently, guard cells lose water directly to theatmosphere. If the rate of evaporative water loss fromthe guard cells exceeds the rate of water regain fromunderlying mesophyll cells, the guard cells will becomeflaccid and the stomatal aperture will close. The guardcells may thus respond directly to the vapor pressuregradient between the leaf and the atmosphere. Closureof the stomata by direct evaporation of water from theguard cells is sometimes referred to as hydropassiveclosure. Hydropassive closure requires no metabolicinvolvement on the part of the guard cells; guardcells respond to loss of water as a simple osmometer(Chapter 1).

Stomatal closure is also regulated by hydroactiveprocesses. Hydroactive closure is metabolically depen-dent and involves essentially a reversal of the ion fluxesthat cause opening (Chapter 1). Hydroactive closure istriggered by decreasing water potential in the leaf meso-phyll cells and appears to involve abscisic acid (ABA) andother hormones. Since the discovery of ABA in the late1960s, it has been known to have a prominent role instomatal closure due to water stress. ABA accumulatesin water-stressed (that is, wilted) leaves and externalapplication of ABA is a powerful inhibitor of stomatalopening. Furthermore, two tomato mutants, known asflacca and sitiens, fail to accumulate normal levels ofABA and both will wilt very readily. The precise role ofABA in stomatal closure in water-stressed whole plantshas, however, been difficult to decipher with certainty.This is because ABA is ubiquitous, often occurring inhigh concentrations in nonstressed tissue. Also, someearly studies indicated that stomata would begin to closebefore increases in ABA content could be detected.

In most well-watered plants, ABA appears to besynthesized in the cytoplasm of leaf mesophyll cells but,

because of intracellular pH gradients, ABA accumulatesin the chloroplasts (Figure 13.9). At low pH, ABA existsin the protonated form ABAH, which freely permeatesmost cell membranes. The dissociated form ABA− isimpermeant; because it is a charged molecule it doesnot readily cross membranes. Thus, ABAH tends todiffuse from cellular compartments with a low pH intocompartments with a higher pH. There, some of itdissociates to ABA− and becomes trapped. It is wellestablished that in actively photosynthesizing mesophyllcells the cytosol will be moderately acidic (pH 6.0 to 6.5)while the chloroplast stroma is alkaline (pH 7.5 to 8.0)(Chapter 7). It has been calculated that if the stroma pHis 7.5 and cytosolic pH is 6.5, the concentration of ABAin the chloroplasts will be about tenfold higher than inthe cytosol.

According to the current model, the initial detec-tion of water stress in leaves is related to its effects onphotosynthesis, described earlier in this chapter. Inhibi-tion of electron transport and photophosphorylation inthe chloroplasts would disrupt proton accumulation inthe thylakoid lumen and lower the stroma pH. At thesame time, there is an increase in the pH of the apoplastsurrounding the mesophyll cells. The resulting pH gra-dient stimulates a release of ABA from the mesophyllcells into the apoplast, where it can be carried in thetranspiration stream to the guard cells (Figure 13.10).

Just how ABA controls turgor in the guard cellsremains to be determined. Evidence indicates that ABAdoes not need to enter the guard cell, but acts instead onthe outer surface of the plasma membrane. PresumablyABA interacts with the high-affinity binding sites on theplasma membrane (see Chapter 21), although the exis-tence of such sites has yet to be confirmed. Nonetheless,there are strong indications that ABA interferes withplasma membrane proton pumps and, consequently, theuptake of K+, or that it stimulates K+ efflux from theguard cells. Either way, the guard cells will lose turgor,leading to closure of the stomata.

H+ H+

LIGHT DARK

ABAH ABAH

pH 7.5 pH 6.5

H+ + ABA− ABAH

hν

H+ H+

StromaStroma

Cytosol

ThylakoidThylakoid

ChloroplastChloroplast

(pH6.5)

FIGURE 13.9 ABA storage in chloroplasts.In the light, photosynthesis drives protonsinto the interior of the thylakoid, creatinga pH gradient between the stroma and thecytosol. The pH gradient favors movementof ABAH into the chloroplast, where it dis-sociates to ABA−. The membrane is lesspermeable to ABA−. In the dark, protonsleak back into the stroma, the pH gradientcollapses, and ABAH moves back into thecytosol.

232 Chapter 13 / Responses of Plants to Environmental Stress

H2O

Substomatal airspace

Apoplast Symplast

Cuticle

FIGURE 13.10 ABA movement in the apoplast. ABA syn-thesized in the roots is carried to the leaf mesophyll cells(heavy arrows) in the transpiration stream (light arrows).ABA equilibrates with the chloroplasts of the photosyn-thetic mesophyll cells or is carried to the stomatal guardcells in the apoplast.

As noted above, wilted leaves accumulate largequantities of ABA. In most cases, however, stomatalclosure begins before there is any significant increase inthe ABA concentration. This can be explained by therelease of stored ABA into the apoplast, which occursearly enough and in sufficient quantity—the apoplastconcentration will at least double—to account for initialclosure. Increased ABA synthesis follows and serves toprolong the closing effect.

Stomatal closure does not always rely on the per-ception of water deficits and signals arising within theleaves. In some cases it appears that the stomata closein response to soil desiccation before there is any mea-surable reduction of turgor in the leaf mesophyll cells.Several studies have indicated a feed-forward controlsystem that originates in the roots and transmits infor-mation to the stomata. In these experiments, plants aregrown such that the roots are equally divided betweentwo containers of soil (Figure 13.11). Water deficitscan then be introduced by withholding water from

FIGURE 13.12 Stomatal closure in a split-root exper-iment. Maize (Zea mays) plants were grown asshown in Figure 13.11. Control plants (open circles)had both halves of the root system well-watered.Water was withheld from half the roots of theexperimental plants (closed circles) on day zero.Stomatal opening, measured as leaf conductance,declined in the plants with water-stressed roots.(From Blackman, P. G., W. J. Davies. 1985. Jour-nal of Experimental Botany 36:39–48. Reprinted bypermission of The Company of Biologists, Ltd).

Time (days)

0.0

0.1

0.2

7

Leaf

con

duct

ance

(cm

s−1

)

6543210−1−2−3

0.3

FIGURE 13.11 An experimental setup for testing theeffects of desiccated roots on ABA synthesis and stom-atal closure. Roots of a single plant are divided equallybetween two containers. Water supplied to one containermaintains the leaves in a fully turgid state while water iswithheld from the second container. Withholding waterfrom the roots leads to stomatal closure, even though theleaves are not stressed.

one container while the other is watered regularly.Control plants receive regular watering of both con-tainers. Stomatal opening along with factors such asABA levels, water potential, and turgor are comparedbetween half-watered plants and fully watered controls.Typically, stomatal conductance, a measure of stom-atal opening, declines within a few days of withholdingwater from the roots (Figure 13.12), yet there is nomeasurable change in water potential or loss of turgorin the leaves. In experiments with day flower (Commelinacommunis), there was a significant increase in ABA con-tent of the roots in the dry container and in the leafepidermis (Figure 13.13). Furthermore, ABA is readilytranslocated from roots to the leaves in the transpirationstream, even when roots are exposed to dry air. Theseresults provide reasonably good evidence that ABA isinvolved in a kind of early warning system that com-municates information about soil water potential to theleaves.

13.5 Plants are Sensitive to Fluctuations in Temperature 233

Percentage of original fresh weight

20

AB

A (

ng/1

00

mg

d.w

t)

100 90 80 70 65 60

40

60

80

100

120

60

80

100

RW

C (

%)

FIGURE 13.13 Effect of air drying on the ABA con-tent of Commelina communis root tips. Root tipswere air dried to the relative water contents shownin the upper curve. Lower curve shows the dra-matic increase in ABA content as the fresh weightdecreases. (From J. Zhang, W. J. Davies, Journal ofExperimental Botany 38:2015–2023, 1987. Reprintedby permission of The Company of Biologists, Ltd.)

Hormones other than ABA may also be involvedin communication between water-stressed roots andleaves. In an experiment with half-watered maize (Zeamays) plants, results similar to those with Commelinawere obtained. One notable exception was that the ABAcontent of the leaves did not increase, and the applicationof cytokinins to the leaves prevented stomatal closure.At least in Zea, it appears that closure is brought aboutby decreased movement of cytokinins out of the dryingroots.

Both hydropassive and hydroactive closure of stom-ata represent mechanisms that enable plants to anticipatepotential problems of water availability through eitherexcessive transpirational loss from the leaves or chronic,but nonlethal, soil water deficit. Although considerableprogress has been achieved in this field over the lastdecade, there is clearly much yet to be learned aboutstomatal behavior and the response of plants to waterstress.

13.5 PLANTS ARE SENSITIVETO FLUCTUATIONSIN TEMPERATURE

Plants exhibit a wide range of sensitivities to extremesof temperature. Some are killed or injured by moder-ate chilling temperatures while others can survive. Eachplant has its unique set of temperature requirementsfor growth and development (Chapter 27). There is anoptimum temperature at which each plant grows anddevelops most efficiently, and upper and lower limits(Chapter 12). As the temperature approaches these lim-its, growth diminishes, and beyond those limits there isno growth at all. Except in the relatively stable climates

of tropical forests, temperatures frequently exceed theselimits on a daily or seasonal basis, depending on the envi-ronment. Deserts, for example, are characteristically hotand dry during the day but experience low night tem-peratures because, in the absence of a moist atmosphere,much of this heat is reradiated into space. Plants at highaltitudes, where much of the daily heat gain is radiatedinto the thin atmosphere every night, experience similartemperature excursions. Plants native to the northerntemperate and boreal forests must survive temperaturesas low as −70◦C every winter. How plants respond totemperature extremes has long captivated plant biolo-gists. In this section, we will consider how chilling stressand high temperature stress effect the physiology of theplant.

13.5.1 MANY PLANTS ARE CHILLINGSENSITIVE

Plants native to warm habitats are injured when exposedto low, nonfreezing temperatures and are considered tobe chilling sensitive. Plants such as maize (Zea mays),tomato (Lycopersicon esculentum), cucumber (Cucurbitasp.), soybean (Glycine max), cotton (Gossypium hirsutum),and banana (Musa sp.) are particularly susceptible andwill exhibit signs of injury when exposed to tempera-tures below 10 to 15◦C. Even some temperate plantssuch as apple (Malus sp.), potato (Solanum tuberosum),and asparagus (Asparagus sp.) experience injury at tem-peratures above freezing (0 to 5◦C).

Outward signs of chilling injury can take a varietyof forms, depending on the species and age of the plantand the duration of the low-temperature stress. Youngseedlings typically show signs of reduced leaf expan-sion, wilting, and chlorosis. In extreme cases, browning

234 Chapter 13 / Responses of Plants to Environmental Stress

and the appearance of dead tissue (necrosis) and/ordeath of the plant will result. In some plants, repro-ductive development is especially sensitive to chillingtemperature. Exposure of rice plants, for example, tochilling temperatures at the time of anthesis (floralopening) results in sterile flowers. Symptoms of chillinginjury reflect a wide range of metabolic dysfunctionsin chilling-sensitive tissues, including: impaired proto-plasmic streaming, reduced respiration, reduced ratesof protein synthesis as well as altered patterns of pro-tein synthesis. One of the immediate plant responsesto a chilling stress is the light-dependent inhibition ofphotosynthesis. Because low temperature inhibits theD1 repair cycle, this leads to chronic photoinhibitionof PSII and PSI in cucumber. Indeed, there appearto be few aspects of cellular biochemistry that are notimpaired in chilling-sensitive tissues following exposureto low temperature.

One explanation for this is that low tempera-ture causes reversible changes in the physical stateof cellular membranes. Membrane lipids consist pri-marily of diacylglycerides containing two fatty acids ofeither 16- or 18-carbon atoms. Some fatty acids areunsaturated, which means that they have one or morecarbon-carbon double bonds (—CH = CH—), whileothers are fully saturated with hydrogen (—CH2—CH2—). Because saturated fatty acids—and lipids thatcontain them—solidify at higher temperatures thanunsaturated fatty acids, the relative proportions of unsat-urated and saturated fatty acids in membrane lipids havea strong influence on the fluidity of membranes. Achange in the membrane from the fluid state to a gel (orsemicrystalline) state is marked by an abrupt transitionthat can be monitored by a variety of physical meth-ods. The temperature at which this transition occurs isknown as the transition temperature.

Chilling-sensitive plants tend to have a higherproportion of saturated fatty acids (Table 13.1) anda correspondingly higher transition temperature. Formitochondrial membranes of the chilling-sensitive plantmung bean (Vigna radiata), for example, the transitiontemperature is 14◦C. Mung bean seedlings grow poorlybelow 15◦C. Chilling-resistant species, on the otherhand, tend to have lower proportions of saturated fattyacids and, therefore, lower transition temperatures. Dur-ing acclimation to low temperature, the proportion ofunsaturated fatty acids increases and transition temper-ature decreases.

The net effect of the transition from a liquidmembrane to a semicrystalline state at low temperatureis similar to the effects of water stress described above.The integrity of membrane channels is disrupted,resulting in loss of compartmentation and soluteleakage, and the operation of integral proteins thatmake up respiratory assemblies, photosystems, andother membrane-based metabolic processes is impaired.

TABLE 13.1 Ratio of unsaturated/saturatedfatty acids of membrane lipids of mitochondriaisolated from chilling-sensitive andchilling-resistant tissues.

Chilling-sensitive tissuesPhaseolus vulgaris (bean) shoot 2.8Ipomoea batatas (sweet potato) tuber 1.7Zea mays (maize) shoot 2.1Lycopersicon esculentum (tomato) green fruit 2.8Chilling-resistant tissuesBrassica oleracea (cauliflower) buds 3.2Brassica campestris (turnip) root 3.9Pisum sativum (pea) shoot 3.8

From data of J. M. Lyons et al., Plant Physiology 39:262, 1964.

Membranes of chilling-resistant are able to maintainmembrane fluidity to much lower temperatures andthereby protect these critical cellular functions againstdamage. However, it is important to appreciate thatthese differences in membrane lipid unsaturation, bythemselves, cannot fully account for the differencesin chilling sensitivity between chilling resistant andchilling sensitive plant genotypes.

13.5.2 HIGH-TEMPERATURESTRESS CAUSES PROTEINDENATURATION

The traditional view is that the high-temperature limitfor most C3 plants is determined by irreversible pro-tein denaturation of enzymes. Reactions in the thylakoidmembranes of higher plant chloroplasts are most sen-sitive to high-temperature damage, with consequenteffects on the efficiency of photosynthesis. Photosys-tem II and its associated oxygen-evolving complex(Chapter 7) are particularly susceptible to injury. Theoxygen-evolving complex is directly inactivated by heat,thereby disrupting electron donation to PSII resultingin the accumulation of P680+, the strongest oxidiz-ing agent in nature (Chapter 7). The result is that anincreasing portion of the absorbed energy cannot beused photochemically by PSII which leads to chronicphotoinhibition. This is easily monitored in intact leavesusing chlorophyll fluorescence (Box 13.1). Other stud-ies have indicated that the activities of Rubisco, Rubiscoactivase, and other carbon-fixation enzymes, may alsobe severely compromised at high temperatures.

Exposure of most organisms to supraoptimal tem-peratures for brief periods also suppresses the synthesisof most proteins including the PSII reaction cen-ter polypeptide, D1. This leads to an inhibition ofthe D1 repair cycle that is critically important toovercome the effects of chronic photoinhibition. In

13.6 Insect Pests and Disease Represent Potential Biotic Stresses 235

contrast, high-temperature stress also induces the syn-thesis of a new family of low molecular mass proteinsknown as heat shock proteins (HSPs). This interestingclass of proteins was originally discovered in Drosophilamelanogaster (fruit fly) but they have since been discov-ered in a variety of animals, plants, and microorganisms.Exposures in the range of 15 minutes to a few hours attemperatures 5◦C to 15◦C above the normal growingtemperature are usually sufficient to cause full inductionof HSPs. HSPs are either not present or present at verylow levels in nonstressed tissues. Initially, interest inHSPs centered on their potential for the study of generegulation. There are, however, several aspects of HSPsthat are of physiological interest.

There are three distinct classes of HSPs in higherplants, based on their approximate molecular mass:HSP90, HSP70, and a heterogeneous group with amolecular mass in the range of 17 to 28 kDa (Table 13.2).One in particular, HSP70, has a high degree of struc-tural similarity—about 70 percent identical—in bothplants and animals. Another protein, ubiquitin, is alsofound in all eukaryote organisms subjected to heat stress

TABLE 13.2 Principal heat shock proteins(HSP) found in plants and their probable functions.Families are designated by their typical molecularmass. The number and exact molecular mass ofproteins in each family vary depending on plantspecies.

HSP Family Probable Function

HSP 110 Unknown.HSP 90 Protecting receptor proteins.HSP 70 ATP-dependent protein assembly or

disassembly reactions; preventingprotein from denaturation oraggregation (molecular chaperone).Found in cytoplasm, mitochondria, andchloroplasts.

HSP 60 Molecular chaperone, directing theproper assembly of multisubunitproteins. Found in cytoplasm,mitochondria, and chloroplasts.

LMW HSPs(17–28 kDa)

Function largely unknown. LMW(low-molecular-weight) HSPs reversiblyform aggregates called ‘‘heat shockgranules’’. Found in cytoplasm andchloroplasts.

Ubiquitin An 8 kDa protein involved in targetingother proteins for proteolyticdegradation.

Based on Vierling, 1990. E. Vierling. 1990. Heat shock proteinfunction and expression in plants. In: R. G. Alscher, J. R. Cumming(eds.), Stress Responses in Plants: Adaptation and Acclimation Mechanisms.New York: Wiley-Liss, pp. 357–375.

and is considered a HSP. Ubiquitin has an importantrole in marking proteins for proteolytic degradation.HSPs are found throughout the cytoplasm as well as innuclei, chloroplasts, and mitochondria. As well, induc-tion of HSPs does not require a sudden temperatureshift: they have been detected in field-grown plants fol-lowing more gradual temperature rises of the sort thatmight be expected under normal growing conditions.

HSPs are synthesized very rapidly following anabrupt increase in temperature; new mRNA transcriptscan be detected within 3 to 5 minutes and HSPs formthe bulk of newly synthesized protein within 30 minutes.Within a few hours of return to normal temperature,HSPs are no longer produced and the pattern of pro-tein synthesis returns to normal. The speed of theirappearance suggests that HSPs might have a criticalrole in protecting the cell against deleterious effects ofrapid temperature shifts. HSP70, for example, appearsto function as a molecular chaperone, or chaperonin.Chaperonins are a class of proteins normally present inthe cell that direct the assembly of multimeric proteinaggregates. There is in the chloroplast, for example, aRubisco-binding protein (HSP60) that helps to assemblethe large and small subunits of Rubisco into a functionalenzyme. It has been suggested that HSP70 functionsto prevent the disassembly and denaturation of multi-meric aggregates during heat stress. At the same time,increased ubiquitin levels reflect an increased demandfor removal of proteins damaged by the heat shock.

In nature, high temperature stress is usually associ-ated with water stress. Since both of these stresses inhibitphotosynthesis, both of these stresses will concomitantlypredispose the plant to photoinhibition. This illustratesthe interactive effects of abiotic stresses to which plantsare constantly exposed in a natural, fluctuating envi-ronment. Furthermore, exposure of plants to abioticstresses such as drought and temperature extremes usu-ally increases the susceptibility of these plants to attackby insects and plant diseases.

13.6 INSECT PESTS AND DISEASEREPRESENT POTENTIALBIOTIC STRESSES

Typically, a plant challenged by insects or potentiallypathogenic microorganisms responds with changes inthe composition and physical properties of cell walls,the biosynthesis of secondary metabolites that serve toisolate and limit the spread of the invading pathogen.These responses are collectively known as a hypersen-sitive reaction.

The hypersensitive reaction is commonly activatedby viruses, bacteria, fungi, and nematodes and occursprincipally in plants outside the pathogen’s normalspecificity range. Although the hypersensitive reaction

236 Chapter 13 / Responses of Plants to Environmental Stress

is complex and can vary depending on the nature ofthe causal agent, there are common features that gen-erally apply. An early event in this sensing/signalingpathway is the activation of defense-related genes andsynthesis of their products, pathogenesis-related (PR)proteins. PR proteins include proteinase inhibitors thatdisarm proteolytic enzymes secreted by the pathogenand lytic enzymes such as β-1,3-glucanase and chiti-nase that degrade microbial cell walls. Also activatedare genes that encode enzymes for the biosynthesisof isoflavonoids and other phytoalexins that limit thegrowth of pathogens. Lignin, callose, and suberin areaccumulated in cell walls along with hydroxyproline-richglycoproteins that are believed to provide structuralsupport to the wall. These deposits strengthen the cellwall and render it less susceptible to attack by theinvading pathogen. Finally, the invaded cells initiateprogrammed cell death, a process that results in theformation of necrotic lesions at the infection site. Cellnecrosis isolates the pathogen, slowing both its devel-opment and its spread throughout the plant. It is notclear at this time to what extent these components of thehypersensitive reaction are sequential or parallel events.

13.6.1 SYSTEMIC ACQUIREDRESISTANCE REPRESENTS APLANT IMMUNE RESPONSE

Some secondary metabolites associated with the hyper-sensitive reaction appear to constitute signal transduc-tion pathways that prepare other cells and tissues toresist secondary infections. Initially the hypersensitivereaction is limited to the few cells at the point of inva-sion, but over a period of time, ranging from hours todays, the capacity to resist pathogens gradually becomesdistributed throughout the entire plant. In effect, theplant reacts to the initial infection by slowly developinga general immune capacity. This phenomenon is knownas systemic acquired resistance (SAR).

The development of SAR is still not completelyunderstood, but one component of the signaling path-way appears to be salicylic acid (Figure 13.14). Salicylic

COOH

OH O

COOH

CH3C

O

Salicylicacid

Aspirin(acetylsalicylic acid)

FIGURE 13.14 The chemical structure of salicylic acidand its commercial derivative acetylsalicylic acid. Sali-cylic acid has been implicated in the immune strategiesof plants.

acid (2-hydroxybenzoic acid) is a naturally occurringsecondary metabolite with analgesic properties. NativeNorth Americans and Eurasians have long used willowbark (Salix sps.), a source of the salicylic acid glyco-side, salicin, to obtain generalized relief from aches andpains.

The relationship between salicylic acid and resis-tance to pathogens did not become apparent until theearly 1990s, when it was observed that both salicylicacid and its acetyl derivative (aspirin), when appliedto tobacco plants, induced PR gene expression andenhanced resistance to tobacco mosaic virus (TMV).Since then, it has been shown in a variety of plantsthat infection is followed by increased levels of sali-cylic acid both locally and in distal regions of the plant(Figure 13.15). For example, when tobacco plants areinoculated with TMV, the salicylic acid level rises asmuch as 20-fold in the inoculated leaves and 5-fold inthe noninfected leaves. Furthermore, the appearance ofPR proteins rises in parallel with salicylic acid. The risein salicylic acid levels usually precedes the developmentof SAR. There are also a number of Arabidopsis mutantsand transgenic plants that are characterized by consti-tutively high levels of both salicylic acid and SAR and,consequently, enhanced resistance to pathogens.

On the other side of the coin, plants with artificiallylow levels of salicylic acid generally fail to establishSAR. For example, bacteria have a gene designatednahG that encodes the enzyme salicylate hydroxylase.Arabidopsis plants transformed with the nahG gene thuscontain little or no salicylic acid. Plants transformedwith the nahG gene also fail to establish SAR andare compromised in their ability to ward off pathogenattack. Salicylic acid levels can also be reduced by directinhibition of the enzyme phenylalanine-ammonia lyase(PAL), which catalyzes the first step in the biosynthesisof salicylic acid. PAL-limited Arabidopsis plants lose their

Pathogen

Pathogen

HR

Primary infection

SAS

SASASASA

MSAMSASS

MSA

Secondary infection

FIGURE 13.15 The possible role of salicylic acid in sys-temic acquired resistance (SAR). The first pathogens toinfect the plant (primary infection) stimulate a localizedhypersensitive reaction (HR) and the synthesis of sali-cylic acid (SA). Salicylic acid is translocated through thephloem to other regions of the plant where it preventssecondary infection by other pathogens. Alternatively,salicylic acid may be converted to methylsalicylic acid(MSA). MSA is moderately volatile and may function asan airborne signal.

13.7 There are Features Common to all Stresses 237

resistance to disease, but resistance can be restored byapplying salicylic acid. Based on results such as these, itis clear that salicylic acid has a significant role in plantdefense responses. However, the mechanism wherebysalicylic acid establishes and maintains SAR is yet to bedetermined.

13.6.2 JASMONATES MEDIATE INSECTAND DISEASE RESISTANCE

On the basis of recent experiments, it appears that jas-monates, especially jasmonic acid and its methyl ester(methyljasmonate) (Figure 13.16), also mediate insectand disease resistance. Jasmonates have been found tooccur throughout plants, with highest concentrations inyoung, actively growing tissues. Methyljasmonate is theprincipal constituent of the essential oil of Jasminiumand high concentrations of jasmonic acid have beenisolated from fungal culture filtrates.

There are some similarities in the action of sal-icylic acid and jasmonates with respect to insect anddisease resistance, but there are also some importantdistinctions. In a study of two fungal resistance genesin Arabidopsis, for example, it was found that expres-sion of one gene was induced by salicylic acid, but notjasmonic acid, while the second gene was induced by jas-monic acid but not salicylic acid. Apparently there are atleast two defensive pathways, one mediated by salicylicacid and one mediated by jasmonates. Jasmonic acid issynthesized from the unsaturated fatty acid, linolenicacid, which has led to the proposal that jasmonic acidfunctions as a type of second messenger.

Another very interesting but somewhat complicat-ing aspect of jasmonates is that their action is not limitedto insect and disease resistance. Through their effect ongene expression, jasmonates modulate a number of otherphysiological processes. These include seed and pollengermination, vegetative protein storage, root develop-ment, and tendril coiling. In most of these effects, thejasmonates appear to work in concert with ethylene.

O CH3

O

COOH

C

FIGURE 13.16 The chemical structures of jasmonic acid(above) and methyljasmonate (below). Jasmonic acid issynthesized from linolenic acid (18:3).

This breadth of jasmonate effects has led some to sug-gest that jasmonates should be elevated to the status ofplant hormones.

13.7 THERE ARE FEATURESCOMMON TO ALL STRESSES

The maintenance of cellular homeostasis is the result ofa complex network of genetically regulated biochemi-cal pathways. Thus, modulation of cellular homeostasisby abiotic and biotic stresses that we have discussedin this chapter can be detected at all levels of cellu-lar organization—from genes to physiological function.Thus, any stress may induce or repress specific setsof genes or gene families through the regulation oftranscription which will reflect changes in protein com-plement within the cell. Many of these proteins willbe enzymes which catalyze specific reactions and con-sequently their presence or absence will alter cellularphysiology. Because homeostasis is a complex networkof interacting genes and biochemical pathways, differ-ent stresses may affect common sets of genes or genefamilies which is consistent with the observation thatthere appears to be overlap or cross talk between sig-nal transduction pathways involved in the physiologicalresponse of plants to various stresses.

All stresses discussed in this chapter inhibit pho-tosynthesis in one way or another, which may lead tochronic photoinhibition. Since chlorophyll fluorescence(Box 13.1) can be exploited as an intrinsic probe ofthe overall function of photosynthesis, chlorophyll flu-orescence is the most widely used technique in plantstress research. Since photosynthesis is extremely sen-sitive to environmental stress, it should not surprise usthat all abiotic and biotic stresses have a negative effecton plant productivity and survival (Table 13.3) (also seeChapter 12). If one assumes that the record yield rep-resents the maximal yield under near-optimal, naturalgrowth conditions, then the difference between theaverage yields and the record yields may be inter-preted to indicate the average losses in yield due tosuboptimal growth conditions as consequence of thecombined effects of abiotic and biotic stresses. The datain Table 13.3 illustrate that environmental stresses havea staggering effect on the crop yields of corn, wheat,sorghum, and potato. The combined effects of abi-otic and biotic stress reduces the yield of these cropsby 70 to 87%! This should be cause for concern forregarding future world food production in the contextof climate change predictions coupled with continuedhuman population growth. However, plants also exhibitan astounding capacity to acclimate to myriad environ-mental conditions. The capacity of plants to sense andsubsequently acclimate on a short-term and long-termbasis to environmental stress is the subject of Chapter 14.

238 Chapter 13 / Responses of Plants to Environmental Stress

TABLE 13.3 The effects of abiotic and biotic stress on average crop yields.

Crop Record Yield (kg/hectare) Average Yield (kg/hectare) Average Loss (kg/hectare) (% loss)

Maize 19,300 4,600 14,700 76Wheat 14,500 1,880 12,620 87Sorghum 20,100 2,830 17,270 86Potato 94,100 28,300 65,800 70

Adapted from Bray, E. A., J. Bailey-Serres, E. Weretilnyk. 2000. Responses to Abiotic Stresses. Biochemistry and MolecularBiology of Plants, pp. 1158–1203. Rockville, MD: American Society of Plant Physiologists.

SUMMARY

The maintenance of a cellular steady-state far fromequilibrium results in an apparently stable conditioncalled homeostasis. Environmental modulation ofhomeostasis may be defined as biological stress. Plantstress usually implies some adverse effect on the physi-ology of a plant. Plants may respond to stress in severalways.

Susceptible plants succumb to a stress, other plantsavoid stress by completing their life cycle during peri-ods of relatively low stress whereas stress resistantplants are able to tolerate a stress. Although light isrequired for the photosynthetic assimilation of CO2,too much light can inhibit photosynthesis and resultsin photodamage from chronic photoinhibition. Onerole of the D1 repair cycle is to overcome the effectsof photodamage to PSII reaction centers. Water stressin natural environments usually arises due to lack ofrainfall, a condition known as drought. Thus, waterdeficit stress is often referred to as drought stress.This leads to desiccation of the protoplasm and cel-lular dysfunction. An immediate response of mostplants to water stress is stomatal closure due to lowturgor in the guard cells. Stomatal closure is triggeredby decreasing water potential in the leaf mesophyll.The hormone abscisic acid (ABA) appears to have asignificant role in stomatal closure. Chilling stressrefers to exposure of plants to temperatures near butabove the freezing point of water. The membranesof chilling-sensitive plants tend to have a higher pro-portion of unsaturated fatty acids and, consequently,change from a fluid to semicrystalline gel state at highertemperatures than chilling-resistant plants. The uppertemperature limit for most plants is determined by acombination of irreversible denaturation of enzymesand problems with membrane fluidity. Plants subjectedto heat stress respond by synthesizing a new familyof low-molecular-weight heat shock proteins. Plantsrespond to insect damage and microbial pathogeninfection with a hypersensitive reaction. The hyper-sensitive reaction includes changes in the compositionand increased strength of the cell wall and the forma-tion of necrotic lesions at the site of infection. These

responses serve to isolate the potential pathogen andprevent its development and spread through the plant.Salicylic acid or its methyl ester may serve as a mobilesignal, participating in systemic acquired resistance, aform of generalized immune response. Another pos-sible signaling agent is jasmonic acid, a derivative ofthe fatty acid linolenic acid. A common feature of allstresses is that they induce or repress specific genes orgene families and they affect photosynthesis negatively.As a consequence, the combination of abiotic and bioticstresses reduces plant productivity and crop yield.

CHAPTER REVIEW

1. Define homeostasis. Define environmental stress.2. If plants require light for photosynthesis, explain

why plants can be exposed to too much light.3. What is the role of the D1 repair cycle?4. Describe how plants may be injured by water

stress.5. How does stomatal closure come about in response

to water stress?6. How do chilling-sensitive and a chilling-tolerant

plants differ in their response to suddenexposures to low, nonfreezing temperatures?

7. What are heat shock proteins?8. Define biotic stress. What are the roles of salicylic

acid and jasmonates in a plant’s response to bioticstress?

9. What are two common features of all stresses?

FURTHER READING

Bostock, R. M. 2005. Signal crosstalk and induced resistance:Straddling the line between cost and benefit. AnnualReview of Phytopathology 43:545–580

Buchanan, B. B., W. Gruissem, R. L. Jones. 2000. Biochem-istry and Molecular Biology of Plants. Rockville MD: Amer-ican Society of Plant Physiologists.

Further Reading 239

Creelman, R. A., J. E. Mullet. 1997. Biosynthesis and actionof jasmonates in plants. Annual Review of Plant Physiologyand Plant Molecular Biology 48:355–381.

Demmig-Adams, B., W. W. Adams III, A. Mattoo. 2006.Photoprotection, Photoinhibition, Gene Regulation and Envi-ronment. Advances in Photosynthesis and Respiration, Vol.21. Dordrecht: Springer.

Harwood, J. L. 1998. Involvement of chloroplast lipids inthe reaction of plants submitted to stress. Lipids in Pho-tosynthesis: Structure, Function and Genetics. Advances inPhotosynthesis, Vol 6, pp. 287–302. Dordrecht: KluwerAcademic Publishers.

Howe, G. A., G. Jander. 2008. Plant immunity to insect her-bivores. Annual Review of Plant Biology 59:41–66.

Iba, K. 2002. Acclimative response to temperature stressin higher plants: Approaches of gene engineering for

temperature tolerance. Annual Review of Plant Biology53:224–245.

Ingram, J., D. Bartels. 1996. The molecular basis of dehydra-tion tolerance in plants. Annual Review of Plant Physiologyand Plant Molecular Biology 47:377–403.

Kessler, A., I. T. Baldwin. 2002. Plant responses to insect her-bivory: The emerging molecular analysis. Annual Reviewof Plant Biology 53:299–328.

Melis, A. 1999. Photosystem-II damage and repair cycle inchloroplasts: What modulates the rate of photodamagein vivo? Trends in Plant Science 4:130–135.

Nishida, I., N. Murata. 1996. Chilling sensitivity in plantsand cyanobacteria: The crucial contribution of mem-brane lipids. Annual Review of Plant Physiology and PlantMolecular Biology 47:541–568.

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NuclearGene

Expression

Chloroplast Mitochondrion

PSII

PQH2

PQ

PSI Complex I

ComplexIV

UQH2

UQ

Nucleus

hν hν

14Acclimation to Environmental Stress

In Chapter 13 we defined stress as a negative effecton plant homeostasis which can be measured when aplant is exposed to a sudden change from an opti-mal condition for growth to some suboptimal abioticcondition such high light, low temperature, drought,or some biotic infestation. Many stress-sensitive plantsmay succumb to such a stress and die. However, in thischapter we focus on plants that exhibit the capacity totolerate a particular stress over time. Such plants arecalled stress resistant or stress tolerant and reflectsthe ability of these plants to acclimate or adjust to thestress. The capacity of a plant to acclimate is, of course,a genetic trait. However, the specific changes broughtabout in response to stress are not themselves passed onto the next generation and thus are nonheritable. Thecapacity of plants to acclimate to changing environmen-tal conditions reflects their remarkable plasticity. As aconsequence, a plant’s physiology and morphology arenot static but are very dynamic and responsive to theirenvironment. The ability of biennial plants and wintercultivars of cereal grains to survive over winter is anexample of acclimation to low temperature. The processof acclimation to a stress is known as hardening andplants that have the capacity to acclimate are commonlyreferred to as hardy species. In contrast, those plants

that exhibit a minimal capacity to acclimate to a spe-cific stress are referred to as nonhardy species. Thus,frost-hardy plants are those that are able to acclimateto low temperature and are able to survive the freezingstress of winter, and drought-hardy plants are able tosurvive water stress. The physiological bases of acclima-tion to various environmental stresses will be the focusof this chapter and will include a discussion of

• the theoretical basis of plant acclimation and plas-ticity as a time-nested phenomenon,

• the role of state transitions in response to changesin light quality,

• the xanthophyll cycle and protection against photo-damage,

• the mechanisms that allow plants to survive waterlimitations,

• low temperature acclimation and freezing tolerance,• excitation pressure as a redox signal for retrograde

regulation of nuclear genes,• photosynthetic acclimation to high temperature,

and• the protective role of O2 as an alternative photo-

synthetic electron acceptor during acclimation.

241

242 Chapter 14 / Acclimation to Environmental Stress

14.1 PLANT ACCLIMATIONIS A TIME-DEPENDENTPHENOMENON

As discussed in Chapter 13, a plant stress usuallyreflects some sudden change in environmental condi-tion. However, in stress-tolerant plant species, exposureto a particular stress leads to acclimation to that spe-cific stress in a time-dependent manner (Figure 14.1).Thus, plant stress and plant acclimation are intimatelylinked with each other. The stress-induced modulationof homeostasis can be considered as the signal for theplant to initiate processes required for the establishmentof a new homeostasis associated with the acclimatedstate. Plants exhibit stress resistance or stress toler-ance because of their genetic capacity to adjust or toacclimate to the stress and establish a new homeostaticstate over time. Furthermore, the acclimation process instress-resistant species is usually reversible upon removalof the external stress (Figure 14.1).

The establishment of homeostasis associated withthe new acclimated state is not the result of a sin-gle physiological process but rather the result of manyphysiological processes that the plant integrates overtime, that is, integrates over the acclimation period.Plants usually integrate these physiological processesover a short-term as well as a long-term basis. Theshort-term processes involved in acclimation can beinitiated within seconds or minutes upon exposure toa stress but may be transient in nature. That meansthat although these processes can be detected very soonafter the onset of a stress, their activities also disappearrather rapidly. As a consequence, the lifetime of theseprocesses is rather short (Figure 14.2, processes a, band c). In contrast, long-term processes are less transient

and thus usually exhibit a longer lifetime (Figure 14.2,processes d, e and f). However, the lifetimes of theseprocesses overlap in time such that the short-term pro-cesses usually constitute the initial responses to a stresswhile the long-term processes are usually detected laterin the acclimation process. Such a hierarchy of short-and long-term responses indicates that the attainmentof the acclimated state can be considered a complex,time-nested response to a stress. Acclimation usuallyinvolves the differential expression of specific sets ofgenes associated with exposure to a particular stress.The remarkable capacity to regulate gene expressionin response to environmental change in a time-nestedmanner is the basis of plant plasticity.

14.2 ACCLIMATION IS INITIATEDBY RAPID, SHORT-TERMRESPONSES

In this section, we will discuss examples of initial, rapidresponses to changes in light, water availability andtemperature that are part of the acclimation process.

14.2.1 STATE TRANSITIONS REGULATEENERGY DISTRIBUTIONIN RESPONSE TO CHANGESIN SPECTRAL DISTRIBUTION

Optimal CO2 reduction requires an efficient supplyof NADPH and ATP that would, in turn, require asteady, balanced electron flow through PSII and PSI(Chapter 7). But if light is not saturating and the deliveryof excitation energy to PSII and PSI is not balanced,then the rate of electron transport and, consequently,

FIGURE 14.1 A schematic relationship betweenstress and acclimation. Under some optimal envi-ronmental condition, a plant is in a homeostaticstate A as indicated by a constant rate of somephysiological process measured over time. Uponthe imposition of an external stress, the rate ofthis physiological process in most cases decreasesrapidly which indicates a disruption of the home-ostatic state A. Plants that are able to adjust tothis stress over time may establish a new physio-logical rate that is either lower (homeostatic stateB) or higher (homeostatic state C) than the orig-inal rate (homeostatic state A). Such plants arecapable of acclimation and are considered stresstolerant. The plant may remain in this new home-ostatic state B or state C through completion ofits life cycle. Alternatively, the plant may return tothe original homeostatic state A when the stress isremoved (broken lines). The black line indicatesthe most probable acclimation response.

STRESS

Stress

Removed

ACCLIMATION

Time

Homeostatic

Homeostatic

Homestatic

Homeostatic

State A

Rat

e of

a

Phy

siol

ogic

al P

roce

ss

State C

State B

State A

Stress

Removed

14.2 Acclimation is Initiated by Rapid, Short-Term Responses 243

Time

a

o TaTb

TcTd Te

Tf

b c d e f g

Phy

siol

ogic

al A

ctiv

ity

FIGURE 14.2 A schematic graph illustrating plant acclima-tion as a time-nested response. Peaks labeled a, b, c, d,e, f, and g represent the appearance and disappearance ofdifferent physiological processes that are initiated by theonset of a stress at time 0. Arrows labeled Ta, Tb, Tc,Td, Te, Tf, and Tg approximates the lifetime of each pro-cess. The longer the arrow, the longer the lifetime. Thus,process a is the most transient but least stable responsewhereas process g is least transient but most stable physi-ological response to the stress. Processes a to g are said tobe nested in a time with short-term processes occurringfirst, followed by the more long-term processes.

photosynthesis will be limited by the photosystemreceiving the least energy. Under natural conditions, theamount of light driving PSII and PSI is not necessarilybalanced or consistent. For example, less energy isrequired to excite P700 (171 kJ) than P680 (175 kJ).Moreover, because of differences in the number ofantenna molecules, their absorption coefficients, and avariety of other factors that influence absorption, thecapacities of PSII and PSI to absorb light, often referredto as their absorption cross-section (σ ), are not equal.This might be expected to place unique constraints onthe overall efficiency of photosynthesis. State transitionsare a short-term mechanism to regulate excitationenergy distribution between the two photosystems thatis required to maintain an efficient flow of electrons toNADP+.

In addition to the inherent inequities of spectraldistribution, plants often face other situations requir-ing rapid adjustments in the amount of energy beingfed into PSII and PSI. Plants growing in the shadeof a canopy, for example, are frequently subject to

sudden transient fluctuations in fluence rate, knownas sunflecks. A sunfleck is a spot of direct sunlightimpinging on the leaf through an open gap in thecanopy. Sunfleck lifetimes are variable, from a few sec-onds’ duration due to wind flutter up to 20 minutesor longer under woodland canopies. Similar fluctu-ations occur when the sun suddenly reappears afterhaving been blocked by extensive cloud cover. In sit-uations such as these the leaf may be subject to asmuch as tenfold increases in energy that, particularlywhen directed to PSII, could have severe damagingeffects.

State transitions are one of the best-understoodmechanisms for short-term regulation of energy distri-bution and is based on reversible phosphorylation ofLHCII protein (Figure 14.3). The phosphorylation ofproteins is a ubiquitous mechanism for regulating manyaspects of gene regulation and response to environmen-tal stimuli in all eukaryote organisms. The phosphoryla-tion of proteins is catalyzed by a class of enzymes knownas protein kinases. Chloroplasts contain a thylakoid

PS II

PS IILight

PS ILight

Protein kinase

Protein phosphatase

LHC IILHC II

PQ

PQH2

Pi P

++

PS II

FIGURE 14.3 Reversible phosphorylation of LHCII. When PSII is overexcited relativeto PSI, plastoquinol (PQH2) accumulates. A high level of plastoquinol activates a pro-tein kinase that phosphorylates LHCII. Addition of a phosphate group weakens theinteraction between LHCII and the PSII core antenna, causing LHCII to dissociatefrom PSII. The input of light to PSII is diminished and PSII slows down, thus allow-ing PSI to oxidize excess PQH2 to plastoquinone (PQ), which, in turn, deactivatesthe protein kinase. LHCII is dephosphorylated by a protein phosphatase, allowingLCHII to reform in association with PSII.

244 Chapter 14 / Acclimation to Environmental Stress

membrane-bound protein kinase capable of phospho-rylating LHCII. The activity of this kinase is sensitiveto the redox state of the thylakoid membrane and isactivated when excess energy drives PSII, resulting in abuildup of reduced plastoquinone (PQH2). Plants thatare exposed to conditions that result in the preferentialexcitation of PSII are considered to be in state 2.

The resulting phosphorylation of LHCII increasesthe negative charge of the protein, causing LHCII todissociate from PSII. The same negative charge alsoloosens the appression of the thylakoid membranes inthe grana stacks, freeing a certain portion of LHCII tomigrate into the PSI-rich stroma thylakoids. This shiftsthe balance of energy away from the PSII complexes,which remain behind in the appressed region, in favorof PSI. The preferential excitation of PSI is referred toas state 1. As the plastoquinone pool becomes reoxi-dized, due to increased PSI activity, the protein kinase isdeactivated and a protein phosphatase enzyme dephos-phorylates the LHCII, causing it to migrate back into theappressed region and recombine with PSII. Recently, ithas been shown that the PSI-H subunit of Arabidopsisthaliana is required for reversible transitions betweenstate 1 and state 2. In Arabidopsis thaliana lacking thePSI-H subunit due to antisense suppression of the genewhich codes for PSI-H, LHCII is unable to transferenergy to PSI, thus impairing state transitions. Thenet result of these state transitions is a very dynamic,continuous adjustment of excitation energy distribu-tion between PSII and PSI. This continual adjustmentof energy input in turn maintains an optimal flow ofelectrons through the two photosystems.

14.2.2 CAROTENOIDS SERVE A DUALFUNCTION: LIGHT HARVESTINGAND PHOTOPROTECTION

The pigment-protein complexes of thylakoid mem-branes contain not only the chlorophyll pigments,but carotenes and xanthopylls as well (Chapter 6).The principal carotene in most higher plant species isβ-carotene, although smaller amounts of α-carotenemay be present in some species. The principalxanthophylls are lutein, violaxanthin, and zeaxanthin(see Figure 6.11). It appears that carotenoids mayserve two principal functions in photosynthesis: lightharvesting and photoprotection. Primarily on thebasis of action spectra, it has long been believed thatthe principal function of carotenoids is to transferabsorbed light energy to chlorophyll. In this sense, thecarotenoids serve a light-harvesting function and theenergy they absorb is eventually transferred from thelight-harvesting complex to reaction centers for use inphotosynthetic electron transport for generation of ATPand NADPH. In contrast to the light-harvesting roleof carotenoids, there is now substantial evidence that

carotenoids also play an important role in protectingthe photosynthetic system from chronic photo-inhibition. This is called photoprotection. Duringperiods of peak irradiance, plants typically ab-sorb more energy than they can utilize in the reductionof carbon dioxide, that is, they are exposed to excesslight (see Figure 13.4). Rapidly growing crops, forexample, may utilize no more than 50 percent ofabsorbed radiation, while other species, such asevergreens, may utilize as little as 10 percent. Anyexcess absorbed energy must be dissipated. If not, theexcess absorbed energy may lead to photoinhibition ofphotosynthesis (see Chapter 13). Prolonged exposureto photoinhibitory conditions may lead subsequentlyto photodamage and uncontrolled destruction of thephotosynthetic apparatus due to the accumulation ofreactive oxygen species (ROS). These are toxic formsof oxygen that may lead to cell damage and ultimatelycell death.

A unique feature of O2 is that, in its ground state,it is in a triplet state (3O2) rather than the usual singletstate that characterizes almost all other molecules. Thisprevents O2 from chemically reacting with most organicmolecules. Thus, this difference between ground stateO2 and almost all other molecules is a major deterrent tospontaneous combustion and allows life to persist evenin an oxygen-rich environment! However, wheneverthe energy absorbed by the photosynthetic apparatusexceeds the capacity to utilize that energy, there is anincreased probability that singlet excited chlorophyll(1Chl) will be converted to triplet excited chlorophyll(3Chl) (Chapter 6). As a consequence of such an inter-system crossing event, triplet chlorophyll may interactwith ground state O2 and convert ground state oxygento singlet excited O2(1O2). This is one example of atoxic ROS. Clearly light can be a very dangerous formof energy, especially for oxygenic photosynthetic organ-isms. Carotenoids present in thylakoid membranes cancompete with ground state oxygen for triplet excitedchlorophyll and prevent the formation of singlet oxy-gen. Thus, carotenoids such as β-carotene are importantin preventing the formation of toxic reactive oxygenspecies.

Formation of singlet excited O2 in chloroplastscan also be prevented by trapping and dissipatingexcess excitation energy before it reaches the reactioncenter. Recent studies have established an importantlink between the dissipation of excess energy nonpho-tochemically as heat (Box 13.1) and the presence ofthe xanthophyll, zeaxanthin. Zeaxanthin is formed fromviolaxanthin by a process known as the xanthophyllcycle (Figure 14.4A). Violaxanthin is a diepoxide;it contains two epoxy groups, one on each ring.Under conditions of excess light, violaxanthin (V) isenzymatically converted to zeaxanthin (Z) throughthe removal of those two oxygens (de-epoxidation).

14.2 Acclimation is Initiated by Rapid, Short-Term Responses 245

Violaxanthin

Zeaxanthin

CH

CHH3 3

3 3

3

C CH

H

CC

H H

H H H H H H H OHCH

CH3 CH3

33

3

H C CH

O

H C

H

CC

H

CC

H

CC

H

CC

CC

CC

CC

CC

CH

CHH3 3

3 3

3

C CH

HHO

CC

H H

H H H H H H HCH

CH3 CH3

33

3

H C CH

H C

H

CC

H

CC

H

CC

H

CC

CC

CC

CC

CCO

Epo

xida

tion

De-

epox

idat

ion

HO

OH

Antheraxanthin

O

CH

CHH3 3

3 3

3

C CH

HHO

CC

H H

H H H H H H HCH

CH3 CH3

33

3

H C CH

H C

H

CC

H

CC

H

CC

H

CC

CC

CC

CC

CC

OH

Ene

rgy

v *

B.

A.

Low light

Chlb

Chla

P680

P680

Ene

rgy

z

v

C.

High light

Chlb

Chla

P680

HEAT

LOW

LIG

HT

HIG

H L

IGH

T

FIGURE 14.4 The xanthophyll cycle and photoprotection. (A) Under high light,de-epoxidation removes the two oxygen (epoxy) groups from the rings of violax-anthin. This is a two-step reaction with the intermediate antheraxanthin contain-ing only one epoxy group. De-epoxidation is induced by high light, low lumenalpH, and high levels of reduced ascorbate. Note that in zeaxanthin, the number ofcarbon–carbon double bonds is increased by two relative to violaxanthin. The xan-thophyll cycle is reversible since, under low-light conditions, zeaxanthin is convertedback to violaxanthin. (B) Violaxanthin (V) acts as a light-harvesting pigment. Violax-anthin is able to transfer its absorbed excitation energy, via chlorophyll b (Chl b) andchlorophyll a (Chl a) to the PSII reaction center to excite P680. (C) Under excesslight, most of the violaxanthin (V) is converted to zeaxanthin (Z), which is unable totransfer its absorbed excitation energy to the PSII reaction center via chlorophyll b.Zeaxanthin loses its absorbed energy as heat.

De-epoxidation is stepwise—removal of the firstoxygen generates an intermediate monoepoxide(antheraxanthin)(A). De-epoxidation is also induced bya low pH in the lumen, which is a normal consequenceof electron transport under high light conditions.The reaction is reversed in the dark as zeaxanthin isagain enzymatically converted back to violaxanthin.Xanthophyll cycle activity can be assessed by measuringthe de-epoxidation state (DEPS) of the cycle pool

which is a measure of the concentration of zeaxanthin(Z) and antheraxanthin (A) relative to the total pool(V + A + Z) (Equation 14.1).

DEPS = Z + 12 A/V + A + Z (14.1)

Although it has been established that the xan-thophyll cycle plays a key role in photoprotection ofthe chloroplast through nonphotochemical quench-ing (NPQ; Box 13.1), the precise molecular mechanism

246 Chapter 14 / Acclimation to Environmental Stress

is still disputed. Through the generation of an Ara-bidopsis thaliana mutant named npq4, one model suggeststhat the zeaxanthin produced by the xanthophyll cyclemay be bound to a specific PSII polypeptide encodedby the PsbS gene. These observations form the basisfor the hypothesis that the xanthophyll cycle acts as areversible, molecular switch to regulate the capacity forNPQ and the safe dissipation of excess absorbed. Underlow light, LHCII has abundant violaxanthin (V) whoseabsorbed excitation energy is rapidly transferred to thereaction center to excite P680 to P680* (Figure 14.4B).However, exposure to high light rapidly converts toviolaxanthin to zeaxanthin within LHCII. Since Z cannot transfer its excitation energy to chlorophyll, Z inthe excited state decays to its ground state by losing itsexcitation energy as heat. The molecular switch modelassumes that zeaxanthin (Z) absorbs light energy directlyand that NPQ is a consequence of the decay of excitedstate Z to its ground state with the concomitant loss ofheat. Thus, zeaxanthin decreases the efficiency of energytransfer to PSII reaction centers by acting as a directquencher of energy. Support for the molecular switchmodel is still equivocal since the presence of PsbS stillhas not been detected in the crystal structure of PSII,and furthermore, NPQ can occur in the absence of Z.

An alternative model for the regulation of NPQsuggests that the conversion of V to Z by the xantho-phyll cycle regulates the aggregation state of LCHII.In the aggregated state, the efficiency of energy trans-fer from LHCII to PSII reaction centers is drasticallydecreased. In this model, aggregated LHCII acts as theenergy quencher which protects PSII reaction centersfrom excess excitation. This suggests that Z is indirectlyinvolved in quenching energy by affecting the physicalstructure of LHCII. Regardless of which model is cor-rect, it appears that the xanthophyll cycle is a ubiquitousprocess for protecting the chloroplast against poten-tially damaging effects of excess light. However, it isinteresting to note that the xanthophyll cycle is absentin cyanobacteria. The regulation of NPQ by the xan-thophyll cycle continues to be an intensive and excitingarea of photosynthesis research.

What are the functional consequences of a stim-ulation of NPQ by high light? Irrespective of themechanism underlying photoprotection through NPQ,the functional consequence of NPQ is a decrease inthe efficiency of energy transfer to PSII reaction cen-tres. Because less energy is transferred to PSII reactioncenters per photon absorbed by LCHII under suchconditions, the efficiency of PSII photochemistry (P680+ energy → P680+ + e), will decrease per photonabsorbed. Thus, the light-inducible xanthophyll cycleprotects PSII reaction centers by dissipating excess exci-tation nonphotochemically as heat.

When plants are subjected to excess light,photoinhibition will occur (Chapter 13) which can

be measured as a decrease in Fv/Fm as a functionof time under high light. However, concomitantly,exposure to light also stimulates the de-epoxidationof the xanthophyll cycle (DEPS) and NPQ overtime (Figure 14.5A, left panel). However, when theplants are allowed to recover from the photoinhibitionby removal from the high light condition, Fv/Fmand DEPS rapidly recover to their original valuesprior to the photoinhibition treatment (Figure 14.5A,right panel). Thus, the responses of Fv/Fm

500

Irradiance (μmol m−2 s−1)

2500 750

30

20

Rat

e of

Pho

tosy

nthe

sis

(μm

ol O

2 e

volv

ed m

−2 s

−1)

10

PhotoinhibitionTime (min)

RecoveryTime (min)

40 800 40 80

0.8

0.6

0.4

0.2Fv/F

m (

, )

DE

PS

(,

)

A.

B.

FIGURE 14.5 A schematic graph illustrating photopro-tection and dynamic photoinhibition. (A) The effects oftime of exposure to photoinhibition and time of recov-ery from this photoinhibition on Fv/Fm (black curve) andDEPS (red curve). The panel on the left illustrates thatthe time-dependent decrease in Fv/Fm is almost a mir-ror image of time-dependent increase in DEPS due toexposure to high light. The panel on the right illustratesthat the time-dependent recovery of Fv/Fm (black bro-ken line) is almost a mirror image of the time-dependentdecrease in DEPS (red broken line). Such mirror imageresponses are characteristic of dynamic photoinhibition.In contrast to dynamic photoinhibition, recovery fromchronic photoinhibition is much slower (blue brokenline). (Adapted from Pocock, T., D. Koziak, D., Rosso,N. P. A. Huner. 2007. Journal of Phycology 43:924–936.)(B) The effects of exposure to either dynamic (redline) or chronic photoinhibition (broken red line) onthe light response curves for O2 evolution. The blackline represents the light response curve for control,non-photoinhibited plants. (Adapted from Osmond, C.B. 1994. In: N. R. Baker, J. R. Bowyer (eds.), Photoinhi-bition of Photosynthesis: From Molecular Mechanisms to theField, pp. 1–24. Oxford: Bios Scientific Publishers.)

14.2 Acclimation is Initiated by Rapid, Short-Term Responses 247

and DEPS to photoinhibition and recovery timeappear to be mirror images of one another. Therapidly reversible inhibition of PSII reaction centersthat is usually a consequence of an increase in thermalenergy dissipation through NPQ is defined as dynamicphotoinhibition. This reflects photoprotection. Incontrast, photodamage or chronic photoinhibitionis defined as the slowly reversible inhibition of PSIIreaction centers that is usually a consequence of damageto the D1 reaction center polypeptide. Thus, the timerequired for the recovery of Fv/Fm from chronicphotoinhibition is much longer than that from dynamicphotoinhibition. Photodamage is only slowly reversiblebecause of its dependence on protein synthesis for theD1 repair cycle (Chapter 13). The difference betweendynamic and chronic photoinhibition can also be seenat the level of O2 evolution (Figure 14.5B). Dynamicphotoinhibition usually results in a rapidly reversibledecrease in photosynthetic efficiency (Chapter 13)measured as the maximum slope of the CO2 lightresponse curve but not a decrease in photosyntheticcapacity measured as the maximum light saturated rateof photosynthesis. In contrast, chronic photoinhibitionusually results in a decrease in both photosyntheticefficiency and photosynthetic capacity which recoververy slowly (Figure 14.5B). Clearly, both photodamageand photoprotection may lead to photoinhibitionof photosynthesis as reflected in a decrease inphotosynthetic efficiency, albeit for different reasons.The former causes a reduction in photosyntheticefficiency due to an alteration in the antenna thatreduces the efficiency of resonance energy transfer tothe PSII reaction center. The latter is due to damage tothe reaction center that reduces the efficiency of chargeseparation rather than energy transfer in the antenna.

14.2.3 OSMOTIC ADJUSTMENT ISA RESPONSE TO WATER STRESS

A pronounced response to water stress in many plantsis a decrease in osmotic potential resulting from anaccumulation of solutes (see Chapter 1 and 2). Thisprocess is known as osmotic adjustment. While someincrease in solute concentration is expected as a resultof dehydration and decreasing cell volume, osmotic

adjustment refers specifically to a net increase in soluteconcentration due to metabolic processes triggered bystress. Osmotic adjustment generates a more negativeleaf water potential, thereby helping to maintain watermovement into the leaf and, consequently, leaf turgor.

Solutes accumulate during osmotic adjustment andthe decreases in osmotic potential due to osmoticadjustment are relatively small, less than 1.0 MPa. Nev-ertheless, the role of solutes in maintaining turgor atrelatively low water potentials represents a significantform of acclimation to water stress. Osmotic adjustmentmay also play an important role in helping partiallywilted leaves to regain turgor once the water supplyrecovers. By helping to maintain leaf turgor, osmoticadjustment also enables plants to keep their stomataopen and continue taking up CO2 for photosynthe-sis under conditions of moderate water stress. Solutesimplicated in osmotic adjustment include a range ofinorganic ions (especially K+), sugars, and amino acids(Figure 14.6). One amino acid that appears to be partic-ularly sensitive to stress is proline. A large number ofplants synthesize proline from glutamine in the leaves.The role of proline is demonstrated by experimentswith tomato cells in culture. Cells subjected to water(osmotic) stress by exposure to hyperosmotic concen-trations of polyethylene glycol (PEG) responded with aninitial loss of turgor and rapid accumulation of proline.As proline accumulation continued, however, turgorgradually recovered. Sorbitol, a sugar alcohol, andbetaine (N,N,N-trimethyl glycine) are other commonaccumulated solutes. Most chemicals associated withosmotic adjustment share the property that they do notsignificantly interfere with normal metabolic processes.Such chemicals are called compatible solutes.

Although osmotic adjustment appears to be a gen-eral response to water stress, not all species are capableof adjusting their solute concentrations. Sugarbeet (Betavulgaris), on the one hand, synthesizes large quantities ofbetaine and is known as an osmotic adjuster. Osmoticadjustment in cowpea (Vigna unguiculata), on the otherhand, is minimal and cowpea is known as an osmoticnonadjuster. Cowpea instead has very sensitive stomataand avoids desiccation by closing the stomata and main-taining a relatively high water potential. It is interesting

Proline

H2COH

H2COH

HO CH

HC OH

HC OH

HC OH

Sorbitol

N CH2 COOHH3CH3CH3C

Glycine betaine

H2C

NH

H2C CH COOH

CH2

+

FIGURE 14.6 Three solutes typicallyinvolved in osmotic adjustment.

248 Chapter 14 / Acclimation to Environmental Stress

to note that there is no long-term advantage of osmoticadjustment over stomatal closure, at least with regardto net carbon gain. While sugarbeet is able to continuephotosynthesis at lower water potentials, the advantageover cowpea is shortlived. After one or two days, exces-sive water loss overrides osmotic adjustment and overthe long term carbon assimilation in sugarbeet declines.

14.2.4 LOW TEMPERATURES INDUCELIPID UNSATURATION ANDCOLD REGULATED GENESIN COLD TOLERANT PLANTS

The study of cold tolerant, herbaceous plants such aswheat (Triticum aestivum), barley (Hordeum vulgare),alfalfa (Medicago), spinach (Spinacea oleracea), and themodel plant species Arabidopsis thaliana has enhancedour understanding of the metabolic and molecular eventsbefore, during, and after acclimation. This has assistedgreatly in the search for metabolic and genetic fac-tors involved in cold tolerance. One of the immediateresponses of cold tolerant plants to low temperature isan increase in the proportion of unsaturated fatty acids(Chapter 13) bound to lipids associated with the plasmamembrane, mitochondrial membranes as well as thy-lakoid membranes. Various biophysical measurementsindicate that this ensures that the membrane can remainin a more fluid and less gel-like state at lower temper-ature which enhances membrane stability and functionat these low temperatures. A change in the membranefrom the fluid state to a more solid state is marked byan abrupt change in the membrane activity. The tem-perature at which this transition occurs is known as thetransition temperature. This means that at tempera-tures above the transition temperature, the membraneremains fluid but becomes more solid or gel-like attemperatures below the transition temperature. Thisallows higher activity of membrane process at lowertemperatures.

Cold acclimation of herbaceous plants induceschanges in gene expression. During acclimation, thereare changes in mRNA transcription, increases inprotein synthesis, and qualitative changes in the patternof proteins synthesized. A major class of cold-inducedgenes encode homologs of late embryogenesis activeproteins (LEA-proteins) that are synthesized late inembryogenesis and during dehydration stress. Thesepolypeptides fall into a number of families based onamino acid sequence similarities (Table 14.1). However,these proteins encoded by cold-regulated genes sharecommon physical properties. (1) They are unusuallyhydrophilic. (2) They remain soluble upon boilingin dilute aqueous buffer. (3) They exhibit relativelysimple amino acid sequences that form amphipathicα-helices. It appears that cor15a and wsc120 interact

TABLE 14.1 Plant cold-regulated genes.

Plant Source Gene Polypeptide Mol. Mass (kD)

Arabidopsis cor15a cor15a 15Alfalfa cas15 cas15 15Wheat wcs120 wcs120 39Barley hva1 hva1 22

with membranes, which enhances their stability to lowtemperature as well as against freezing.

The promoter regions of certain cold-regulatedgenes are activated in response to low temperature anddehydration stress. Analyses of these promoter regionsof cold-regulated genes of Arabidopsis thaliana led to theidentification of a DNA regulatory element called thedehydration responsive element (DRE). The DRE hasa conserved core C-repeat sequence of CCGAC thatimparts responsiveness to low temperature and dehy-dration. Specfic proteins that bind to the DRE arecalled C-repeat binding f actors (CBFs). Thus, CBFsare transcriptional activators that are involved in reg-ulating the expression of cold-regulated genes. It isconcluded that cold acclimation is regulated by a familyof DRE-containing genes that are, in turn, induced by afamily of CBF transcriptional factors.

There is good reason to believe that ABA might beinvolved in cold acclimation of herbaceous tissues. Anincrease in endogenous ABA levels has been observedduring cold acclimation in several species and theamount of increase is greater in cold-tolerant varietiesthan in cold-sensitive varieties. In addition, significantlevels of cold tolerance can be induced by the applicationof ABA to intact plants, callus, and suspension cultures.In both intact alfalfa seedlings and suspension-culturedcells of winter rapeseed (Brassica napus), exogenous ABAcan induce up to 50 to 60 percent survival comparedwith a normal cold-acclimation treatment. ABA alsoinduces the synthesis of new proteins. In alfalfa, someof the induced proteins are unique to ABA treatment,but some are common to both low-temperature andABA treatment. Further research is required to eluci-date the precise role of ABA in cold acclimation and theinduction of genes associated with cold tolerance.

14.2.5 Q10 FOR PLANT RESPIRATIONVARIES AS A FUNCTIONOF TEMPERATURE

Enzymes and enzyme reactions are sensitive to temper-ature (Chapter 8, Box 1). Enzyme reactions typically areconsidered to have a Q10 of about 2, which means thatthe rate of the reaction doubles for each 10◦C rise intemperature. The rate of reaction increases with tem-perature until an optimum is reached, beyond which

14.3 Long-Term Acclimation Alters Phenotype 249

20

Temperature (�C)

100 30 40

4.0

3.0

2.0

Q1

0 f

or R

espi

rati

on

1.0

FIGURE 14.7 The temperature dependence of the Q10 forplant respiration. The broken line indicates the expectedrelationship between Q10 and temperature whereas thesolid line represents the actual relationship for plant res-piration. (Adapted from Atkin, O. K., M. G. Tjoelker.2003. Trends in Plant Science 8:343–351.)

the rate usually declines sharply. The decline in enzymeactivity is normally caused by thermal denaturation asa result of protein unfolding. It is usually assumed thatQ10 is independent of the temperature range over whichit is measured. However, this is not true for plant respira-tion. In fact, the Q10 increases linearly upon short-termincreases in temperature from 40◦ to 0◦C (Figure 14.7).This dynamic temperature response of Q10 for respira-tion appears to be consistent across diverse plant taxa.Since Q10 measures the temperature-dependent changein respiration rate, an increase in ambient temperaturewill cause a greater change in rates of respiration inplants native to cold, Arctic climates than plants nativeto hotter climates. Further, other abiotic factors such asirradiance and water deficit can also influence the Q10for plant respiration.

Why does the Q10 for plant respiration vary as afunction of short-term exposure to temperature? Thismay be explained on the basis of two primary factors: (1)the effect of temperature on the Vmax (Chapter 8, Box1) of enzymes involved in respiratory carbon metabolismas well as on the maximum rate of respiratory electrontransport (Chapter 10); (2) the effect of temperature onsubstrate availability. Short-term exposures to low tem-perature reduces the flux of carbon through glycolysisand the TCA cycle because low temperature will reducethe activities of the various enzymes involved in thesepathways. In addition, low temperature will decreasethe fluidity of the inner membrane, which decrease therate of respiratory electron transport. As a consequence,short-term exposure to low temperature will reduce therates of CO2 evolution and O2 consumption. However,at moderate to high temperatures, it is not enzymeactivity that limits the rate of reaction but rather theavailability of substrates such as ADP and ATP. At high

temperatures, mitochondrial membranes may becomeleaky to protons, and therefore, reduce the capacity tosynthesize ATP by chemiosmosis (Chapter 5 and 10).

14.3 LONG-TERM ACCLIMATIONALTERS PHENOTYPE

In this section, we will discuss specific examples ofslower, long-term responses to changes in light, wateravailability and temperature that are part of the accli-mation process that result in phenotypic alterations.

14.3.1 LIGHT REGULATES NUCLEARGENE EXPRESSION ANDPHOTOACCLIMATION

The process whereby adjustments are made to the struc-ture and function of the photosynthetic apparatus inresponse changes in growth irradiance is called pho-toacclimation. One consequence of photoacclimationis a change in pigment composition which results inan altered visible phenotype. It is important to notethat photoacclimation requires growth and develop-ment. For example, photoautrophs grown under highlight typically exhibit as decrease in total chlorophyllper leaf area compared to the same plants grown atlow irradiance. Thus, the leaves of high-light plants areusually a pale green or yellow-green compared to a darkgreen phenotype of the same species grown at low light.Functionally, high-light plants exhibit a photosyntheticlight response curve for CO2 assimilation that is distinctfrom that of plants grown under low light, whether mea-sured as net photosynthesis (Figure 14.8A) or as grossphotosynthesis (Figure 14.8 B). Typically, plants grownunder high light have a higher photosynthetic capacity,that is, a higher light saturated rate of photosynthesisthan low-light plants. In contrast, high-light plants mayhave a lower photosynthetic efficiency, that is, a lowerinitial slope, compared to the same plants grown at lowlight (Figure 14.8B).

Many green algae may exhibit an even moredramatic change in phenotype in response to growthat either high or low light than terrestrial plants(Figure 14.9A). The high light phenotype illustratesone mechanism of photoacclimation which involvesthe modulation of the size and composition of thelight-harvesting complex (LHCII) of PSII coupled witha change in Rubisco content (see Chapter 7). There isnow a consensus that the content of LHCII decreaseson a leaf area basis as the growth irradiance increases(Figure 14.9B), which is coupled with increased xan-thophyll cycle activity and increased photoprotectionthrough NPQ. Since the bulk of the chlorophyll aand chlorophyll b is bound to LHCII (Chapter 7), adecrease in the amount of LHCII results in a decrease

250 Chapter 14 / Acclimation to Environmental Stress

50 10 15

Irradiance(mol m−2 day−1)

Gro

ss P

hoto

synt

hesi

s(m

ol C

o 2 m

−2 d

ay−1

)N

et P

hoto

synt

hesi

s(m

ol C

o 2 m

−2 d

ay−1

)

20

HL

LL

25

0.3

0.2

0.1

0

0.2

0.1

HL

A

B

LL

FIGURE 14.8 Photosynthetic light response curves forAlocasia macrorrhiza. This C3 plant was grown undereither high light (25 mol m−2 day−1) or low light (1.7 molm−2 day−1). (A) Light response curves for net CO2 gasexchange. Recall (Chapter 8) that the apparent or net rateof CO2 uptake by a leaf (AP) = the gross rate or actualphotosynthetic rate (GP)—[rate of respiration (R) + therate of photorespiration (PR)]. In the dark, GP = 0 andPR = 0. In the dark gas exchange is due to respirationand measured as CO2 evolution. Thus, in (A) the valuefor net photosynthetic rate in the dark (irradiance = 0)is a negative number indicating net CO2 evolution fromthe leaf. At higher irradiance, the rate of photosyntheticCO2 uptake exceeds both respiration and photorespi-ration such that there is a net uptake of CO2. Rates oflight-dependent CO2 uptake are arbitrarily given positivevalues whereas rates of CO2 evolution are given negativevalues. Thus, gross photosynthesis (GP) represents pho-tosynthetic CO2 uptake rates that have been correctedfor rates of respiratory CO2 evolution, that is, GP = AP+ R +PR. In (B), gross photosynthetic rates were calcu-lated assuming minimal contributions from photorespi-ration. (Adapted from Pearcy, R. W. 1996. Photosynthesis:A Comprehensive Treatise, A. S. Raghavendra (ed.), pp.250–263. Cambridge: Cambridge University Press.)

in total chlorophyll content as well as an increase in theratio of chlorophyll a/chlorophyll b. Functionally, thisresults in a decrease in light-harvesting efficiency underlow, light-limiting irradiance which may be detectedas a decrease in the initial slope of the photosyntheticlight response curve in high-light plants compared tolow-light plants (Figure 14.8B). Under light-saturatedconditions, the activity of the Calvin Cycle limits therate of photosynthesis (Chapter 13). Thus, high-lightplants exhibit higher photosynthetic capacity becausethey exhibit a higher total Rubisco content per leafarea than the same plants grown under low light.

Furthemore, the rate of growth of plants is usuallyfaster under high-light than low-light conditions. Thisis reflected in higher rates of respiration in high-lightplants than low-light plants. This is indicated by thefact that when measuring net CO2 assimilation in thedark (Figure 14.8A, 0 irradiance), the rate of CO2evolution due to respiration is greater in the high-lightthan low-light plants.

Experiments utilizing single cell green algal speciessuch as Dunaliella tertiolecta and Chlorella vulgarisindicate that the light-dependent change in the contentof LHCII is modulated in response to the redox state ofthe plastoquinone (PQ) pool (see Chapter 7). Photosyn-thetic electron transport can be inhibited specifically atthe Cyt b6/f complex with a compound called DBMIB(2,5-dibromo-6-isopropyl-3-methyl-1,4-benzoquin-one) (Figure 14.9C). In the presence of this compound,there is a net accumulation of PQH2 because, althoughPSII is able to convert PQ to PQH2, PSI can notoxidize this pool because of the chemical block at theCyt b6/f complex (Figure 14.9C). Thus, under theseconditions, the PQ pool remains largely reduced andtranscription of the nuclear Lhcb genes coding for themajor LHCII polypeptides is repressed. This results inan inhibition of the biosynthesis of LHCII polypeptides,which decreases the LHCII polypeptide content. As aconsequence, this results in a yellow phenotype typicalof high-light grown algal cells (Figure 14.9A, compareDBMIB and HL). Alternatively, photosyntheticelectron transport can also be inhibited specificallyat PSII (Figure 14.9C) in the presence of DCMU(3-(3,4-dichlorophenyl)-1,1-dimethylurea; see Figure7.13). Under these conditions, PSII is unable toreduce PQ to PQH2 and PQ accumulates becauseany PQH2 pool is oxidized by PSI. This produces agreen phenotype (Figure 14.9, DCMU) which mimicslow-light-grown cells (Figure 14.9A, LL).

How can we rationalize this phenotypic response togrowth under high light? Exposure to high light poten-tially exposes plants to an imbalance in energy budget.This energy imbalance is a consequence of the fact thatmore light is absorbed than can be utilized metabolicallyeither through the reduction of CO2 and NO−

3 or bythe oxidation of carbon by respiration (Chapter 8, 10,and 11). Under such conditions, the PQ pool tends to bein a reduced state. Continued exposure to such condi-tions can lead to chronic photoinhibition (Chapter 13)which, if it persists over the long-term, may lead tocell death. Energy balance can be attained under highlight by reducing the efficiency of light absorptionunder high light. This is accomplished by producinga smaller LHCII (Figure 14.9B) with high concen-trations of zeaxanthin which decreases the absorptioncross section (σ ) of PSII and decreases the probabil-ity of light absorption by PSII. One consequence of

14.3 Long-Term Acclimation Alters Phenotype 251

LL HL DCMU DBMIB

RC RC

Low light

A.

B.

C.

High light

DCMU DBMIB

hν

PSII

PQH2

PQ(ox) (red) (ox)

(red) (ox) (red)

Cytb6/f

Cytb6/f

PC

PChν

PSI

LHCII

COREANTENNA

FIGURE 14.9 Photoacclimation to high light. (A)An illustration of cell cultures of the green alga,Dunaliella tertiolecta grown under either low light(LL) or high light (HL) and either in the presenceof DBMIB or DCMU. (B) A model to illustrate thedifference in LHCII between an organism grownat low light and the same organism grown at highlight. RC represents the reaction centre complex ofPSII. Photoacclimation is thought to be a result ofchanges in LHCII content with little or no change ineither the content of the core antenna (Chapter 7;CP47 and CP43) or the reaction center complex(D1 and D2, Chapter 7). (C) A simplified model ofphotosynthetic linear electron transport illustratingthe coupled reduction and oxidation of PSII, the PQpool, the Cyt b6/f complex, plastocyanin (PC) andPSI. (See Chapter 7 for details.) DCMU inhibits PSIIactivity. DBMIB inhibits the Cyt b6/f complex.

this decreased efficiency for light absorption and energytransfer is enhanced photoprotection and increased tol-erance to growth conditions that typically cause photo-inhibition.

An energy imbalance due to high light appears tobe sensed through changes in the redox state of the PQpool. Such an energy imbalance due to overexcitationof PSII is called excitation pressure. Thus, theredox state of the PQ pool appears to induce a signaltransduction pathway which regulates the expression ofthe Lhcb genes present in the nucleus. The regulationof nuclear genes by organelles such as the chloroplastis called retrograde gene regulation (Figure 14.10).However, retrograde regulation is not restricted tochloroplast-nucleus interactions. Like the chloroplast,the mitochondrion also regulates nuclear geneexpression. This illustrates that important molecularcommunication pathways exist between the chloroplast,the mitochondrion, and the nucleus to regulate nucleargene expression. The chloroplast-nucleus and the

mitochondrion-nucleus signal transduction pathway(s)remain elusive but an intensive area of research.

The maintenance of cellular energy balanceis called photostasis, which is dependent uponchloroplast-mitochondrial interactions. For example,the mitochondrial Moc1 protein is thought to regulatethe transcription of mitochondrial genes involvedin the maintenance of the mitochondrial respiratoryelectron transport. However, under high light, themoc1 mutant, which lacks this mitochondrial protein,is unable to up-regulate rates of respiration to matchthe production of fixed carbon by photosynthesis.The block in mitochondrial electron transport slowsthe rate of respiratory carbon metabolism which,in turn, causes a feedback inhibition in the rate ofphotosynthetic electron transport. This also results inthe reduction of the PQ pool in the chloroplast. Thisis an excellent example of the link between chloroplastand mitochondrial metabolism and its importance inthe regulation of cellular redox balance.

252 Chapter 14 / Acclimation to Environmental Stress

FIGURE 14.10 A model illustrating retrograde reg-ulation. The redox state of the plastoquinonepool (PQ) in the chloroplast can regulate nucleargene expression. Similarly, the redox state of themitochondrial ubiquinone pool (UQ) can regu-late nuclear gene expression. The regulation ofnuclear gene expression by the chloroplast andthe mitochondrion is called retrograde regulation.The chloroplast and the mitochondrion commu-nicate through carbon metabolic pathways.

NuclearGene

Expression

Chloroplast Mitochondrion

PSII

PQH2

PQ

PSI Complex I

ComplexIV

UQH2

UQ

Nucleus

hν hν

14.3.2 DOES THE PHOTOSYNTHETICAPPARATUS RESPOND TOCHANGES IN LIGHT QUALITY?

Terrestrial plants growing in extremely shaded habitatsbelow the canopy of a tropical rainforest floor receiveabout 1 percent or less of the photosynthetically activeradiation (PAR) (Chapter 6) incident at the canopy level.Furthermore, the light that reaches the forest floor isenriched in far-red and green light because the red andblue light are absorbed by the canopy leaves of the tallertrees. Thus, acclimation to natural shade conditionswould appear to be a complex interaction of responsesto both light intensity and light quality. Acclimationin response to light quality would involve regulationby the plant photoreceptor, phytochrome (Chapter 22).A major problem in past experiments to test the dif-ferential effects of light quality versus light intensityon acclimation of the photosynthetic apparatus is thefact that differences in light quality were not applied atequal photon fluence rates (Chapter 6). Thus, this makesinterpretations of many of these experiments equivocal.Experiments with pea and corn where equal photon flu-ence rates of red and far-red light were used, the ratioof chlorophyll a : chlorophyll b did not change signifi-cantly. This is consistent with the observations no majorchanges in thylakoid membrane structure and compo-sition were observed in pea and corn plants exposed todifferences in light quality of equal photon fluence rates.However, the ability to adjust the structure and com-position of the photosynthetic apparatus in responseto light quality appears to be strongly species depen-dent. When the shade fern, Asplenium australasicum, was

grown in red light, changes in leaf pigment compositionmimicked acclimation to high light whereas growth ofthis species under blue light mimicked acclimation tolow light. These results are consistent with the fact thatshade plants grow in habitats enriched in far-red andgreen light but depleted in the blue and red regions ofthe visible spectrum. Furthermore, very little is knownregarding the contents of phytochrome and other pho-toreceptors in shade adapted plants. Thus, it appearsthat acclimation of the photosynthetic apparatus to lightquality is highly species dependent.

A dramatic example of acclimation to light qual-ity is exhibited by cyanobacteria (Figure 14.11). Recallthat the photosynthetic apparatus of these prokaryoticphotoautrophs exhibit similarities as well as impor-tant differences from that of their eukaryotic pho-toautotrophic counterparts (Chapter 7). Like plantsand green algae, cyanobacteria contain both photo-system I (PSI) and photosystem II (PSII) and, as a con-sequence, are oxygenic. However, unlike these eukary-otes, cyanobacteria use extrinsic, thylakoid membrane,pigment-protein complexes called phycobilisomes toharvest light energy for photosynthesis. The major pig-ments of many cyanobacteria covalently bound to thephycobilisomes include phycocyanin (PC) and phyco-erythrin (PE). Cyanobacteria that accumulate PC andPE such as Fremyella diplosiphon are capable of adjustingthe pigment composition of phycobilisomes in responseto changes in light quality. This ability to adjust toambient light color is called complementary chro-matic adaptation (CCA). CCA is a consequence of achange in the proportion of PC and PE in response tolight quality (Figure 14.11). When exposed to growth

14.3 Long-Term Acclimation Alters Phenotype 253

A. B.

RED

GREEN

FIGURE 14.11 The effect of light quality on the phenotypes of Fremyella diplosiphonThe filamentous cyanobacterium, F. diplosiphon, grown on agar plates and acclimatedto either green light (A) or to red light (B). Below each culture is a model illustratingthe change in pigment composition of the phycobilisome associated with PSII inresponse to growth under either green or red light. (Adapted from Kehoe, D. M.,A. Gutu. 2006. Responding to color: The regulation of complementary chromaticadaptation. Annual Review of Plant Biology 57:127–150 (with permission)).

under red light, Fremyella diplosiphon exhibits the char-acteristic blue-green phenotype due to the fact that PCmost effectively absorbs red light. In contrast, duringgrowth under green light, Fremyella diplosiphon exhibitsa distinctive red phenotype because PE most effec-tively absorbs green light. Since this acclimation processis photoreversible, it appeared to share features thatwere common to the red/far red photoreversible, phy-tochrome response characteristic of terrestrial plants.However, recently it has been shown that CCA is regu-lated by both the redox state of the PQ pool as well asby multiple, as yet unidentified, photoreceptors.

Clearly, CCA is due to changes in the expression ofgenes present in the genome of a cyanobacterium to pro-duce the different phenotypes. The reversible changesin phenotype associated with CCA reflect a remarkableplasticity of cyanobacteria to respond to changes in ambi-ent light color to maximize absorption of light energyfor photosynthesis and growth. Thus, this phenomenonshould be called complementary chromatic acclimationrather than complementary chromatic adaptation.

14.3.3 ACCLIMATION TO DROUGHTAFFECTS SHOOT–ROOT RATIOAND LEAF AREA

One of the long-term effects of water deficit is a reduc-tion in vegetative growth. Shoot growth, and especiallythe growth of leaves, is generally more sensitive thanroot growth. In a study in which water was withheldfrom maize (Zea mays) plants, for example, there wasa significant reduction of leaf expansion when tissuewater potentials reached −0.45 MPa and growth wascompletely inhibited at −1.00 MPa. At the same time,

normal root growth was maintained until the waterpotential of the root tissues reached −0.85 MPa andwas not completely inhibited until the water potentialdropped to −1.4 MPa. Reduced leaf expansion is bene-ficial to a plant under conditions of water stress becauseit leads to a smaller leaf area and reduced transpiration.Traditionally, the effect of low water potential on cellenlargement has been attributed to a loss of turgor inthe cells in the growing region. Plant cell enlargementoccurs when water moves in to establish full turgorfollowing stress relaxation in the cell wall (Chapter 16and 17). It should not be too surprising, then, that anearly consequence of limited water supply would bereduced growth. Thus, although the cells are able tomaintain turgor, it does not appear to be sufficient tomaintain a full rate of growth based on cell enlargement.

The preceding discussion applies primarily toshoots and leaves that are actively growing. Manymature plants, such as cotton (Gossypium hirsutum),subjected to prolonged water stress will respond byaccelerated senescence and abscission of the olderleaves. In the case of cotton, only the youngest leavesat the apex of the stem will remain in cases of severewater stress. This process, sometimes referred to as leafarea adjustment, is another mechanism for reducingleaf area and transpiration during times of limited wateravailability. So long as the buds remain viable, newleaves will be produced when the stress is relieved.

As noted above, roots are generally less sensitivethan shoots to water stress. Apparently, osmotic adjust-ment in roots is sufficient to maintain water uptakeand growth down to much lower water potentials thanis possible in leaves. Relative root growth may actu-ally be enhanced by low water potentials, such that the

254 Chapter 14 / Acclimation to Environmental Stress

root–shoot ratio will change in favor of the propor-tion of roots. An increase in the root–shoot ratio asthe water supply becomes depleted is clearly advanta-geous, as it improves the capacity of the root system toextract more water by exploring larger volumes of soil. Achanging root–shoot ratio is accompanied by a changein source–sink relationships with the result that a largerproportion of photosynthate is partitioned to the roots.Delivery of carbon to the roots can continue, however,only to the extent that carbon supply can be maintainedby photosynthesis or mobilization of reserves stored inthe leaves. In addition, to changes in leaf area, someplants respond to growth under water deficit conditionsby reducing stomatal frequency. By reducing the num-ber of stomates per leaf area, a plant can reduce potentialwater loss due to transpiration when water is limitinggrowth.

An early response to water deficits is closure ofstomates to conserve water. In some plants this maylead to low internal leaf CO2 concentrations which willlimit photosynthetic capacity. A limited photosyntheticcapacity in the presence of light can result in exposureof plants to excess light and photoinhibition. Sinceleaf O2 levels will remain higher than CO2 when leafstomates are closed, O2 will be preferentially consumedthrough photorespiration and the action of the enzymeRubisco (Equation 14.1). Just as for the reduction ofCO2 (Equation 14.1), the continuous consumption ofO2 by Rubisco (Equation 14.2)

RuBP + CO2 → 2PGA (14.1)

RuBP + O2 → PGA + P-Glycolate (14.2)

requires the continuous regeneration of RuBP by theCalvin Cycle (Chapter 8). This requires a constant sup-ply of NADPH and ATP generated by photosyntheticelectron transport (see Figure 13.4). The fixation of onemole of O2 through photorespiration consumes moreenergy (5ATP + 3NADPH) than the fixation of onemole of CO2 (3ATP + 2NADPH). Consequently, thephotorespiratory pathway may play an important rolein maintaining photostasis when CO2 is limiting. Sucha role for photorespiration is supported by the factthat photorespiratory mutants of Arabidopsis thaliana aremore sensitive to high light under water stress.

14.3.4 COLD ACCLIMATION MIMICSPHOTOACCLIMATION

For more than a century it has been known that thegrowth of winter varieties of cold tolerant herbaceousplant species such as rye (Secale cereale L. cv Mus-keteer) and wheat (Triticum aestivum L. cv Kharkov)at low temperatures results in enhanced freezing tol-erance measured as LT50, the freezing temperatureat which 50 percent of a population of plants are

killed (Figure 14.12A). The enhanced freezing tol-erance is strongly correlated with the expression ofcor genes. However, in addition to enhanced freezingtolerance, cold acclimated winter varieties of wheat,rye, barley, spinach as well as Arabidopsis thaliana alsoexhibit a decreased sensitivity to photoinhibition eventhough the plants were never exposed to high light(Figure 14.12B). Thus, the decreased sensitivity to pho-toinhibition exhibited by these cold acclimated plantsmimics photoacclimation to high light. However, in

20

40

60

80

100

−16−14−12−10−8−40

% P

lant

s ki

lled

Temperature (�c)

100

100

Fv/F

m (

% o

f co

ntro

l)

Photoinhibition Time (h)

A.

B.

FIGURE 14.12 (A) The schematic graph illustrating theeffect of growth temperature on freezing tolerance mea-sured as LT50. Cold-tolerant plants grown at 25◦C (blackline) exhibit an LT50 of −7◦C whereas the cold-tolerantplants that are grown at 5◦C (red line) exhibit an LT50of about −12◦C. Thus, cold-acclimated plants exhibit anincreased freezing tolerance. (B) A schematic graph illus-trating the effect of growth temperature on sensitivity tophotoinhibition. The maximal photochemical efficiencyof PSII measured as Fv/Fm of the plants grown at 25◦C(black line) decreases to a greater extent than that ofplants grown at 5◦C (red line). Thus, cold-acclimatedplants exhibit a decreased sensitivity to photoinhibition(Adapted from Gray, G. R., L. V. Savitch, A. G. Ivanov,N. P. A. Huner. 1996. Plant Physiology 110:61–71).

14.4 Freezing Tolerance in Herbaceous Species is a Complex Interaction Between Light and Low Temperature 255

these winter varieties, this is accomplished with minimalchanges in the structure and composition of LHCII.

How is this possible? Growth of cold-tolerantwinter wheat and winter rye at low temperature stim-ulates photosynthetic capacity with minimal changesin photosynthetic efficiency or in the ratios of chloro-phyll a/chlorophyll b. This stimulation in photosyn-thetic capacity is the result of the following. First, coldacclimation enhances the transcription and translationof genes encoding major regulatory enzymes of stro-mal and cytosolic carbon metabolism such as Rubisco,chloroplastic FBPase, cytosolic FBPase, and sucrose-Psynthase (SPS), as well as increased fructan biosynthesisin the vacuole (Chapter 9). This results in higher totalenzyme activity and a higher flux of carbon throughthe sucrose biosynthetic pathway. Second, this is cou-pled to higher rates of carbon export from the leavesin the light due to enhanced sink activity (Chapter 9).In contrast to short-term exposure to low temperature,growth at low temperature stimulates rates of respiratorycarbon metabolism. Third, cold acclimation suppressesphotorespiration which also enhances net carbon gain.Thus, subsequent exposure of these cold-acclimatedplants to increasing irradiance stimulates their photo-synthetic capacity even further, which is translated intoincreased growth rates and biomass production at lowtemperature in addition to the stimulation of NPQvia the xanthophyll cycle. Thus, these cold-acclimatedplants exhibit a resistance to photoinhibition becauseof their enhanced capacity to utilize the absorbed lightfor carbon metabolism, biomass production and growthcoupled with the dissipation of any excess absorbedlight through NPQ. It is important to appreciate that,to exhibit these characteristics, cold-tolerant plants mustgrow and develop at low temperatures.

Similar to cold-acclimated winter varieties ofwheat, rye, barley, spinach, as well as Arabidopsisthaliana, cold-acclimated Chlorella vulgaris also exhibitsa decreased sensitivity to photoinhibition. However, incontrast with these terrestrial plants, cold-acclimatedChlorella vulgaris exhibits the same yellow-pale-greenphenotype as high-light grown cells (Figure 14.9,HL) even though these cold-acclimated cells havenot grown under high light. It has been shownthat cold acclimation of Chlorella vulgaris mimicsphotoacclimation because growth at low temperatureinduces a comparable energy imbalance or excitationpressure as growth at high light. Why does this occur?Unlike wheat, rye, and Arabidopsis, Chlorella vulgarisis unable to up-regulate photosynthetic capacityduring cold acclimation when measured on a per-cellbasis. Low growth temperature reduces the rate ofintersystem electron transport as well as enzyme activityinvolved in carbon metabolism without affecting lightabsorption, and energy transfer from LHCII to P680and its subsequent photooxidation to P680+. Under

these conditions, PQH2 accumulates because PSIIreduces the PQ pool faster than PSI and ultimatelyCO2 assimilation can oxidize this pool. Concomitantly,this chloroplast redox signal represses Lhcb geneexpression by retrograde regulation (Figure 14.10).The cyanobacterium, Plectonema boryanum, also showsa similar phenotypic response to growth at either lowtemperature or high light as Chlorella vulgaris. However,the cyanobacterium responds by decreasing the size ofits phycobilisome associated with PSII (Chapter 7).

The discussion above illustrates the remarkableplasticity with which photoautrophs respond to energyimbalances as a consequence of growth at either highlight or low temperature. Although low temperatureand high light cause a similar imbalance in cellularenergy budget, the response of photoautrophs to energyimbalance or excitation pressure is species dependent.It appears that many terrestrial cold-tolerant plants canattain photostasis by combining an up-regulation of pho-tosynthetic capacity and increased growth rates to utilizeabsorbed light energy with photoprotection throughxanthophyll cycle activity. In contrast, many green algaeattain photostasis by reducing light-harvesting efficiencyby making a smaller light-harvesting complex throughretrograde regulation due to their inability to utilizethe absorbed energy through carbon metabolism andgrowth.

14.4 FREEZING TOLERANCE INHERBACEOUS SPECIES IS ACOMPLEX INTERACTIONBETWEEN LIGHT AND LOWTEMPERATURE

Cold acclimation and the development of maximumfreezing tolerance in overwintering herbaceous plantssuch as winter wheat, winter rye, spinach, and Arabidopsisthaliana requires active growth and development at lowtemperature. As a result, leaves of these plant speciesdeveloped at low temperature are anatomically, mor-phologically, physiologically, and biochemically distinctfrom the same plants developed at warm temperatures.For example, these herbaceous species grown at low tem-perature exhibit a short, compact growth habit, thickerleaves due to an increase in leaf mesophyll cell sizeand/or an increase in the number of palisade cell lay-ers, and an increase in cell cytoplasm associated witha decrease in leaf water content. Photosynthesis con-tinues during the cold acclimation period and providesthe necessary energy required for this low-temperaturegrowth, which leads to the cold-acclimated state andmaximum freezing tolerance (Equation 14.3).

256 Chapter 14 / Acclimation to Environmental Stress

TABLE 14.2 Effects of growth temperature andgrowth irradiance on LT50 of winter rye (Secalecereale L. cv Musketeer). Growth irradiance is inunits of μmol photons m−2 s−1).

Growth GrowthTemperature (◦C) Irradiance LT50 (◦C)

20 50 low light −420 250 moderate light −620 800 high light −8

5 50 low light −85 250 moderate light −16

Cold Stress → Growth/Development

→ Cold Acclimation → Freezing Tolerance (14.3)

As indicated in Table 14.2, exposure to low tem-perature (5◦C) is an absolute requirement for the attain-ment of maximum freezing tolerance in winter rye(LT50 = −16◦C). However, low temperature by itselfis not sufficient to induce maximum freezing toler-ance since growth at low temperature but low lightreduces LT50 by 50% (Table 14.2). Thus, light is alsoan absolute requirement for the attainment of maxi-mum freezing tolerance in winter cereals (Table 14.2).This light-dependent increase in freezing tolerance atconstant low-growth temperature occurs independentlyof either photoperiod or light quality. Thus, the lightdependence of LT50 in herbaceous winter cereals doesnot appear to be phytochrome-dependent. Further-more, increasing the growth irradiance at temperaturesthat normally do not induce freezing tolerance (20◦C)results in a doubling of LT50 from −4 under low light to−8◦C under high light, which is consistent with the lightdependence of freezing tolerance. However, high lightcan not compensate for low temperature for the induc-tion of maximum freezing tolerance in winter cereals.Thus, the attainment of maximum freezing toleranceappears to be the result of an additive effect of bothlow-growth temperature and growth irradiance.

14.4.1 COLD ACCLIMATED PLANTSSECRETE ANTIFREEZEPROTEINS

Overwintering plants can tolerate freezing because oftheir ability to control the freezing event itself. As longas freezing of water is confined to the apoplast, thatis the cell wall and the extracellular space, the plantwill survive. Alternatively, if freezing occurs intracel-lularly, the plant will die. Cold acclimation in manyplants is associated with the secretion of antifreezeproteins (AFPs) from the cytoplasm into the apoplast.AFPs have been reported in ferns, gymnosperms, as

well as mono- and dicotyledonous angiosperms. AFPsinhibit ice crystal growth by binding to the surface ofa growing ice crystal via hydrogen bonding betweenspecific hydrophilic amino acids present in the AFP andwater within the crystal lattice of ice. The presenceof AFPs in cold-tolerant plants is not constitutive butrequires exposure to low temperature and they accumu-late in virtually all plant tissue including seeds, stems,leaves, flowers, and roots. When winter rye plants coldacclimate, the gaseous plant hormone, ethylene, is pro-duced. Ethylene induces the transcription of the familyof genes that encode AFPs. Upon translation, the AFPsare secreted via the endoplasmic reticulum, Golgi bod-ies, and vesicles that fuse with the plasma membraneand are deposited on the surface of the cell wall wherethey inhibit ice crystal formation.

Do AFPs alter LT50? The answer is an unequivocalno. If this is so, what is the role of AFPs? Although AFPshave a minimal effect on LT50, AFPs have a significanteffect on the rate of ice crystal formation. Thus, AFPsmost likely enhance winter survival by slowing the rateof extracellular freezing.

14.4.2 NORTH TEMPERATE WOODYPLANTS SURVIVE FREEZINGSTRESS

Boreal deciduous trees, conifers, and shrubs such aspaper birch (Betula papyrifera), trembling aspen (Popu-lus tremuloides), and willow (Salix sp.)—all found as farnorth as the arctic circle—survive because they are ableto acclimate to the below-freezing winter temperatures.During their normal growing season, these plants willsuffer injury or death if exposed to freezing tempera-tures. Even a light frost during the spring or summermay be lethal to plants that are actively growing. Yet,cold-acclimated stems of these species may survive tem-peratures as low as −196◦C (liquid nitrogen) withoutapparent injury.

Acclimation of woody species to freezing stressis a common phenomenon in nature, but the precisemechanism by which acclimation is achieved is notwell understood. It is known that acclimation in woodytissues occurs in two distinct stages (Figure 14.13). Itbegins in the autumn when growth and photosynthesisceases and the plant enters dormancy. This first stageof acclimation is induced by short days and is thoughtto be under the control of phytochrome. Acclimation atthis stage can be inhibited by long days and early frost.Thus, in woody species, it is essential that the plantenter the dormant stage prior to the onset of frost toprevent freezing damage.

The second stage of acclimation is triggered byexposure of the overwintering tissue to low tempera-ture, corresponding to the first frost (Figure 14.13). Atthis stage, respiratory activity is sufficient to provide the

14.5 Plants Adjust Photosynthetic Capacity in Response to High Temperature 257

Time (month of year)

Tiss

ue t

empe

ratu

re (

ºC)

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Lowesttemperaturetested

–60

–20

+20

Sept. Oct. Nov.

Short-dayphotoperiodresponse

FIGURE 14.13 Acclimation to low temperature in woodystems. The curve depicts the lowest survival tempera-ture as a function of time of year. Note that significantdecreases in survival temperature correspond to shorten-ing daylength and the time of the first frost.

energy necessary for the numerous metabolic changesrequired to attain the maximum cold-acclimated state.There are increases in the level of organic phosphatesand the conversion of starch to sugars. Glycopro-teins accumulate and the protoplasm becomes generallymore resistant to dehydration. Fully acclimated cellscan withstand temperatures far below those normallyexperienced in nature.

14.5 PLANTS ADJUSTPHOTOSYNTHETICCAPACITY IN RESPONSETO HIGH TEMPERATURE

Plants that can acclimate to high temperatures arecalled thermotolerant plants. Photosynthetic capacitymeasured as the maximum light-saturated rate of CO2assimilation is sensitive to temperature (Figure 14.14).The C3 and the C4 plant illustrated in Figure 14.14exhibit temperature optima for photosynthesis that aredependent upon the growth condition to which theseplants were exposed. Plants exposed to cool tempera-tures generally exhibit a lower temperature optimum forphotosynthesis than those exposed to high temperatures.Such a shift in temperature optima reflects photosyn-thetic acclimation to temperature which has been shown

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L. divaricata

Hot

T. oblongifolia

A

B

FIGURE 14.14 A schematic graph illustrating the abilityof thermotolerant C3 and C4 plants to adjust photosyn-thetic capacity. Temperature profiles for light-saturatedphotosynthetic rates are plotted as a function of leaftemperature for a C4 plant, T. oblongifolia (A), and aC3 plant, L. divaricata (B). (Adapted from Berry, J. A.,O. Bjorkman. 1980. Annual Review of Plant Physiology31:491–543.)

to be reversible and not due to temperature-dependentstomatal limitations. Rather, such photosynthetic accli-mation appears to be due to a combination of changesin the temperature stability of thylakoid membranes aswell as the enzymes of the PCR cycle such as Rubisco.Although photosynthetic acclimation of C3 overwin-tering species such as wheat and rye requires growthand development at cold temperatures, this is not thecase for all plants. For example, reversible photosyn-thetic acclimation in the C3 species, Nerium oleander,is observed in fully expanded, mature leaves exposedto either high or low temperatures. This may be due,in part, to the different absolute temperature ranges towhich a particular species can acclimate.

The maximum rate of photosynthesis of the C4plant, T. oblongifolia, appears to be more sensitive toleaf temperature than that of L. divaricata, a C3 plant(Figure 14.14). However, such differential responses donot reflect a general difference between C3 and C4species. Rather, such differences have been shown tobe very species dependent within plants that exhibiteither C3 or C4 photosynthesis. Clearly, plants exhibitextraordinary plasticity to adjust photosynthetically tochanges in temperature.

258 Chapter 14 / Acclimation to Environmental Stress

14.6 OXYGEN MAY PROTECTDURING ACCLIMATION TOVARIOUS STRESSES

Although the oxygen evolving complex (OEC) associ-ated with PSII results in the light-dependent evolutionof oxygen (Chapter 7), O2 can also act as alternativeelectron acceptor for photosynthetic electron transport.Thus, photosynthetic electron transport may also con-sume oxygen. Even under normal conditions, up to5 percent to 10 percent of the photosynthetic electronsthat are generated by PSI may react with molecularoxygen rather than with NADP+. This has importantfunctional consequences for active chloroplasts. Thephotoreduction of oxygen by PSI is called the Mehlerreaction and results in the production of another toxic,reactive oxygen species known as a superoxide radical(O−

2 ) (a radical is a molecule with an unpaired electron).To counteract the accumulation of this radical, photo-synthetic organisms have evolved mechanisms to protectthemselves from excess light and the potential ravagesof O2. An effective system for the removal of super-oxide is the ubiquitous enzyme superoxide dismutase(SOD). SOD is found in several cellular compartments

FIGURE 14.15 Oxygen as an alternativeelectron acceptor in chloroplasts. (A) TheAsada-Halliwell pathway. O2 can be pho-toreduced by PSI directly to generatethe superoxide free radical, O−

2 (Mehlerreaction). Superoxide dismutase (SOD)then converts this radical to hydrogenperoxide (H2O2). Hydrogen peroxide isalso toxic and is reduced via the chloro-plastic enzyme, ascorbate peroxidase, towater and ascorbate is oxidized to mon-odehydroascorbate (MDHA). Ascorbate(vitamin C) is regenerated through theaction of the enzyme, dehydroascorbatereductase, through the consumption ofreduced glutathione (GSH). Oxidized glu-tathione (GSSH) is, in turn, reduced by theenzyme glutathione reductase, which usesNADPH as reductant. (B) Chlororespira-tory pathway. NAD(P)H dehydrogenase(Ndh) present in thylakoid membranesconsumes stromal NAD(P)H and passesthe electrons (e) directly to plastoquinone(PQ). The plastid terminal oxidase (PTOX)present in thlylakoid membranes oxidizesplastoquinol and reduces O2 to water. Thestromal pool represents any metabolicpathway present in the stroma that gener-ates reducing power (see Chapter 8). Ndhmay also participate in cyclic electron trans-port around PSI.

Lumen

Ndh PQCyt

PSI

PC

fd

e e

ee

e

NAD(P)H

PQ

PTOX

Stomal Pool

PSI

b6f

Chlororespiratory pathwayB.

Asada-Halliwell pathwayA.

SODPSIO2

– H2O2

Ascorbate GSSH

MDHA GSH

NADPH

NADP+

O2

O2

H2O

Thylakoidmembrane

including the chloroplast. It is able to scavenge and inac-tivate superoxide radicals by forming hydrogen peroxideand molecular oxygen (Equation 14.4):

2O2− + 2H+ → H2O2 + O2 (14.4)

The H2O2, in turn, is reduced to water in the chloroplastby sequential reduction with ascorbate (vitamin C), glu-tathione, and NADPH (Figure 14.15A). It is interestingto note that plant chloroplasts normally exhibit relativelyhigh concentrations of ascorbate (0.5 to 1.0 μmol mg−1

Chl in Arabidopsis thaliana), which can vary dependingon the growth conditions. Reduction of H2O2 is neces-sary in order to prevent its reaction with O−

2 to form thehighly toxic hydroxyl radical (OH·), another exampleof an ROS that can rapidly damage proteins. This path-way for the protection against ROS is known as theAsada-Halliwell Pathway (Figure 14.15A).

In addition to the photoreduction of oxygen by PSIthrough the Mehler reaction, chloroplasts, also exhibitthe capacity to reduce O2 in the dark through the chlo-rorespiratory pathway (Figure 14.15B). Under normalgrowth conditions, this chloroplastic respiratory path-way results in the reduction of the thylakoid PQ pool inthe dark. This pathway involves an NAD(P)H dehydro-genase (Ndh) that reduces PQ nonphotochemically by

Chapter Review 259

using stromal reductants (stromal pool). However, it hasbeen proposed that subsequently the thylakoid-boundplastid terminal oxidase (PTOX) couples the oxidationof plastoquinol with the reduction of O2 to water evenin the light. Recently, the gene for a plastid terminaloxidase (PTOX) was identified in Arabidopsis thalianaas well as Chlamydomonas reinhardtii. The sequence ofPTOX is very similar to the alternative oxidase (AOX)present in mitochondria (Chapter 10).

Since O2 can act as an alternative electron acceptorfor the photosynthetic electron transport chain eitherthrough the Mehler reaction or through the chlorores-piratory pathway, both of these processes representpotential mechanisms to keep the PQ pool oxidized anddecrease the probability of chronic photoinhibition dur-ing acclimation irrespective of the stress which initiatesthe short-term and long-term mechanisms of acclima-tion. However, the specific contribution of the Mehlerreaction and chlororespiration to photoprotection dur-ing acclimation to stress appears to be both speciesdependent as well as dependent upon the specific stressto which a plant is exposed. Furthermore, irrespectiveof whether the short-term or long-term mechanism ofacclimation is in response to high light, water deficits,or temperature, the maintenance of photostasis, that isa balance in energy budget, appears to be an importantfeature of the newly attained acclimated state.

SUMMARY

Stress-resistant or stress-tolerant plants exhibit theability to acclimate or adjust to the environmentalstress. Although specific changes in physiology broughtabout by acclimation to a stress are not heritable, thecapacity of plants to acclimate reflects their remark-able plasticity. Plants exhibit stress tolerance becauseof their genetic capacity to acclimate and establisha new homeostatic state over time. Since acclima-tion represents an integrated hierarchy of short- andlong-term responses, the attainment of the new accli-mated state is a complex, time-nested response to astress. Short-term mechanisms usually take place veryearly in the acclimation response and include: (1) statetransitions in response to rapid changes in light qualityto balance energy distribution between PSII and PSI;(2) the induction of the xanthophyll cycle and non-photochemical quenching (NPQ) to photoprotect PSIIagainst sudden exposures to excess light and the poten-tial for chronic photoinhibition; (3) the accumulationof compatible solutes for cellular osmotic adjustmentto prevent water loss upon exposures to water deficitsand the stimulation of photorespiration to consumeexcess energy; (4) the modulation of membrane fluidityand the induction of specific cold regulated genes inresponse to low-temperature acclimation.

A unique characteristic of plant respiration isthat its Q10 is not constant but varies as a functionof short-term changes in temperature. In contrast tothe short-term mechanisms that initiate acclimationresponses, the long-term mechanisms of acclimationmay result in phenotypic changes. Photoacclimation tohigh light leads to a reduction in chlorophyll contentwith concomitant decreases in the abundance of PSIIlight-harvesting complex polypeptides and increasesin Rubisco content relative growth at low light. Thisresponse is a consequence of retrograde regulation ofnuclear genes coding for PSII light-harvesting polypep-tides by the redox status of the PQ pool. This responseto high light is mimicked by acclimation to low tem-perature because both excess irradiance and low tem-perature increase the reduction state of the PQ poolin a similar way. This can be measured as excita-tion pressure which is a measure of the proportion ofclosed PSII reaction centers. Although photoautrophsacclimate to light quality, the extent of such accli-mation appears to be species dependent. The mostdramatic example of acclimation to light quality isexhibited by many cyanobacteria through a processcalled complementary chromatic adaptation. Growthand development of winter cereals at low tempera-ture stimulates photosynthetic and respiratory capacitywhich results in an increased tolerance to photoin-hibition. In response to growth under water deficitconditions, plants reduce shoot–root ratios and totalleaf area to reduce water loss due to transpiration. Coldacclimation of herbaceous as well as woody plants leadsto increased freezing tolerance which is measured asLT50. Antifreeze proteins do not affect LT50 butcontrol the rate of extracellular ice formation. Plantscapable of acclimating to high temperature are con-sidered thermotolerant and exhibit the ability to shiftthe temperature profile for maximum photosyntheticcapacity towards high temperatures. Many plants canutilize O2 as an alternative electron acceptor in pho-tosynthetic electron transport during acclimation tovarious stresses through either the Mehler reaction orchlororespiration. Photostasis is the maintenance of abalance between energy absorbed versus energy uti-lized through metabolism and growth. The attainmentof photostasis appears to be a common feature of accli-mation to high light, water deficits, and temperature.

CHAPTER REVIEW

1. How are stress and acclimation related to oneanother?

2. How is the PQ pool of photosynthetic electrontransport involved in regulating state transitions?

3. What is the role of the xanthophyll cycle inphotoprotection from excess irradiance?

260 Chapter 14 / Acclimation to Environmental Stress

4. List three compatible solutes. What istheir role in acclimation to water stress?

5. What are cor genes?6. How does the Q10 for plant respiration vary

with short-term changes in temperature?7. What is the phenotypic change associ-

ated with photoacclimation in terrestrialplants and green algae? What are the func-tional consequences of these changes?

8. Define retrograde regulation. How is itinvolved in regulation of photoacclimation?

9. Define complementary chromatic adaptation.Explain why this name should be considered a mis-nomer.

10. Explain why acclimation to low temperaturecan mimic photoacclimation in green algae.

11. Why do cold-acclimated winter cereals exhibitan increased tolerance to photoinhibition?

12. Describe two phenotypic changes associ-ated with acclimation to water stress inherbaceous plants. How do these phenotypicchanges mitigate the effects of water deficits?

13. Define LT50. What abiotic factors regulatemaximum LT50 in herbaceous plants?

14. Where are antifreeze proteins localized. What istheir role?

15. What role can O2 play in acclimation to variousstresses?

FURTHER READING

Anderson, J. M. 1995. The grand design of photosynthesis:Acclimation of the photosynthetic apparatus to environ-mental cues. Photosynthesis Research 46:129–139.

Anderson, J. M. 1986. Photoregulation of the composition,function, and structure of thylakoid membranes. AnnualReview of Plant Physiology 37:93–136.

Apel, K., H. Hirt. 2004. Reactive oxygen species: Metabolism,oxidative stress and signal transduction. Annual Review ofPlant Biology 55:373–399.

Atkin, O. K., M. G. Tjoelker. 2003. Thermal acclimation anddynamic response of plant respiration to temperature.Trends in Plant Science 8:343–351.

Buchanan, B. B., W. Gruissem, R. L. Jones. 2000.Biochemistry and Molecular Biology of Plants. Rockville,MD: American Society of Plant Physiologists.

Demmig-Adams, B., W. W. Adams III, A. Mattoo. 2006.Photoprotection, Photoinhibition, Gene Regulation and Envi-ronment. Advances in Photosynthesis and Respiration, Vol.21. Dordrecht: Springer.

Griffith, M., M. W. F. Yaish. 2004. Antifreeze proteins inoverwintering plants: A tale of two activities. Trends inPlant Science 9:399–405.

Huner, N. P. A., G. Oquist, G. F. Sarhan. 1998. Energy bal-ance and acclimation to light and cold. Trends in PlantScience 3:224–230.

Ingram, J., D. Bartels. 1996. The molecular basis of dehydra-tion tolerance in plants. Annual Review of Plant Physiologyand Plant Molecular Biology 47:377–403.

Kehoe, D. M., A. Gutu. 2006. Responding to color: The reg-ulation of complementary chromatic adaptation. AnnualReview of Plant Biology 57:127–150.

Mittler, R. 2006. Abiotic stress, the field and environment andstress combination. Trends in Plant Science 11:15–19.

Nishida, I., N. Murata. 1996. Chilling sensitivity in plantsand cyanobacteria: The crucial contribution of mem-brane lipids. Annual Review of Plant Physiology and PlantMolecular Biology 47:541–568.

Nott, A., H.-S. Jung, S. Koussevitsky, J. Chory. 2006.Plastid-to-nucleus retrograde signalling. Annual Reviewof Plant Biology 57:739–759.

Oquist, G., N. P. A. Huner. 2003. Photosynthesis of overwin-tering evergreen plants. Annual Review of Plant Biology54:329–355.

Pearcy, R. W. 1996. Acclimation to sun and shade. In: A. S.Raghavendra (ed.), Photosynthesis: A Comprehensive Trea-tise, pp. 250–263. Cambridge: Cambridge UniversityPress.

Potters, G., T. P. Pasternak, Y. Guisez, K. J. Palme, M.A. K. Jansen. 2007. Stress-induced morphogenicresponses: Growing out of trouble? Trends in PlantScience 12:98–105.

Sack, L., N. M. Holbrook. 2006. Leaf hydraulics. AnnualReview of Plant Biology 57:361–381.

Smirnoff, N. 1995. Environment and Plant Metabolism: Flexi-bility and Acclimation. Bios Scientific Publishers.

Stitt, M., V. Hurry. 2002. A plant for all seasons: Alterationin photosynthetic carbon metabolism during cold accli-mation in Arabidopsis. Current Opinion in Plant Biology5:199–206.

Thomashow, M. F. 1999. Plant cold acclimation: Freezingtolerance genes and regulatory mechanisms. AnnualReview of Plant Physiology and Plant Molecular Biology50:571–599.

15Adaptations to the Environment

Previously, we defined stress as a negative effect onplant homeostasis which can be measured when a plantis exposed to a sudden change from some optimal con-dition for growth to some suboptimal abiotic conditionsuch high light, high or low temperature, and waterdeficit. In Chapter 14, we discussed the capacity ofmany C3 plants to acclimate or adjust to such stressesover time. The myriad mechanisms available for accli-mation at the whole plant, physiological, biochemical,and molecular levels in response to a change in theirenvironment appear to be integrated in a time-nestedfashion. In Chapter 14 we discovered that the abilityof plants to acclimate to specific stresses varied as afunction of the species examined. The capacity of aplant to acclimate reflects the remarkable plasticity inthe form and function of many plants. However, it isnot the specific phenotypic and physiological changesinduced during acclimation period that are passed on tothe next generation, but rather, it is the inherent abilityto change in the first place that is heritable. Thus, plas-ticity is the genetic trait that characterizes plants ableto tolerate environmental change during the normal lifecycle of a plant from seed germination, to vegetativegrowth, to reproductive development, and ultimatelyto sensescence. These latter topics are discussed in

detail in Chapters 16 to 27. In contrast to acclimation,adaptation refers to heritable modifications in struc-ture or function that increase the fitness of the organ-ism in a stressful environment. This means that plantadaptation to the environment takes place over a muchlonger time scale than acclimation. Since adaptation isa consequence of natural selection, adaptation occursover an evolutionary time scale. Adaptation is a conse-quence of the complement of genes found in the genomeof a plant species as a result of natural selection over thecourse of many, many generations. Thus, it is possiblethat some plants adapted to a very special niche mayexhibit limited plasticity to acclimate to sudden changesin their environment.

In this chapter we will discuss

• the physiology of plants adapted to low-light versushigh-light environments,

• the role of C4 photosynthesis in adaptation of plantsto high temperature,

• CAM photosynthesis and its relationship to adapta-tion to desert environments, and

• the impact of physiology on the structure of plantcommunities, biomes, as well as global weatherpatterns.

261

262 Chapter 15 / Adaptations to the Environment

15.1 SUN AND SHADE ADAPTEDPLANTS RESPONDDIFFERENTIALLY TOIRRADIANCE

Shade plants adapted to growth on the floor of a tropicalrainforest not only may receive less than 1 percent ofthe incident photon fluence rate measured at the top ofthe canopy, but are also exposed to light enriched in thegreen and far-red wavelengths of the visible spectrum.Plants are classified as either sun or shade species,depending on the ability to acclimate to changes inirradiance. The ability to acclimate to irradiance is aninherited trait and is determined by species genotypewhich is a consequence of many, many successive gen-erations of natural selection to the light environmentexperienced by the species in its native habitat. Obli-gate shade plants are adapted to survive extreme shadeconditions and exhibit minimal capacity to acclimate tohigh-light conditions. Generally, leaves of shade speciesare thinner and exhibit higher chlorophyll contents thanthose of sun species (Table 15.1). Chloroplasts of obli-gate shade species are highly stacked with as many as 100thylakoids per granum which occupy most of the stromalvolume whereas the number of appressed thylakoids ina granum of sun species typically varies from 5 to 30 (seeChapter 5). In addition to their higher chlorophyll con-tent, shade adapted species typically exhibit lower ratiosof chlorophyll a to chlorophyll b (2.0 to 2.5) comparedto sun species (3.2 to 3.6). Since chlorophyll b is associ-ated with the light-harvesting complexes of PSI (LHCI)and PSII (LHCII) (Chapter 7), the lower chlorophyll a:chlorophyll b ratio is indicative of more light harvest-ing complex present in shade adapted species than insun species. This may be reflected in a higher photo-synthetic efficiency at low, light-limiting irradiance inshade adapted species than in sun species (Figure 15.1A).However, the light-saturated rate of photosynthesis isgenerally lower in shade species (Figure 15.1A) due toa lower Rubisco content in shade species than in sunspecies. In contrast to obligate shade species, sun species

TABLE 15.1 A comparison of averagechlorophyll contents of leaves from sun and shadespecies grown in their native habitats.

Fresh weight Total Chlorophyllper leaf area per fresh weight per leaf area

Species (g dm−2) (mg g−1) (mg dm−2)

Shade 0.80 3.1 5.3Sun 2.5 1.9 4.7

(Adapted from Boardman, N. K. 1977. Comparative photosynthesis ofsun and shade plants. Annual Review of Plant Physiology 28: 355–377.)

exhibit remarkable plasticity in their ability to changelight-saturated rates of photosynthesis in response toincreasing growth irradiance (Figure 15.1B). As a con-sequence of their limited ability to acclimate to growthirradiance, clones of Solidago virgaurea native to shadedhabitats were incapable of acclimating to high lightintensity. High light caused a reduction in both photo-synthetic efficiency and photosynthetic capacity. Theseresults are consistent with the observation that exposureof leaves of obligate shade plants to high light usu-ally leads to chronic photoinhibition followed by leafchlorosis and ultimately to death (Chapter 13 and 14). Incontrast, sun species are less susceptible to chronic pho-toinhibition than shade species because of their greatercapacity to convert the absorbed light into fixed carbonand prevent damage to PSII. On the other hand, obli-gate sun plants exposed to growth under extremely lowirradiance may cause etiolation (see Chapter 5) whichmay be fatal because they are growing below theirlight compensation point. This is the irradiance at

30

20

Gro

ss R

ate

of P

hoto

synt

hesi

s(μ

mol

CO

2 m

−2 s

−1)

10

1000

Irradiance (μmol m−2 s−1)

2000

Shade Plant

Sun Plant

0

20

15

10

Net

Rat

e of

Pho

tosy

nthe

sis

(μm

ol C

O2 d

m−2

min

−1)

5

500 1000 1500 2000

LL

IL

HL

Irradiance (μmol m−2 s−1)

0

0

A.

B.

FIGURE 15.1 The photosynthetic response of sun andshade species to irradiance. (A) The light response curvesof a sun species (Atriplex patula) and a shade species(Asarum caudatum) grown under natural conditions. (B)The light response curves of the sun species, Atriplexpatula, to increasing growth irradiance. Growth irradi-ance included low light (LL, 2 mW cm-2), intermediatelight (IL, 6.3 mW cm-2) and high light (HL, 20 mWcm−2). (Adapted from Boardman, N. K. 1977. Compar-ative photosynthesis of sun and shade plants. AnnualReview of Plant Physiology 28:355–377.)

15.2 C4 Plants are Adapted to High Temperature and Drought 263

which the rate of photosynthesis equals the rate ofrespiration. Below the light compensation point, plantsare respiring faster than photosynthesis can assimilateCO2. Clearly, long-term exposure to such conditionswill be fatal for a plant. Typically, shade adapted plantsexhibit lower light-compensation points than sun speciesbecause the rate of respiration is lower in the formerthan the latter.

It appears that the physiological plasticity for accli-mation to growth irradiance varies considerably betweengenotypes. The extent of this plasticity reflects anadaptation to a plant’s natural habitat. Obligate shadeadaptation appears to preclude acclimation to highlight whereas obligate sun adaptation appears to pre-clude acclimation to extreme low-light conditions. Thus,adaptation for high photosynthetic efficiency under oneextreme regime appears to preclude the maintenance ofhigh photosynthetic efficiency under the other extreme.

15.2 C4 PLANTS ARE ADAPTEDTO HIGH TEMPERATUREAND DROUGHT

15.2.1 THE C4 SYNDROME IS ANOTHERBIOCHEMICAL MECHANISM TOASSIMILATE CO2

Plants that incorporate carbon solely through the PCRor Calvin cycle are generally known as C3 plants becausethe product of the first carboxylation reaction cat-alyzed by the enzyme, Rubisco (Equation 15.1), is thethree-carbon acid, 3-phosphoglyceric acid (PGA)

RuBP + CO2 → 2PGA (15.1)

(Chapter 8). Certain other groups, known as C4 plants,are distinguished by the fact that the first product is a

four-carbon acid oxaloacetate (OAA). C4 plants alsoexhibit a number of specific anatomical, physiological,and biochemical characteristics that constitute the C4syndrome. One particular anatomical feature charac-teristic of most C4 leaves is the presence of two distinctphotosynthetic tissues (Figure 15.2). In C4 leaves thevascular bundles are quite close together and each bun-dle is surrounded by a tightly fitted layer of cells calledthe bundle sheath. Between the vascular bundles andadjacent to the air spaces of the leaf are the more looselyarranged mesophyll cells. This distinction betweenmesophyll and bundle sheath photosynthetic cells, calledKranz anatomy (see below), plays a major role in theC4 syndrome.

C4 plants are generally of tropical or subtropicalorigin representing nearly 1,500 species spread throughat least 18 different angiosperm families (3 monocots,15 dicots). Interestingly, no one family has been foundto express the C4 syndrome exclusively—all 18 familiescontain both C3 and C4 representatives. This suggeststhat the C4 cycle has arisen rather recently in evolu-tion of angiosperms and in a number of diverse taxaat different times. Under conditions of high fluencerates and high temperature (30◦ to 40◦C) the photo-synthetic rate of C4 species may be two to three timesgreater than that of C3 species. C4 plants also have aspecialized leaf anatomy comprised of mesophyll cellsand bundle-sheath cells. The anatomical arrangementsand their impact on leaf photosynthesis are exploredfurther in the next section. Generally, C4 plants appearto be better equipped to withstand drought and are ableto maintain active photosynthesis under conditions ofwater stress that would lead to stomatal closure andconsequent reduction of CO2 uptake by C3 species.All of these features appear to be a consequence ofthe CO2-concentrating capacity of C4 plants and theresulting suppression of photorespiratory CO2 loss.

FIGURE 15.2 Leaf of a C4 grass.Cross-section of a leaf of maize (Zeamays), showing the arrangement of mes-ophyll and bundle-sheath cells. Notehigh concentration of chloroplasts inbundle-sheath cells. (From Esau, K.1977. Anatomy of Seed Plants. New York:Wiley. Reprinted by permission.)

264 Chapter 15 / Adaptations to the Environment

The key to the general C4 cycle is the enzymephosphoenol pyruvate carboxylase (PEPcase), whichcatalyzes the carboxylation of phosphoenol pyruvate(PEP) using the bicarbonate ion HCO−

3 as the sub-strate rather than CO2 (Figure 15.3). The product ofthe PEPcase reaction, the 4-carbon acid, oxaloacetate(OAA), is moderately unstable. It is quickly reducedto either malate or transaminated to aspartate—both ofwhich are more stable—and transported out of the mes-ophyll cell into an adjacent bundle-sheath cell. Once inthe bundle-sheath cell, the C4 acid undergoes a decar-boxylation reaction and the resulting CO2 is availablefor reduction to triose sugars via the PCR cycle in thebundle-sheath chloroplast. The C3 acid—either pyru-vate or alanine—that remains after decarboxylation isthen transported back into the mesophyll cell. Herethe alanine or the pyruvate is converted to pyruvate.The pyruvate is then phosphorylated by the enzyme,pyruvate, phosphate dikinase, to regenerate the orig-inal acceptor molecule, phosphoenol pyruvate (PEP)(Figure 15.3).

There are certain similarities between the PCRcycle and C4 metabolism. Like Rubisco, the PEP-case carboxylation reaction is virtually irreversible and,consequently, energetically very favorable. Reducingpotential is required at some point to remove the prod-uct and ATP is required to regenerate the acceptormolecule, PEP, and thus keep the reaction going. Avery significant difference, however, is that once in the

bundle-sheath cell, the C4 acid is decarboxylated, givingup the CO2 originally assimilated in the mesophyll cell.This decarboxylation means that, unlike the C3 cycle,the C4 cycle does not of itself result in any net carbonreduction. The plant relies ultimately on the operationof the PCR cycle in the bundle-sheath chloroplast forthe synthesis of triose phosphates.

The principal effect of the C4 cycle is that it con-centrates CO2 in the bundle-sheath cells where theenzymes of the PCR cycle are located. By shuttling theCO2 in the form of organic acids it is possible to buildmuch higher CO2 concentrations in the bundle-sheathcells than would be possible relying on the diffusionof CO2 alone. Results of studies employing radiola-beled 14CO2 have indicated the concentration of CO2in bundle-sheath cells may reach 60 mM; about tenfoldhigher than that in C3 plants. Higher CO2 concen-trations would suppress photorespiration and supporthigher rates of photosynthesis. Under optimal condi-tions, C4 crop species can assimilate CO2 at rates twoto three times that of C3 species. All this productivitydoes not, however, come free. There is an energy costto building the CO2 concentration in the bundle-sheathcells. For every CO2 assimilated, two ATP must beexpended in the regeneration of PEP. This is in addi-tion to the ATP and NADPH required in the PCRcycle. Thus the net energy requirement for assimila-tion of CO2 by the C4 cycle is five ATP and twoNADPH.

COO-

C = O

CH2

COO-COO-

C

CH2

CO2 HCO3-

NADHNAD+

C4Acid

C4Acid

CO2 PGA

PCRRuBP

SugarsStarch

C3Acid

C3Acid

Phosphoenolpyruvate( PEP )

2AMP + PPi2ATP + Pi

Pi

PO

MESOPHYLL CELL BUNDLE-SHEATH CELL

Oxaloacetate

FIGURE 15.3 Schematic of the general plan of the C4 photosynthetic carbon assimila-tion cycle. Initial carboxylation of phosphoenol pyruvate (PEP) to oxaloacetate (OAA)in the mesophyll cell is followed by translocation of a C4 acid to the bundle-sheathcell where it undergoes a decarboxylation. The resulting C3 acid is returned to themesophyll cell to complete the cycle. The CO2 released in the decarboxylation step isassimilated by the PCR cycle in the bundle-sheath chloroplast. The C4 cycle servesto concentrate CO2 in the bundle-sheath cell and suppress photorespiration.

15.2 C4 Plants are Adapted to High Temperature and Drought 265

15.2.2 THE C4 SYNDROME IS USUALLYASSOCIATED WITH KRANZ LEAFANATOMY

The anatomy of a typical C4 leaf is shown in Figure 15.2.Recall that the photosynthetic parenchyma cells in a typ-ical C3 leaf are organized into two distinct tissues—anupper region of tightly packed palisade cells and themore loosely arranged spongy mesophyll cells border-ing large air spaces (Chapter 8). The C4 leaf, on theother hand, is generally thinner than the C3 leaf, the vas-cular bundles are closer together, and the air spaces aresmaller. Moreover, there is only one type of mesophyllcell, loosely arranged in the fashion of the spongy mes-ophyll in the C3 leaf. Surrounding each vascular bundleis a sheath of tightly packed, thick-walled cells contain-ing large numbers of chloroplasts. Indeed, C4 plantscan often be recognized by the prominent, dark-greenveins. Because of the wreathlike configuration of thesebundle-sheath cells, this arrangement is known as Kranzanatomy (Kranz, wreath, German).

The characteristic anatomical arrangement of theC4 leaf ensures a short diffusion path for CO2 to theinitial carboxylation site in the mesophyll cell. A shortdiffusion path together with the uniform distribution ofthe initial carboxylating enzyme throughout the cytosolmakes for a more efficient CO2 trapping. Moreover, nomesophyll cell is more than two or three cells distantfrom a bundle-sheath cell, which no doubt facilitates thetransfer of the C4 and C3 acids. The high chloroplastdensity in the bundle-sheath cell is apparently necessaryto process the high concentrations of CO2 generatedby the C4 system. Finally, the close juxtaposition of thePCR cycle cells to the vascular tissue means that product(i.e., sugars) can be quickly exported from the leaf whenrequired. Overall it is safe to conclude that the effective-ness of the C4 system is enhanced by these anatomicaladaptations. It should be pointed out, however, thatsome C3 dicots do have well-developed bundle sheaths,although they generally have few or no chloroplasts.Thus the presence of Kranz anatomy cannot, in itself,be taken as evidence of the C4 syndrome.

15.2.3 THE C4 SYNDROME HASECOLOGICAL SIGNIFICANCE

C4 plants exhibit a number of physiological attributesthat appear to be immediate consequences of theirunique carbon metabolism. It is generally perceived thatthese physiological attributes may lead to higher photo-synthetic productivity under certain conditions and havesignificant ecological consequences. Unlike C3 plants,photosynthesis of C4 plants is not inhibited by O2, andthey exhibit no postillumination CO2 burst and have avery low CO2 compensation point (Table 15.2). TheCO2 compensation concentration is the ambient carbon

TABLE 15.2 A comparison of significantfeatures of C3 and C4 plants.

C3 C4

Photorespiration yes not detectableCO2 compensation (μlCO21−1)

20–100 0–5

Temperature optimum(◦C)photosynthesis 20–25 30-45Rubisco 20–25PEPcase 30–35Quantum yield as afunction of temperature

declining steady

Transpiration Ratio 500–1000 200–350Light saturation (μmolephotons m−2 s−1)

400–500 does notsaturate

dioxide concentration at which the rate of CO2 uptake(for photosynthesis) is balanced by the rate of CO2evolution (by respiration). In a closed environment, theCO2 compensation concentration would be the stableCO2 concentration in the air when CO2 uptake andevolution have come into equilibrium. For C3 plants,the CO2 compensation point falls into the range of 20to 100 μl CO2 per liters whereas the values for C4 plantsare in the range of 0 to 5 μl l−1.

On the basis of the above observations, it is reason-able to conclude either that photorespiration is absentfrom C4 plants or that the process is suppressed. How-ever, although the activity of the glycolate pathway(Chapter 8) is very low in some C4 plants (Sorghumbicolor, for example), most C4 plants appear to have bothperoxisomes and the metabolic machinery to supportphotorespiration. The weight of evidence thus favorsthe conclusion that C4 plants do photorespire, but atmuch reduced rates. The high level of CO2 developed inthe bundle-sheath cells would tend to suppress photores-piration by outcompeting O2 for binding to Rubisco. Inaddition, the anatomical and biochemical adaptations ofC4 leaves ensures that any CO2 that might escape thebundle-sheath cell is trapped and reassimilated by PEP-case in the mesophyll cells, before it has the opportunityto escape from the leaf. Thus C4 leaves are not onlyefficient CO2 absorbers, but also effectively trap andrecirculate any CO2 that might be produced in the leaf.

15.2.4 THE C4 SYNDROME ISDIFFERENTIALLY SENSITIVETO TEMPERATURE

In addition to the virtual absence of photorespiration,most C4 plants tend to have a higher temperature

266 Chapter 15 / Adaptations to the Environment

0.02

0.04

0.06

0.08

Qua

ntum

yie

ld. (m

ole

CO

2 p

er a

bsor

bed

mol

e ph

oton

s)

Encelia californica, C3

Atriplex rosea, C4

10 20 30 40

Leaf temperature. ( C)

FIGURE 15.4 Quantum yield for uptake of CO2 in a C4and C3 plant as a function of leaf temperature. (FromEhleringer, J., O. Bjorkman. 1977. Plant Physiology59:86–90. Copyright American Society of Plant Physi-ologists.)

optimum (30–45◦C) than C3 plants (20–25◦C). Thisdifference is due primarily to the differential stabilityof the photorespiration relative to photosynthesis. Athigh temperatures between 40◦ and 50◦C, the rate ofphotosynthesis decreases to a greater extent than therate of photorespiration. This is at least in part dueto the higher temperature stability of some of the C4cycle enzymes compared to C3 enzymes. As a result,photosynthetic efficiency or the quantum yield of pho-tosynthesis in C3 plants tends to decline with increasingleaf temperature while it remains fairly constant in C4plants (Figure 15.4). The decline of quantum yield in C3plants is due, in part, to decreased carboxylation activ-ity of Rubisco at the higher temperatures. This trendis exacerbated by changes in the relative solubility ofCO2 and O2. The solubility of gases generally declineswith increasing temperature, but the solubility of CO2 isaffected more than the solubility of O2. Consequently,higher temperatures increasingly favor oxygenation byRubisco.

Another interesting feature of C4 plants is theirgeneral low-temperature sensitivity. While there aresome cold-tolerant C4 species, most perform poorly, ifat all, at low temperature. Zea mays, for example, willnot grow at temperatures below 12◦ to 15◦C. This lowerlimit for growth is probably set, in part, by the enzymepyruvate, phosphate dikinase (Figure 15.3), which is coldlabile and experiences a substantial loss of activity below

12◦C. This would inhibit the regeneration of PEP, anecessary substrate for PEP carboxylase (Equation 15.2).

PEP + CO2 → OAA (15.2)

15.2.5 THE C4 SYNDROME ISASSOCIATED WITH WATERSTRESS

Photosynthesis in most situations is limited by avail-able CO2 and water. In C3 plants, even moderate waterstress will initiate closure of the stomata (Chapter 13)and reduce the available supply of CO2. The low CO2compensation point of C4 plants means that they canmaintain higher rates of photosynthesis at lower CO2levels. Thus C4 plants gain an advantage over C3 plantswhen the stomata are partially closed to conserve waterduring a period of water stress (Chapter 13). An effectivemeasure of this advantage is the value of the transpi-ration ratio (TR). The transpiration ratio relates theuptake of CO2 to the loss of water by evaporation (tran-spiration) from the leaf. The inverse of transpirationratio, called water use efficiency (WUE), is often citedin ecological literature.

TR = moles H2O transpired/moles CO2 assimilated(15.3)

WUE = moles of CO2 assimilated/

moles H2O transpired (15.4)

Transpiration ratios for C4 plants are typically inthe range of 200 to 350, while for C3 plants valuesin the range of 500 to 1000 are often cited. The lowtranspiration ratio for C4 plants reflects their capac-ity to maintain high rates of photosynthesis whileeffectively conserving water. Even under ideal condi-tions, CO2 supply limits photosynthesis in C3 plantsto the extent that light saturation occurs at fluencerates about 25 percent of full sunlight. C4 plants,on the other hand, never really saturate, even at fullsunlight (Figure 15.5). Even so, C4 photosynthesisis not necessarily more efficient than C3 photosyn-thesis. At leaf temperatures below 30◦C the quan-tum yield for C4 plants is actually lower than forC3 plants—that is, C4 photosynthesis is less efficient(Figure 15.4). The low quantum efficiency of C4 plantsreflects an additional light requirement that can beaccounted for by the ATP required by the pyruvate,phosphate dikinase reaction (Figure 15.3). How can thelower photosynthetic efficiency of C4 plants at lowertemperatures be reconciled with their apparent higherproductivity? Recall that C4 plants are native to trop-ical or subtropical habitats where there is usually anabundance of light. C4 plants generally exhibit higherlight-saturated rates of photosynthesis than C3 plants,which reflects a higher photosynthetic capacity (Table 15.1,Figure 15.5) due to their higher content of compo-nents of photosynthetic electron transport and certain

15.3 Crassulacean Acid Metabolism is an Adaptation to Desert Life 267

500

0

1000 1500 2000

Fluence rate (μmole m−2 s−1 PAR)

Rat

e of

CO

2 U

ptak

e

C3

C4

FIGURE 15.5 Fluence response curves for typical C3 andC4 plants.

photosynthetic enzymes such as Rubisco. Thus, C4plants can take advantage of some of the excess avail-able light to generate the ATP needed to run the C4cycle, concentrate the CO2, and increase net carbonassimilation.

However, C4 plants are not competitive in allsituations. C3 plants may equal or even exceed C4 plantsin productivity under conditions of lower temperaturesand lower irradiance, and where water availability isnot limiting as generally found in temperate climates.This differential sensitivity to environmental conditionsbetween C3 and C4 plants is reflected in the observationthat, to the frustration of homeowner and farmer alike,many of our more aggressive weeds are C4 species.These include crabgrass (Digitaria sanguinalis), Russianthistle (Salsola kali), and several species of pigweed(Amaranthus) that often take over during the hot, drymonths in the middle of summer. Many of the morehighly productive crop species also fall within the C4group, including sugarcane (Saccharum officinarum),sorghum (Sorghum bicolor), maize (Zea mays), and millet.However, plant productivity is ultimately dependentupon the growth environment. Consequently, theadaptations in leaf anatomy coupled to the biochemistryof CO2 assimilation that distinguish C4 plants fromC3 plants have a dramatic effect on the geographicaldistribution of these species.

15.3 CRASSULACEAN ACIDMETABOLISM IS ANADAPTATION TO DESERTLIFE

Another CO2 concentrating mechanism biochemicallycomparable to that of C4 plants is crassulacean acid

metabolism (CAM)—so named because it was origi-nally studied most extensively in the family Crassulaceae.This specialized pattern of photosynthesis has now beenfound in some 23 different families of flowering plants(including the Cactaceae and Euphorbiaceae), one fam-ily of ferns (the Polypodiaceae), and in the primitiveplant Welwitschia. Like C4 plants, however, most fami-lies, with the exception of Crassulaceae and Cactaceae,are not exclusively CAM. Most families will have C3representatives as well and some are known to con-tain all three photosynthetic patterns; C3, C4, andCAM.

The unique features of CAM permit a remarkabledegree of water conservation. Individual species utiliz-ing this pathway are thus especially adapted to survivalin extremely dry, or xerophytic, habitats. They are also,without exception, succulent plants—characterized bythick, fleshy leaves (or, as in the cacti, photosyntheticstems) whose cells contain large, water-filled vacuoles.Other than succulence no particular anatomical mod-ifications appear to be required for CAM. However,although succulence appears prerequisite for CAM, notall succulent plants exhibit CAM.

One of the most striking features of CAM plantsis an inverted stomatal cycle—the stomata open mainlyduring the nighttime hours and are usually closed duringthe day. This means that CO2 uptake also occurs mainly atnight (Figure 15.6). In addition, CAM plants are char-acterized by an accumulation of malate at night and itssubsequent depletion during daylight hours and storagecarbohydrate levels that fluctuate inversely with malatelevels. Nocturnal stomatal opening supports a carboxy-lation reaction producing C4 acids that are stored inthe large, watery vacuole (Figure 15.6). Accumulation ofthe organic acids leads to a marked acidification of thesecells at night. The acids are subsequently decarboxylatedduring daylight hours and the resulting CO2 is fixed bythe PCR cycle.

As in C4 plants, the enzyme PEP carboxylaseis central to CAM operation. During the night,the immediate product, OAA, is rapidly reduced byNAD-dependent malate-dehydrogenase to malate,which is then stored in the vacuole. During the daylighthours, malate is retrieved from the vacuole, decarboxy-lated (by NAD-malic enzyme in the Crassulaceae),and the CO2 diffuses into the chloroplast where it isconverted to triose phosphates by the C3 PCR cycle.The large amounts of PEP required to support thecarboxylation reaction appear to be derived from thebreakdown of starch and other storage glucans bythe enzymes of the glycolytic pathway (Chapter 10).The fate of C3 acid (pyruvate or PEP) resulting fromdecarboxylation is uncertain, but the weight of evidenceis that it is reduced to triose phosphate, which in turncan be converted back to glucose or starch (Figure 15.6).

268 Chapter 15 / Adaptations to the Environment

DARK LIGHT

Stomata: OPEN CLOSED OPEN

Acid contentCO2

Uptake

8 10 12 2 4 6 8 10 12NoonMidnight

2 4 6 8

CO2H2O

CO2H2O

HCO3−

CO2

H2O

PEP

Pi

OAA

StarchNADH

NAD+

Malate

Malic acid

Chloroplast

Vacuole

Starch

PCRCycle

CO2

Pyruvate

Malate

Malic acid

Vacuole

Chloroplast

Triose - PTriose - P

FIGURE 15.6 An outline of crassulacean acid metabolism. Above: Curves illustratingstomatal opening, CO2 uptake, and changing acid content of cell vacuoles over a24-hour period. Stomata open in the dark to admit CO2 and close during the day toconserve water. Below, right: While the stomata are closed during the day, storedCO2 is released to be assimilated via the PCR cycle.

15.3.1 IS CAM A VARIATION OF THE C4SYNDROME?

CAM and C4 plants share certain similarities, but thereare significant differences. A comparison between CAMand the C4 cycle is unavoidable since they both usecytoplasmic PEPcase to form C4 acids from PEP andbicarbonate, and in both cases the acids are subse-quently decarboxylated to provide CO2 for the PCRcycle. However, there are two significant differences.The first is that the C4 cycle requires a specializedanatomy by which C4 carboxylation is spatially sepa-rated from the C3 PCR cycle— in CAM both occur in

the same cells but are separated in time. Second, in CAMthere is no closed cycle of carbon intermediates as thereis in C4 plants. Instead, the PEP required as substratefor the carboxylation reaction is derived from storedcarbohydrate. The C3 product of decarboxylation isdisposed of by a variety of metabolic paths, which nodoubt includes the resynthesis of storage carbohydrate.Thus CAM is cyclic in time only. Since CAM occursin the more primitive ferns and Welwitschia whereasthe C4 cycle is found only in angiosperms, it appearsthat CAM preceded C4 photosynthesis in evolutionarytime.

15.4 C4 and CAM Photosynthesis Require Precise Regulation and Temporal Integration 269

15.3.2 CAM PLANTS AREPARTICULARLY SUITEDTO DRY HABITATS

As mentioned above, CAM represents a particularly sig-nificant adaptation to exceptionally dry habitats. ManyCAM plants are true desert plants, growing in shallow,sandy soils with little available water. Nocturnal open-ing of the stomata allows for CO2 uptake during periodswhen conditions leading to evaporative water loss are ata minimum. Then, during the daylight hours when thestomata are closed to reduce water loss, photosynthesiscan proceed by using the reservoir of stored CO2. Thisinterpretation is supported by the transpiration ratio forCAM plants, in the range of 50 to 100, which is substan-tially lower than that for either C3 or C4 plants. There isa price to be paid, however. Rates for daily carbon assim-ilation by CAM plants are only about one-half those ofC3 plants and one-third those of C4. CAM plants canbe expected to grow more slowly under conditions ofadequate moisture. On the other hand, CO2 uptake byCAM plants will continue under conditions of waterstress that would cause complete cessation of photosyn-thesis in C3 plants and severely restrict carbon uptakeby C4 plants. CAM plants enjoy the further advantageof being able to retain and reassimilate respired CO2,thus preventing loss of carbon and helping to maintain afavorable dry weight through extended periods of severedrought.

While some species, in particular the cacti, areobligatory CAM plants, many other succulents exhibit afacultative approach to CAM. One well-studied exampleis the ice plant, Mesembryanthemum crystallinum, a fleshyannual of the family Aizoaceae. Under conditions ofabundant water supply, Mesembryanthemum assimilatescarbon as a typical C3 plant—there is no significantuptake of CO2 at night and no diurnal variation in leafcell acidity. Under conditions of limited water avail-ability or high salt concentration in the soil, CAMmetabolism is switched on. Although carbon assimi-lation by CAM is slower than with conventional C3photosynthesis, its higher water use efficiency permitsphotosynthesis to continue in times of water stress andthe plant is better able to complete its reproductivedevelopment.

15.4 C4 AND CAMPHOTOSYNTHESIS REQUIREPRECISE REGULATION ANDTEMPORAL INTEGRATION

In addition to regulation of PCR cycle enzymes dis-cussed earlier, successful operation of C4 photosynthesisand CAM also requires regulation of starch-PEP inter-conversions; storage and retrieval of malate, PEPcase

and Rubisco competition for CO2; and the temporaloperation of PEP carboxylation. PEPcase is a cytoplas-mic enzyme found in virtually all cells of higher plantswhere it serves a variety of important metabolic func-tions. However, plants with the C4 and CAM modes ofphotosynthesis contain specific isozymes1 with higheractivity levels than associated with C3 or nonphotosyn-thetic cells. Based on a variety of physiological andbiochemical considerations, it appears that PEPcaseactivity in C4 and CAM plants must be regulated bylight–dark transitions. In C4 plants, for example, PEP-case activity should be high in the light in order tomaximize availability of CO2 for the PCR cycle inthe bundle-sheath cells. Although the substrate PEPconsumed in C4 photosynthesis is derived from thebundle-sheath chloroplast, PEP is also a critical inter-mediate in glycolysis. Glycolysis is also a cytoplasmicmetabolic sequence that represents the first stage inrespiration.

Continued high PEPcase activity at night couldresult in uncontrolled utilization of PEP and seriouslyimpair respiratory metabolism. In the case of CAM it isclear that efficient operation requires the carboxylationand decarboxylation reactions—which occur within thesame cell—not be allowed to compete with each other atthe same time. In CAM, PEP carboxylase activity mustbe high at night when atmospheric CO2 is available, butshould be switched off during daylight hours in orderto avoid a futile recycling of CO2 derived from malate.Competition for CO2 is not a problem at night sinceRubisco and the PCR cycle are inoperative in the dark.

PEPcase regulation was initially studied in CAMplants but it is now evident that PEPcase activity in C4plants as well as CAM is subject to reversible activationby light–dark conditions and inhibition by malate—aform of feedback inhibition in which accumulated prod-uct reduces the activity of the enzyme. In C4 plants,light induces an increase in the catalytic activity of theenzyme while at the same time reducing its sensitiv-ity to inhibition by the product molecule, malate. Thisresults in a fivefold increase in activity in the light whenCO2 assimilation and malate production is required toboost CO2 levels in the bundle-sheath cells. In the caseof CAM, the situation is reversed. Enzyme extractedduring the night part of a diurnal cycle exhibits a highaffinity for PEP and is relatively insensitive to inhibitionby malate. Enzyme extracted during the day has a lowaffinity for PEP and is more sensitive to inhibition bymalate.

1Isozymes are different species of enzyme that catalyze thesame reaction. Isozymes are frequently coded by differentgenes and consequently have different protein structures.Although they act on the same substrate, isozymes may betissue- or organ-specific and subject to regulation by differentchemical or environmental factors.

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Recent evidence from studies of Bryophyllum andKalanchoe, both CAM plants, and the C4 plants Zeamays and Sorghum have shown that PEPcase exists intwo states; the biochemically more active form of theenzyme is phosphorylated but the less active form is not.From this it can be concluded that PEPcase activity isregulated by a light-sensitive protein kinase. Evidencefor reversible activation of a protein kinase has beenobtained from both in vivo and in vitro experiments.The reaction requires ATP and phosphorylation occursat a serine residue near the N-terminal end of theenzyme molecule. Just how light activates the proteinkinase is not known at this stage, although there is someindication that it is related to photosynthetic electrontransport. It may be that photophosphorylation suppliesthe ATP required for phosphorylation of the enzyme.

It is interesting to note that light appears to have theopposite effect in the two systems—the protein kinaseis activated by light in C4 plants but inactivated duringthe day in CAM. The effect of light and darkness inCAM plants, however, may be indirect. Studies withBryophyllum have indicated that CAM physiology andthe sensitivity of PEPcase to inhibition by malate inCAM plants may be controlled by an endogenous cir-cadian rhythm. Whatever the mechanism, spatial andtemporal coordination of C4 and C3 metabolism in C4and CAM plants is an important area where we canexpect to see exciting advances in the future.

15.5 PLANT BIOMES REFLECTMYRIAD PHYSIOLOGICALADAPTATIONS

Physiology has an impact beyond the individual plant.The physiology of a plant community or ecosystemis defined as ecophysiology. At an even larger scale,biomes represent a collection of ecosystems that arecharacterized by distinctive vegetation that is related to aparticular set of physical and environmental conditions.As examples of plant communities adapted to differentenvironments, we will discuss two specific plant biomes:the tropical rainforest biome and the desert biome.

15.5.1 TROPICAL RAIN FOREST BIOMESEXHIBIT THE GREATEST PLANTBIODIVERSITY

A tropical rainforest is a diverse and complex ecosys-tem with a number of unique geographic and physi-cal characteristics. All of the principal rainforest sys-tems of the world are located between the latitudes of23◦ 30′ N (Tropic of Cancer) and 23◦ 30′ S (Tropic ofCapricorn). The equatorial location ensures that rain-forest vegetation is subject to both high irradiance andhigh temperature. Moreover, at these latitudes seasonal

variations in both the quantity of light and photoperiod,as well as temperature, are very small. Rainfall is veryhigh, with annual rainfalls of at least 1800 to 2000 mmper year and as high as 4000 mm per year. In this pre-vailing ever-wet tropical climate, with neither water nortemperature limiting, the rainforests rank among themost highly productive ecosystems in the world. Cover-ing only about 8 percent of the global land mass, tropicalrainforests contain more than 40 percent of the world’sbiomass.

There are three principal rainforest regions world-wide. The largest by far is located in the Amazon Riverbasin of northern South America and extends, in smallpatches, through Central America into southern Mex-ico. The Amazon basin rainforest itself, stretching fromthe Atlantic Ocean in the east to the Andean moun-tains in the west, covers more than 4 million km2. Thesecond region is the African rainforest, located alongthe equatorial coast of eastern Africa and extendinginland through the Congo River basin to the mountainsof east-central Africa. Third, the Malaysian rainforestoccurs in patches throughout the Malaysian Archipelagoof southeastern Asia, primarily on the islands of Sumatra,Borneo, and New Guinea. While there are other areas,such as along the coast of northwestern United Statesand southwestern Canada, that also experience heavyrainfalls, these are not true rainforests. These coastalforests represent a southern extension of the northernconiferous, or boreal, forest, which is characterized byshorter growing seasons and persistent snow cover inthe winter.

It is generally believed that rainforests are populatedby a larger number of plant species than the rest of theworld’s ecosystems combined. At the same time, noone species accounts for more than 10–15 percent ofthe trees. There may be several hundred species perhectare and the individuals of each species may bewidely scattered. Rainforest vegetation is dominated bytall, broad-leaved woody evergreens that form a canopyso dense that little light reaches the forest floor. As aresult, available light on the forest floor is very low,limiting understory vegetation. The bulk of the canopyis located about 30 to 50 m above the ground, althoughmany individuals, called emergents, may stand as highas 45 to 70 m. Some mature trees may reach diametersof 3 m, but most rainforest trees are relatively slendercompared with their height.

Also abundant in the rainforests are lianas, largewoody vines that entwine the trees, and epiphytes,plants that grow on the trunks and branches of otherplants. In one report, it was found that as many as60 percent of the trees in a Liberian rainforest carriedepiphytes, with as many as 45 to 65 species on a singletree. Although epiphytes do grow on other plants, theyare not parasites. Epiphytes are photoautotrophs that

15.5 Plant Biomes Reflect Myriad Physiological Adaptations 271

grow only in the illuminated portion of the canopy. Itis believed that they obtain water either directly fromrain and/or from the humid atmosphere of the canopy.Nutrients are obtained from atmospheric dust and thesurfaces of the plants on which they grow. Commonepiphytes include orchids, ferns, and bromeliads.

Soils of tropical rainforests typically retain very littlein the way of plant nutrients and are effectively infer-tile. But how can a forest that grows on such a poor,nutrient-free soil achieve such extraordinarily high lev-els of productivity and biomass? The answer to thisquestion is found in the rapid and thorough recyclingof nutrients. In the moist, warm climate of the rain-forest, litter is decomposed much more rapidly than itis in a comparable temperate forest. Large populationsof termites, ants, fungi, and other detritivores ensurethat the litter is rapidly broken down. The high averagetemperature, coupled with the absence of any cold sea-son, allows decomposers to work with high efficiencyyear-round. Rainforest trees also have a high capacity fornutrient uptake. The shallow, spreading root systems ofrainforest trees form extensive mycorrhizal associationsthat sweep the passing ground water clean of dissolvednutrient ions. There is some evidence that the mycor-rhizal fungi (Chapter 3) may also be in contact with thedecomposing litter, such that the transfer of nutrientsis immediate and direct. This means that, unlike in atemperate forest where the soil is the principal nutri-ent reservoir, the trees themselves represent the largestreserve of nutrients in a rainforest. When vegetation isshed through storm damage or natural senescence, itis rapidly decomposed and the nutrients captured forrecycling into new growth before they have much of achance to enter the soil.

15.5.2 EVAPOTRANSPIRATION IS AMAJOR CONTRIBUTOR TOWEATHER

In any heavily vegetated region, there is to be expecteda continual cycling of water between land surfaces(including vegetation) and the overlying atmosphere.The overall balance of land–atmosphere water exchangeis known as the hydrologic cycle. Rainwater is returnedto the atmosphere in the form of water vapor. Depend-ing on the amount of vegetation cover, transpirationmay be the principal source of water vapor (Chapter 2).In a rainforest, direct evaporation of soil water appearsto be negligible, in part because of the normally heavycanopy cover and in part because the litter on the for-est floor may act as a mulch to reduce evaporation ofsoil water. However, another significant source of watervapor is the direct evaporation of water interceptedby the canopy during a rainfall (Figure 15.7). Evapo-ration of intercepted rainwater occurs simultaneouslywith transpiration and it is not possible to distinguish

Condensation

Rainfall(100%)Evaporation

(25%)Transpiration

(50%)

Runoff (streams and groundwater)(25%)

Evapotranspiration

FIGURE 15.7 Rainfall and latent energy cycle in a tropicalrainforest. Fifty percent of the rainfall is returned to theatmosphere through transpiration. Another 25 percent isreturned by evaporation, primarily of water interceptedby the canopy, and 25 percent is accounted for as runoff.

between water vapors arising from the two sources. Forthis reason, the term evapotranspiration is used toidentify the transfer of water vapor from vegetated landsurfaces to the atmosphere, regardless of the source ofthe vapor.

Recycling of water through evapotranspiration isclearly a significant factor in the hydrologic cycle ofrainforests. In the case of the Amazon basin, for example,water vapor enters the region from the Atlantic Oceanand is driven inland by the predominantly easterly tradewinds. To this vapor is added at least an equal amountof vapor produced by evapotranspiration as the windssweep across the dense forests. Vapors from these twosources mix to form the clouds that eventually producerain across the basin.

The significance of evapotranspiration is not limitedto the water economy of a group of plants or its con-tribution of water vapor to the atmosphere. The latentheat flux (LE) that accompanies the evaporation andcondensation of water over tropical forests is an impor-tant factor in the redistribution of energy throughoutthat region of the atmosphere between ground level andan altitude of 5 to 10 miles (8 to 16 km), known as thetroposphere. The troposphere contains most of the

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moisture in the atmosphere and is therefore the regionwhere clouds and convection currents, or winds, arefound. Consequently, evapotranspiration has the poten-tial to influence both local climate and weather patternsthroughout the atmosphere. The basis for this influenceis the high positive heat exchange that occurs whenwater evaporates and condenses.

15.5.3 DESERT PERENNIALS AREADAPTED TO REDUCETRANSPIRATION AND HEATLOAD

The word desert2 often conjures up images of an aridregion with no plants, just a hot sun over barren, shiftingsands. Most deserts do receive some precipitation andare populated by a diverse and interesting plant lifethat is uniquely adapted to extremely dry conditions.In contrast to our usual image of a hot desert, colddeserts also exist on Earth. The Taylor Dry Valleyin Antarctica is the coldest, most arid, barren deserton Earth. Although no terrestrial plant life exists onthis desert, the biodiversity of microorganisms in theperennially ice covered lakes in this area of Antarctica isastounding.

Most deserts have one or, at most, two seasons ofpredictable rainfall interspersed with extended periodsof drought. Other deserts may receive a significant por-tion of the total annual precipitation as episodic rainfall,which comes as a single brief period of high-intensityrainfall. The result is that seasonality of precipitation,rather than mean annual rainfall values, has greater rel-evance to the distribution of desert vegetation and theirlife cycles. The most distinctive property of deserts isnot so much low precipitation per se, but that poten-tial evapotranspiration far exceeds rainfall on an annualbasis. Because deserts typically have long periods ofcloudless weather, the solar radiation load is high andmost of what little rainfall is received is rapidly lostthrough evaporation. As a result, there is very littleopportunity to store any significant amount of moisturein the soil.

The two principal constraints to plant survival ina desert environment are the availability of water andexcessive heat load. Availability of nutrients, with thepossible exception of nitrogen, is not normally a problemin desert soils. In order to cope with extended droughtinterspersed with only occasional periods of light rain-fall and high summer temperatures, desert perennials

2The term desert is an anthropocentric term referring to dryplaces that are usually hot for at least a portion of the year.Arid and semiarid are classifications based on annual rainfall.Arid regions receive less than 250 mm of rain annually andsemiarid regions between 250 and 450 mm.

have developed numerous morphological and metabolicadaptations that are designed either to conserve wateror to increase water-use efficiency and reduce heatload. Encelia farinosa (brittlebrush) is a C3 broadleaf,drought-deciduous shrub which withstands drought byexploiting seasonal leaf polymorphism. In response towinter rains, Encelia produces large, glabrous (smoothand hairless) leaves. As the dry season approaches andwater supply becomes increasingly tenuous, Encelia pro-duces progressively smaller, thicker leaves. In addition,both the adaxial and abaxial surfaces of the smaller leavesdevelop dense mats of dead, air-filled trichomes (epi-dermal hairs). This mat of trichomes gives the leavesa ‘‘white’’ appearance and increases the reflectance ofthe leaf. The early ‘‘green’’ leaves reflect approximately15 percent of the incident solar radiation, while thelate-season ‘‘white’’ leaves reflect up to 70 percent. Thisrepresents a significant reduction in the heat load onthe leaf, enough to reduce leaf temperature by as muchas 5 to 10◦C at midday in the summer. A reduced heatload serves to reduce transpiration and thus improvewater-use efficiency. It will also help to maintain leaftemperature in the optimum range for photosynthesis.These morphological changes thus enable Encelia andsimilar broadleaf desert species to resist drought stressand maintain a positive net photosynthesis well into thedry season. Sagebrush (Artemisia tridentata), like mostcold-desert shrubs, has a relatively high proportion ofits biomass invested in roots and probably occupies agreater volume of soil than most of its competitors.The high root–shoot ratio appears to be a means toensure efficient extraction of water from the soil whenit is recharged by precipitation during the winter/springseason.

Desert succulents such as members of the Cac-taceae (the true cacti) and their relatives in the familyEuphorbiaceae are often viewed as the quintessentialdesert plants. However, as succulents the cacti are nottrue xerophytes and are actually found more commonlyin semideserts and coastal temperate deserts. The cactiare stem-succulents: leaves have been reduced to thornsand photosynthesis is taken over by the stem. The thick,fleshy stems are composed of very large cells that storecopious amounts of water, and stomates are few andsunken. Another critical aspect is that they utilize cras-sulacean acid metabolism (CAM), a carbon-assimilationpathway that is inextricably tied to morphological suc-culence. Cacti and other succulents appear to reach theirgreatest development in the Sonoran, where they makeuse of the bimodal rainfall regime in order to main-tain their tissue water charge. The Sonoran desert, forexample, is where one finds the giant saguaro cactus(Carnegiea gigantea) and the organ-pipe cactus (Steno-cereus thurberi). Also prominent in the warm deserts areleaf-succulent members of the family Agavaceae, suchas Agave deserti. The succulent photosynthetic stems of

Summary 273

most cacti are oriented vertically rather than horizon-tally. Such stems are thus designed not only to conservewater but also to minimize heat load. An elongated, orcolumnar, shape presents a minimum surface area to thesun when it is overhead during midday.

15.5.4 DESERT ANNUALS AREEPHEMERAL

Annual plants are well represented in desert and semi-arid regions. What perennials there are tend to bedeep-rooted but widely spaced, so that they providerelatively little competition for annual species. Desertannuals are not xerophytic, but survive arid and semiaridconditions as dormant seed. Unlike perennials, whichrely on physiological and morphological adaptationsto help them survive periods of drought, desert annu-als must germinate, establish their entire biomass, andset seed each season. Their entire life cycle must becompleted during periods of adequate moisture and,because those wet period may be relatively short, theseannuals are often referred to as ephemerals. Not sur-prisingly, the life cycle of desert annuals is keyed toseasonal distribution of precipitation. Where precip-itation falls predominantly in the winter, the annualflora is active in the winter and early spring. In regionsexperiencing predominantly summer rains, the annualflora is predominantly summer-active. In those regionsof the Sonoran Desert that experience bimodal precip-itation, there are distinct winter- and summer-activeannual flora. Desert winter annuals tend to exhibit C3photosynthesis, reflecting the fact that winter/springtemperatures are lower and moisture is generally avail-able over a longer period of time. Summer annuals,on the other hand, tend to be C4 species, reflectingthe advantages offered by higher water-use efficiency inview of the short-lived water availability from summerrains.

The key to survival of desert annuals is a relativelyrapid life cycle. Their seeds, able to survive in the soilthrough extended periods of drought, will quickly ger-minate when adequate water is available during thenormal wet season. Desert annuals characteristicallyexhibit high growth rates, which allows them to reachmaturity, flower, and set seed within the relatively briefperiod of time that moisture is available. High growthrates are supported by at least three factors: a highshoot–root ratio, high stomatal conductance (support-ing rapid CO2 uptake), and high photosynthesis rates.An example is Cammisonia claviformis, a Mojave Desertwinter annual with relatively large leaves. Its roots areshallow, which allows them to absorb more efficientlylight rainfall and drippings from nighttime condensa-tion. Leaves tend to be amphistomatic (i.e., stomata arefound on both surfaces), which, accompanied by a high

stomatal conductance, supports a high rate of carbondioxide uptake.

This chapter has indicated that the inherent plas-ticity of plants is reflected in the myriad adaptationsto specific and, at times, extreme environments towhich plant communities are exposed. The adaptationsdiscussed not only have a significant impact on the bio-diversity and geographical distribution of plant biomesbut also influence local weather patterns.

SUMMARY

The physiological plasticity for acclimation to growthirradiance varies considerably between genotypes.Obligate shade adaptation appears to preclude accli-mation to high light whereas obligate sun adaptationappears to preclude acclimation to extreme low lightconditions. Plants with the C4 photosynthetic pathwayare adapted to hot, dry climates. C4 species evolved amechanism for avoiding the impact of photorespiratorycarbon dioxide loss by concentrating carbon dioxidein the carbon-fixing cells. C4 plants exhibit a divisionof labor between mesophyll cells, which pick up car-bon dioxide from the ambient air, and bundle-sheathcells, which contain the PCR cycle and actually fixcarbon. Mesophyll cells contain the enzyme PEPcarboxylase, which catalyzes the carboxylation of phos-phoenolpyruvate. The product C4 acid is transportedinto the bundle-sheath cell. There it is decarboxylatedand the resulting carbon dioxide is fixed via the PCRcycle. C4 plants have a very low carbon dioxide com-pensation concentration and low transpiration ratio.This means C4 plants are able to maintain higher ratesof photosynthesis at lower carbon dioxide levels, evenwhen the stomata are partially closed to conserve waterduring periods of water stress. Plants that exhibit cras-sulacean acid metabolism (CAM) are adapted to hot,desert conditions. CAM plants maintain higher ratesof photosynthesis in habitats with minimal access towater. CAM plants exhibit an inverted stomatal cycle,opening for carbon dioxide uptake at night (when waterstress is lower) and closing during the day (when waterstress is high). The carbon dioxide is stored as organicacids, again through the action of PEP carboxylase.Decarboxylation during the day furnishes the necessarycarbon dioxide for photosynthesis. The water use effi-ciency of CAM is higher than that of C4 plants which,in turn, is higher than that of C3 plants. The differ-ence in water use efficiency is an important factor thatdetermines the geographical distribution of C3, C4 andCAM plants.

Plant biomes reflect myriad adaptations of plantcommunities to specific environments. Althoughthe soils of tropical rainforests are extremely poorin nutrients, these biomes exhibit the greatest

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plant biodiversity. Since water is not limiting inrainforest biomes, the extent of evapotranspirationfrom tropical rainforest biomes is so significantthat it affects weather patterns. Desert perennialsare adapted to maximize photosynthesis but at thesame time minimize heat load and water loss underarid conditions. They exhibit rather slow rates ofvegetative growth. In contrast to desert perennials,desert annuals are ephemeral. They exploit short,episodic rainfall to germinate, grow rapidly, sene-scence, and produce seed to survive the next dry spell.

CHAPTER REVIEW

1. Define adaptation. List five traits that areassociated with shade adaptation.

2. Trace the path of carbon in a typical C4 leaf, fromentry into the leaf through the stomatato its export in the vascular tissue. Howdoes this differ from the C3 pathway?

3. A distinctive feature of the bundle-sheath cells in atypical C4 leaf is a high density of chloroplasts.What advantage does this offer? Why is itadvantageous to have the PCR cycle locatedin the bundle-sheath cells in a C4 leaf?

4. Review the ecological significance of the C4 cycle.In what situations can a C3 species be more pro-ductive than a C4 species?

5. In what significant way does crassulacean acidmetabolism (CAM) differ from C4 metabolism?

6. Under which conditions would C3 plantsoutcompete C4 and CAM plants? Why is this so?

7. Define biome. How does the extent of evap-otranspiration affect weather patterns?

8. Would you predict that a tropical rainforest has agreater proportion of C3 versus C4 species?Explain why this would be so.

9. How do adaptations of perennial desert plants toarid conditions differ from those of desert annuals?

FURTHER READING

Anderson, J. M. 1986. Photoregulation of the composition,function and structure of thylakoid membranes. AnnualReview of Plant Physiology 37:93–136.

Buchanan, B. B., W. Gruissem, R. L. Jones. 2000. Biochem-istry and Molecular Biology of Plants. Rockville MD: Amer-ican Society of Plant Physiologists.

Bush, M. B., J. R. Flenley. 2007. Tropical Rainforest Responsesto Climate Change. Berlin: Springer.

Cushman, J. C., H. J. Bohnert. 1999. Crassulacean acidmetabolism: Molecular genetics. Annual Review of PlantBiology 50:305–332.

Edwards, G. E., V. R. Franceschi, E. V. Voznesenskaya.2004. Single-cell C4 photosynthesis versus the dual-cell(Kranz) paradigm. Annual Review of Plant Biology55:173–196.

Leegood, R. C., T. D. Sharkey, S. von Caemmerer. 2000.Photosynthesis: Physiology and Metabolism. Advances in Pho-tosynthesis, Vol. 9. Dordrecht: Kluwer.

Matsuoka, M., R. T. Furbank, H. Fukayama, M. Miyao. 2001.Molecular engineering of C4 photosynthesis. AnnualReview of Plant Biology 52:297–314.

Nobel, P. S. 2005. Physicochemical and Environmental PlantPhysiology. Burlington: Elsevier Science & Technology.

Potters, G., T. P. Pasternak, Y. Guisez, K. J. Palme, M.A. K. Jansen. 2007. Stress-induced morphogenicresponses: Growing out of trouble? Trends in PlantScience 12:98–105.

Robichaux, R. H. 1999. Ecology of Sonoran Desert Plants andPlant Communities. Tuscon: University of Arizona Press.

Sack, L., N. M. Holbrook. 2006. Leaf hydraulics. AnnualReview of Plant Biology 57:361–381.

Smith, S. D., R. K. Monson, J. E. Anderson. 1997. Physi-ological Ecology of North American Desert Plants. Berlin:Springer Verlag.

16Development: An Overview

The development of a mature plant from a single fertil-ized egg follows a precise and highly ordered successionof events. The fertilized egg cell, or zygote, divides,grows, and differentiates into increasingly complex tis-sues and organs. In the end, these events give rise tothe complex organization of a mature plant that flowers,bears fruit, senesces, and eventually dies. These events,along with their underlying genetic programs and bio-chemistry, and the many other factors that contribute toan orderly progression through the life cycle, constitutedevelopment. Understanding development is one of themajor goals of plant biologists and looking for changesin the pattern of development is one way to enhancethat understanding

In this chapter we highlight major stages in plantdevelopment, including

• the meaning of the terms growth, differentiation,and development,

• the nature of plant meristems,• the development, maturation, and germination of

seeds,• the pattern of development from embryo to adult,

and• senescence and programmed cell death.

16.1 GROWTH,DIFFERENTIATION,AND DEVELOPMENT

Three terms routinely used to describe various changesthat a plant undergoes during its life cycle are growth,differentiation, and development. The chapters thatfollow focus on factors that regulate growth, differenti-ation, and development, so it is necessary to clarify howthese three terms are used.

16.1.1 DEVELOPMENT ISTHE SUM OF GROWTHAND DIFFERENTIATION

Development is an umbrella term, referring to thesum of all of the changes that a cell, tissue, organ,or organism goes through in its life cycle. Developmentis most visibly manifested as changes in the form of anorgan or organism, such as the transition from embryoto seedling, from a leaf primordium to a fully expandedleaf, or from the production of vegetative organs to theproduction of floral structures.

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16.1.2 GROWTH IS AN IRREVERSIBLEINCREASE IN SIZE

Growth is a quantitative term, related only to changes insize and mass. For cells, growth is simply an irreversibleincrease in volume. For tissues and organs, growthnormally reflects an increase in both cell number andcell size.

Growth can be assessed by a variety of quantitativemeasures. Growth of cells such as bacteria or algaein culture, for example, is commonly measured as thefresh weight or packed cell volume in a centrifuge tube.For the angiosperms and other so-called higher plants,however, fresh weight is not always a reliable measure.Most plant tissues are approximately 80 percent water,but water content is highly variable and fresh weight willfluctuate widely with changes in ambient moisture andthe water status of the plant. Dry weight, determinedafter drying the material to a constant weight, is ameasure of the amount of protoplasm or dry matter(i.e., everything but the water). Dry weight is usedmore often than fresh weight, but even dry weightcan be misleading as a measure of growth in certainsituations. Consider the example of a pea seed that isgerminated in darkness. In darkness, the embryo in theseed will begin to grow and produce a shoot axis that mayreach 10 to 12 inches in length. Although we intuitivelysense that considerable growth has occurred, the totaldry weight of the seedling plus the seed will actuallydecrease compared with the dry weight of the seedalone prior to germination. The dry weight decreasesin this case because some of the carbon stored in therespiring seed is lost as carbon dioxide. In the light,this lost carbon would be replaced and even augmentedby photosynthesis but photosynthesis doesn’t operatein darkness. In a situation such as this, either freshweight or the length of the seedling axis would be abetter measure of growth. Length, and perhaps width,would also be suitable measures for an expanding leaf.Length and width would not only provide a measureof the amount of growth, but a length-to-width ratiowould also provide information about the pattern of leafgrowth.

It should be obvious that many parameters could beinvoked to measure growth, dependent to some extenton the needs of the observer. Whatever the measure,however, all attempts to quantify growth reflect a fun-damental understanding that growth is an irreversibleincrease in volume or size.

While cell division and cell enlargement normallygo hand-in-hand, it is important to keep in mind thatthey are separate events; growth can occur without celldivision and cell division can occur without growth.For example, cell division is normally completed veryearly in the development of grass coleoptiles and thesubstantial elongation of the organ that follows is duealmost entirely to cell enlargement. Years ago, it was

shown that wheat (Triticum sp.) seeds could be made togerminate even after having been irradiated with gammaradiation sufficient to block both DNA synthesis andcell division. The result was a small seedling producedby cell enlargement alone. Such seedlings generally didnot survive more than two or three weeks but, exceptfor having abnormally large cells, their morphology wasmore or less normal. On the other hand, cell divisioncan also proceed without cell enlargement. Duringthe early stages of embryo development in flowers, forexample, a portion of the embryo sac goes througha stage in which cell division continues to produce alarger number of increasingly smaller cells, with nooverall increase in the size of the embryo sac.

16.1.3 DIFFERENTIATION REFERSTO QUALITATIVE CHANGESTHAT NORMALLY ACCOMPANYGROWTH

Differentiation refers to differences, other than size,that arise among cells, tissues, and organs. Differen-tiation occurs when cells assume different anatomicalcharacteristics and functions, or form patterns. Differen-tiation begins in the earliest stages of development, suchas, when division of the zygote gives rise to cells that aredestined to become either root or shoot. Later, unspe-cialized parenchyma cells may differentiate into morespecialized cells such as xylem vessels or phloem sievetubes, each with a distinct morphology and unique func-tion. Differentiation does not lend itself easily to quan-titative interpretation but may be described as a seriesof qualitative, rather than quantitative, changes. Finally,although growth and differentiation are normally con-current events, examples abound of growth withoutdifferentiation and differentiation without growth.

Differentiation is a two-way street and is not deter-mined so much by cell lineage as by cell position withrespect to neighboring cells. Thus, even though someplant cells may appear to be highly differentiated orspecialized, they may often be stimulated to revertto a more embryonic form. For example, cells iso-lated from the center of a tobacco stem or a soybeancotyledon and cultured on an artificial medium may bestimulated to reinitiate cell division, to grow as undif-ferentiated callus tissue, and eventually to give rise toa new plant (Figure 16.1). It is as though the cellshave been genetically reprogrammed, allowing them toreverse the differentiation process and to differentiatealong a new and different path. This ability of dif-ferentiated cells to revert to the embryonic state andform new patterns without an intervening reproduc-tive stage is called totipotency. Most living plant cellsare totipotent—something akin to mammalian stemcells—and retain a complete genetic program eventhough not all of the information is used by the cell

16.2 Meristems are Centers of Plant Growth 277

FIGURE 16.1 Shoot regeneration in callus culture. A pieceof pith tissue from the center of a tobacco stem wasexplanted onto a medium containing mineral salts, vita-mins, sucrose, and hormones. The tissue proliferated asan undifferentiated callus (left) for several weeks beforeregenerating new plantlets (right). These plantlets canbe eventually planted into soil and will produce a maturetobacco plant.

at any given time. In this sense, development does notreflect a progressive loss of genetic information, onlythe selective use of that information in order to achieveparticular developmental ends.

Not all cells are totipotent. Highly specialized cellswhose development has been locked in, such as by excep-tionally thick and rigid secondary cell walls or severelymodified protoplasts, are not capable of renewed differ-entiation. On the other hand, it is probable that all tissuescontain at least some potentially totipotent cells—cellsthat have the morphogenetic potential of a zygote. Plantdevelopment proceeds in an orderly fashion because thatpotential is carefully limited. When those limitations areremoved, totipotent cells simply revert to the zygoticstate and begin the developmental program anew.

There is increasing evidence that the genetic pro-gram being read in a particular cell may also dependon the position of that cell with respect to other cellsand tissues and the inputs it receives from its neighbors.The position of a cell determines its interaction with itsneighbors as well as its relationship to nutrient and hor-mone gradients. The importance of position has recentlybeen demonstrated in a study of young Arabidopsis roots.Arabidopsis roots lend themselves well to this kind ofstudy because of their relatively simple structure. Theroot tip is comprised of a single layer each of epidermis,cortex, endodermis, and pericycle surrounding a vascu-lar bundle. This simplified organization makes it easierto track individual cells than is possible in more complexroots with multiple cell layers. Root cells were labeledwith marker genes that were expressed differently invascular and root cap cells and then a laser microscopewas used to surgically remove cells in a region adjacentto the root cap called the quiescent center. The deadcells were displaced toward the root tip and replaced by

daughters of adjacent vascular cells. In their new posi-tion, the former vascular cells expressed the root capmarker. This is a strong indication that the reading ofthe genetic program in a cell is at least to some extentdetermined by positional information rather than celllineage.

16.2 MERISTEMS ARE CENTERSOF PLANT GROWTH

Unlike animals, which are characterized by a general-ized growth pattern, plant growth is limited to discreteregions where the cells retain the capacity for contin-ued cell division. These regions are called meristems(Gk. merizeim, to divide). Two such regions are theapical meristems located at the tips of roots and stems.These regions of active cell division are responsible forprimary growth, or the increase in the length of rootsand stems.

The tip of the root is covered by a root cap, whichprovides mechanical protection for the meristem asthe root grows through the abrasive soil medium. Theroot cap also secretes polysaccharides, which form amucilaginous matrix called mucigel. Mucigel lubricatesthe root tip as it moves through the soil. The rootcap along with its coating of mucigel is also involvedin perception of gravity by roots (Chapter 23). Theroot apical meristem (RAM) is a cluster of dividingcells located at the tip of the root just behind the rootcap (Figure 16.2). Each time a cell in the meristemdivides, one of the two daughter cells will be retainedto continue cell division while the second daughter cellproceeds to elongate, thus increasing the length of theroot and pushing the root tip through the soil. In thecenter of the meristem is a region of slowly dividing cellscalled the quiescent zone. Cell divisions responsible fornew tissues in the elongation root and regeneration ofthe root cap take place around the periphery of thequiescent zone.

The shoot apical meristem (SAM) is structurallymore complex than the root apical meristem. This isunderstandable because in addition to producing newcells that elongate and extend the length of the axis ofthe shoot, the shoot apical meristem must also formprimordia that give rise to lateral organs such as leaves,branches, and floral parts. At the same time it mustperpetuate itself by maintaining a small population ofundifferentiated, dividing cells. Similar to the root apicalmeristem, each time a cell divides in the SAM, onedaughter cell is left behind to elongate and move theshoot apex forward while the other daughter cell remainswithin the meristem to continue dividing.

The meristem of a typical dicot shoot is a small,shiny dome that can be seen in detail with the aid ofan electron microscope (Figure 16.3). A microscopic

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FIGURE 16.2 Schematic diagram of a young Ara-bidopsis root tip showing the principal regions ofthe root apical meristem.

Epidermis

Cortex

Stele

Endodermis

Quiescent center

Lateral rootcap

Columella

examination of a typical SAM reveals that the cells ofthe meristem are usually organized into two distinctregions. Cells in the outermost two to four layers,called the tunica, undergo only anticlinal divisions, ordivisions perpendicular to the surface of the meristem.The tunica thus contributes only to surface growth andits products contribute to the peripheral tissues of thestem. Underlying the tunica is the corpus, a body ofcells that divide in various planes and contribute to thebulk of the shoot.

One of the ongoing mysteries of plant developmentis why and how leaves and branches form where theydo. Part of the reason for this uncertainty is that a lotof things happen at the apex in rapid succession andit is difficult to separate these various events. Newer

methods for the study of gene expression in situ,however, are beginning to provide some insights as towhich genes appear to be turned on and which genesare turned off when the lateral appendages begin todevelop (see Chapter 20).

Leaves form as small swellings or primordia on thelateral flanks of the meristem. In some plants a pairof leaf primordia will arise on opposite sides of themeristem with successive pairs arising at 90 degrees tothe last. In other plants, the leaf primordia arise in aspiral pattern. The points at which the leaf primordiaform are referred to as nodes. In its early stages, the leafprimordium develops as a peg-like extension. As a leafprimordium elongates, however, marginal meristemsdevelop on opposite sides of the primordium, leading

Ax

FB

Lp

A. B.

FIGURE 16.3 Shoot apical meristem. (A) Scanning electron micrograph of a flow-ering meristem of Brassica napus. Ax, apex; Lp, leaf primordium; FB floral bud (B).Schematic cross-section of a shoot apical meristem, showing position of the tunica,corpus, and leaf primordia. (A from Sawhney, V. K., P. L. Polowick. 1986. AmericanJournal of Botany 73:254–263. With permission of the American Journal of Botany.Original photograph kindly provided by V. K. Sawhney and P. L. Polowick.)

16.3 Seed Development and Germination 279

to lateral growth that gives rise to the typical flattenedblade. As each leaf primordium forms, a bud primordiumis also formed in the axil where the leaf joins the stem.These axillary buds will eventually give rise to branches.While all this has been going on, cells in the regionsbetween successive leaves, or the internodes, continueto divide and elongate, thereby increasing the length ofthe stem.

Tissues that are derived directly from the rootand shoot apical meristems are called primary tissues.The stems and roots of woody plants, however, growin diameter as well. An increase in diameter resultsfrom the activity of a meristem called the vascularcambium. Tissues laid down by the vascular cambiumare called secondary tissues, so the vascular cambium isresponsible for secondary growth. The primary tissueof roots and shoots contains a central core of vascular,or conducting, elements. Characteristically, the xylemlies toward the center of the vascular core and thephloem lies at the outer edge of the core (Figure 16.4).The vascular cambium develops between the xylem andphloem and produces new xylem toward the inside andnew phloem toward the outside. Because of its heavycell walls and eventual lignification, xylem is a rigid andlong-lasting tissue that eventually occupies the bulk ofmost woody stems or trunks. Phloem is a more fragiletissue and with each year’s new growth the previousyear’s cells tend to be pushed outward and crushed. Asa result, the xylem continues to transport water andminerals for several years, but a large tree seldom hasmore than one year’s worth of functioning phloem.

Epidermis

Cortex

Primary phloem

Secondary phloemVascular cambium

Secondary xylem

Primary xylem

Pith

FIGURE 16.4 A schematic transverse section through aone-year-old elderberry (Sambucus) stem showing thelocation of the secondary vascular cambium. The vas-cular cambium arises between the primary phloem andprimary xylem and adds new, or secondary, xylem cellsto the inside and new, or secondary, phloem cells to theoutside (arrows). This is repeated annually to add girthto the stem. The secondary xylem develops heavy, lig-nified walls and becomes the woody tissue of the stem.The phloem is a soft tissue and each year’s new growthcrushes the previous year’s phloem. When you peel the‘‘bark’’ off a young stem, the vascular cambium is wherethe tissues separate.

16.3 SEED DEVELOPMENTAND GERMINATION

The life of an individual plant begins when an eggnucleus in the maternal organs of a flower is fertilizedby a sperm nucleus to form a zygote. Growth anddifferentiation of the zygote produces an embryo con-tained within a protective structure called a seed. Underappropriate conditions, the embryo within the seed willrenew its growth and will continue to develop into amature plant.

16.3.1 SEEDS ARE FORMEDIN THE FLOWER

Flowers appear to vary enormously in structure, yet allflowers follow the same basic plan. A generic flowerconsists of four whorls or circles. The two outermostwhorls—the sepals and petals—are vegetative struc-tures; and the two innermost—the stamens and pistil—are the male and female reproductive structures, respec-tively. At the base of the pistil, or female structure, is theovary, which contains one or more ovules (Figure 16.5).

Within each ovule, a single large diploid cell, calledthe megaspore mother cell, undergoes mitosis to pro-duce four megaspore cells. Only one megaspore cellsurvives and that cell undergoes meiotic division toproduce an embryo sac with eight haploid nuclei. Sub-sequent cell division produces a mature embryo sac inwhich the eight nuclei are segregated into seven cells(Figure 16.6). One of those cells is the egg. Another isthe large central cell containing two polar nuclei.

The male structures, or stamens, surround the pistiland consist of an anther perched on a stalk, or filament.

StigmaStyleOvary

Receptacle

Pedicel

Sepal

Petal

Filament

AntherStamen

Pistil

FIGURE 16.5 A typical perfect flower consists of 4 whorls:sepals, petals, stamens, and the pistil. All four whorls areconsidered modified leaves. The sepals are often greenor inconspicuous and the petals are brightly colored.In some flowers, the sepals and petals may both be col-ored. Pollen, containing the sperm nucleus, is producedin the anthers of the stamens. The female egg cells areproduced in the ovary at the base of the pistil. Pollen istransferred to the stigma or stigmatic surface of the pis-til, where it sends out a pollen tube that grows down thestyle and delivers the sperm nucleus to the egg.

280 Chapter 16 / Development: An Overview

Polar nuclei

Egg cell

Integuments

Micropyle

Stalk

FIGURE 16.6 Diagram of a mature eight-nucleate,seven-celled embryo sac. The pollen tube delivers thesperm nuclei through the micropyle, or space betweenthe integuments. The stalk is in effect a placenta thatanchors the ovule to the ovary wall.

The anther contains a large number of microsporemother cells, each of which undergoes meiotic divisionto form uninucleate, single-celled microspores. Themicrospores subsequently become encased in heavy,resistant outer walls and the nucleus divides mitoti-cally, forming two cells—a tube cell and a generativecell—within the original spore wall. This is the maturepollen grain (Figure 16.7).

Mature pollen grains are shed from the anthersand carried to the stigmatic surface of the pistil byinsects, wind, or some other vector. Once the pollengrain lands on the stigmatic surface—an event calledpollination—the pollen grain takes up water and sendsout a pollen tube that grows down the style of the pistiltoward the ovule (Figure 16.7 A, C). The tube nucleusmigrates down the pollen tube and appears to direct itsgrowth. The cell wall of the generative cell breaks downand the generative nucleus divides once to form twosperm nuclei that follow the tube nucleus down the tubeas it elongates. Pollen tube growth toward the embryosac requires a calcium gradient, but the precise signalingmechanism remains uncharacterized.

In the final stage, the elongating pollen tube entersthe ovule by growing through the micropyle (the spacebetween the ends of the surrounding integuments) andreleases the two sperm nuclei into the embryo sac. Ulti-mately, one of the two sperm nuclei enters the egg celland fertilizes the egg cell nucleus to form the zygote.The second sperm nucleus enters the large central celland fuses with the two polar nuclei to form a triploidendosperm nucleus. The endosperm nucleus will goon to form the primary nutritive tissue, or endosperm,for the developing embryo. The involvement of two

sperm nuclei in this way is called double fertiliza-tion, a characteristic unique to the flowering plants orangiosperms.

16.3.2 SEED DEVELOPMENTAND MATURATION

Seeds come in various guises. To the plant ecologist, aseed is a propagule or device for effecting propagationof the individual or population. To the morphologist aseed is more rigorously defined as a mature ovule. Thephysiologist, however, views a seed as a mature embryo

Sperm nuclei

Tube nucleus

Generative cell

Tube cell nucleus

A.

B. C.

FIGURE 16.7 (A) Pollen grains and pollen tubes. Maize(Zea mays) pollen grains were germinated on an agar sur-face. Elongating pollen tubes are indicated by the arrows.(Photograph courtesy of D. B. Walden). (B) Diagram of amature pollen grain showing the tube cell and generativecell. The diploid generative nucleus divides meioticallyto produce two haploid sperm nuclei. (C) A germinatingpollen grain. The pollen tube grows down the style of thepistil and delivers the sperm nuclei to the embryo sac.

16.3 Seed Development and Germination 281

surrounded by nutritive tissues and encased in a protec-tive seed coat. It is fair to say that germinating seeds andyoung seedlings have been a favored system for studyingvegetative development since the time of Darwin andbefore. Treating seeds with chemical mutagens is nowone of the most convenient systems for inducing muta-tions that provide clues about developmental processes(Box 16.1).

The development of a seed begins with the fertilizedovule, or zygote. The early stage of seed developmentis characterized by extensive cell divisions that form theembryo and, in endospermic seeds, the tissues that storenutrients that will support the eventual germination ofthe seed and seedling development.

The first division of the zygote is usually transverseand immediately establishes polarity of the embryo.The upper cell is destined to become the embryo itselfwhile the lower cell produces a stalk-like suspensor thatanchors the embryo at the base of the embryo sac. Thetypical dicot seed will then pass through several recog-nizable stages (Figure 16.8). During the early stages ofembryo development, cell division occurs throughoutthe entire cell mass but at the heart-shape stage boththe shoot and root apical meristems begin to organizeas centers of cell division.

Throughout the development of the embryo, thereis a continuous flow of nutrients from the parent plantinto the endosperm or the cotyledons. In some cases,such as the cereal grains and most other monocots,

Apical meristem

CotyledonsProtoderm

Suspensor

Radicle

Primary vasculartissue

A. B. C.

FIGURE 16.8 Major stages in the development of a dicotembryo. (A) The globular stage. The original zygoteundergoes numerous divisions to produce a suspensorand the 64-cell globular embryo. The protoderm is des-tined to become the epidermis in the mature embryo.The stalk-like suspensor anchors the embryo at thebase of the embryo sac and assists in the absorption ofnutrients from the surrounding endosperm. (B) Theheart-shaped stage. Cells destined to become the apicalmeristem begin to organize at the base of the emerg-ing cotyledons. (C) The torpedo stage. The embryo isno longer attached by the suspensor, the shoot-root axisis now clear, the apical meristem is beginning to func-tion, and the primary vascular tissue (xylem and phloem)begins to organize. Cellular detail is not shown in (C) inorder to emphasize the difference in scale.

the endosperm is retained until maturity and may com-prise the bulk of the seed (Figure 16.9 A, B). These arecalled endospermic seeds. The endosperm of matureendospermic seeds consists of cells filled with starchalong with protein and some small amounts of lipid.In some monocot seeds, most notably the cereal grainssuch as Triticum (wheat), Hordeum (barley), and Avena(oats), the endosperm is surrounded by one or moredistinctive layers of cells, called the aleurone. Aleuronecells are distinguished by the presence of numerousprotein bodies and are the source of enzymes neededto mobilize nutrients during germination. Endosper-mic dicot seeds, such as castor bean (Ricinus communis),have retained a significant amount of endosperm and atmaturity the cotyledons are thin, leaflike structures. Innonendospermic dicot seeds, such as Pisum (pea) andPhaseolus (bean), the cotyledons enlarge at the expense ofthe endosperm and may occupy as much as 90 percentof the seed volume at maturity (Figure 16.9C). Bothendosperm and cotyledons contain large quantities ofstored carbon (in the form of carbohydrates, lipids, andprotein), mineral elements, and hormones that supportthe growth and development of the seedling until it canestablish itself as a photosynthetically competent plant.

The late, or maturation, stage of seed developmentis characterized by cessation of embryo growth and thedevelopment of desiccation resistance. Maturation isterminated by a dramatic desiccation in which the watercontent of the seed is reduced from 80 or 90 percentto approximately 5 percent. Surrounding the matureseed is a hard coat derived from maternal tissues (theinteguments) which surrounded the seed during itsdevelopment in the ovary. Comprised of heavy-walledcells and covered with a thick, waxy cuticle, the seedcoat often presents a significant barrier to the uptake ofboth water and oxygen by the seed.

16.3.3 SEED GERMINATION

Because seeds are severely dehydrated, any metabolicreactions take place so slowly they are scarcelydetectable. Seeds are thus quiescent, or resting, organsthat represent a normal hiatus in the life cycle of aplant. The embryo appears to be in a state of suspendedanimation, capable in some cases of surviving adverseconditions for long periods of time. Resumption ofembryo growth, called germination, is dependentupon a number of factors, but three are especiallyimportant: adequate water to re-hydrate the tissues, thepresence of oxygen to support aerobic respiration, anda ‘‘physiological’’ temperature. Although many seedswill germinate over a wide range of temperatures, theoptimum range for most seeds is 25◦C to 45◦C. Thelower limit is highly variable, depending on the species.The upper limit generally reflects the temperaturewhich denatures proteins.

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BOX 16.1DEVELOPMENT INA MUTANT WEED

Many aspects of plant development have proven dif-ficult to dissect using exclusively physiological andbiochemical approaches. This is especially true of thecomplex metabolic sequences that connect a develop-mental event with the original signal that initiates thatevent. Metabolic events associated with a signal trans-duction pathway often represent a very small portion ofall the biochemical processes within a plant and may notbe discernible against this background of biochemical‘‘noise.’’

Because all development can be traced back to theexpression of genes, one way around this dilemma isto incorporate genetic mutations into a research pro-gram. The use of mutant genes to study metabolism anddevelopment has been a constructive approach since thepioneering work of G. Beadle and E. Tatum with Neu-rospora mutants in the 1940s. Over the years, mutantsin maize, tomato, pea, and a host of other plants haveprovided important insights into normal development.More recently, however, recombinant DNA techniqueshave enabled us to ask questions about genes and devel-opmental events in ways never before possible. Theprincipal strategy is to identify and genetically mapa mutation that modifies the physiological response ofinterest. It is then possible to physically isolate and clonethe wildtype gene and, based on its nucleotide sequence,deduce the amino acid sequence of the encoded proteinthat normally operates in the pathway. This providesinvestigators with useful information about the functionof the wildtype gene and a means to further probe thestep that it controls. Furthermore, by studying interac-tions between mutant genes and multiple input signals,

it is possible to dissect complex interactions betweendifferent pathways.

One organism that has come to dominate this newapproach to the study of plant development is Arabidop-sis thaliana, a member of the Brassicaceae, or mustardfamily. A. thaliana (mouse-ear cress) is a small, herba-ceous weed that grows in dry fields and along roadsidesthroughout the temperate regions of the northern hemi-sphere. Most Arabidopsis are winter annuals. Their seedsgerminate in the fall, forming an overwintering vegeta-tive rosette. In the spring, as the days grow longer, thestem elongates and flowers.

There are many reasons why Arabidopsis has becomesuch a popular experimental tool and model system formolecular genetic experiments on development. Ara-bidopsis is easily grown in the laboratory and its life cycleis complete in 5 to 6 weeks. It is also easily crossedor self-fertilized and produces prodigious numbers ofseeds (up to 10,000 per plant). The Arabidopsis genome,which has recently been completely sequenced, is one ofthe smallest known plant genomes (approximately 108

nucleotides, or 25,498 genes). Finally, mutants are easilyinduced by treating the seeds with chemical mutagens.The surviving seeds are then germinated and mutantprogeny are recovered for analysis. The small size ofthe plant together with its rapid growth and fecun-dity make it easy to screen for mutants with reasonablefrequency.

One must always be cautious with so-called modelsystems. What is learned about Arabidopsis, for example,may well extrapolate to all brassicas, but not necessarilyto all plants. Nonetheless, Arabidopsis is an ideal plantfor doing molecular genetic and developmental experi-ments. The isolation and study of mutants in Arabidopsishas already made significant contributions to many areasof plant development. Moreover, what is learned aboutphysiology and development in Arabidopsis gives inves-tigators important clues about what to look for in otherplants.

The initial step in germination of seeds is the uptakeof water and rehydration of the seed tissues by the pro-cess of imbibition. Like osmosis, imbibition involvesthe movement of water down a water potential gradient.Imbibition differs from osmosis, however, in that it doesnot require the presence of a differentially permeablemembrane and is driven primarily by surface-acting ormatric forces. In other words, imbibition involves thechemical and electrostatic attraction of water to cellwalls, proteins, and other hydrophilic cellular materi-als. Matric potential, like osmotic potential, is alwaysnegative. Hydration causes a swelling of the imbibingmaterial, which may generate substantial forces (called

imbibition pressure). Imbibition pressure developedby a germinating seed will cause the seed coat to rupture,thus permitting the embryo to emerge.

One of the first events to follow the rapid influxof water is the leakage of low molecular weight solutesfrom the seed tissues. This apparently reflects an unsta-ble configuration of cell membranes related to the severedesiccation. The leakage does not persist, however, asthe cells quickly repair themselves and a more sta-ble membrane configuration returns shortly after rehy-dration.

Imbibition of water is followed by a general acti-vation of seed metabolism within minutes of water

16.3 Seed Development and Germination 283

Endosperm

Scutellum(Cotyledon)

Primary leaves

Coleoptile

Mesocotyl

Radicle

Coleorhiza

Pericarp

A.

B.

C.

FIGURE 16.9 (A) Maize (corn, (Zea mays)) kernel showingthe location of the embryo underneath the pericarp. Eachcorn grain is actually an entire fruit and the seed coat hasfused with the surrounding ovary wall to form a com-pound structure called the pericarp. (B) Cross section ofa mature maize kernel (along the dotted line in A) show-ing the principal structures of the embryo. (C) A beanseed (Phaseolus vulgaris) showing the embryo and partof one cotyledon. The seed coat and one cotyledon havebeen removed in order to reveal the embryo.

entering the cells, initially utilizing a few mitochon-dria and respiratory enzymes that had been conserved inthe dehydrated state. Renewed protein synthesis is alsoan early event, utilizing preexisting RNA transcripts andribosomes, as existing organelles are repaired and neworganelles are formed. This is followed closely by (1) therelease of hydrolytic enzymes that digest and mobilizethe stored reserves, and (2) renewed cell division and cellenlargement in the embryonic axis. Detailed respiratorypathways have been studied thoroughly in only a fewspecies of seeds, but it is believed that glycolysis and thecitric acid cycle are active to varying degrees in most, ifnot all, seeds. These pathways produce the carbon skele-tons and ATP that are required to support growth anddevelopment of the embryo. The pentose phosphatepathway is also important in seeds as it produces thereducing potential (in the form NADPH) required forthe reductive synthesis of fatty acids and other essentialcellular constituents. The pentose phosphate pathwayalso generates intermediates in the synthesis of aro-matic compounds and perhaps nucleic acids. Seeds thatstore carbon reserves principally in the form of fats andoils will carry out the synthesis of hexose sugars viagluconeogenesis.

The mobilization of stored carbon in seeds has beenmost extensively studied in cereals (see Chapter 19). Thisis because the endosperm of cereals has long served as aprincipal source of nutrition for man and domesticatedanimals, as well as a basic feedstock in the brewingindustries. These needs have provided a strong incentivefor research into the mobilization of starch reserves incereal grains. In nonendospermic dicot seed such asthe legumes (peas, beans), the initial stages of radicleelongation appear to depend on reserves stored in thetissues of the radicle itself. Later, carbon reserves aremobilized from the cotyledons and transported to theelongating axis.

In most species, germination is considered com-plete when the radicle emerges from the seed coat.Radicle emergence occurs through a combination ofcell enlargement within the radicle itself and imbibitionpressures developed within the seed. Rupture of theseed coat and protrusion of the radicle allows it to makedirect contact with water and nutrient salts required tosupport further growth of the young seedling.

16.3.4 THE LEVEL AND ACTIVITIESOF VARIOUS HORMONESCHANGE DRAMATICALLYDURING SEED DEVELOPMENT

Seed development is characterized by often dramaticchanges in the levels of the principal plant hormones(Figure 16.10). In most seeds, cytokinin (CK) levels arehighest during the very early stages of embryo develop-ment when the rate of cell division is also highest. As the

284 Chapter 16 / Development: An Overview

H2O

Early Mid Late

Nutrientreserves

CK

GA,IAAGA

ABAIAA

QU

IES

CE

NC

E

Seed DevelopmentGermination and

Seedling Development

FIGURE 16.10 The activity of hormones during seeddevelopment and germination. Cytokinins (CK) arepresent during the early stages of development, whencell division is most active. The concentration ofcytokinins then declines while the concentrations ofgibberellin (GA) and auxin (IAA) increase to supportactive cell enlargement. Abscisic acid (ABA) concentra-tion increases in the later stages to prevent precociousgermination. After a quiescent period, the release of gib-berellins activates nutrient mobilization and IAA levelsincrease to stimulate cellular enlargement in the youngseedling.

cytokinin levels decline and the embryo enters a periodof rapid cell enlargement and differentiation, both auxinand gibberellin (GA) levels increase. In the early stagesof embryogenesis, there is little or no detectable abscisicacid (ABA). It is during the latter stages of embryodevelopment, as GA and IAA levels begin to decline,that ABA levels begin to rise. ABA levels generally peakduring the maturation stage when seed volume and dryweight also reach a maximum. The significance of thesechanges will become more apparent as we look at thespecific effects of individual hormones in later chapters.Maturation of the embryo is characterized by cessationof embryo growth, accumulation of nutrient reserves,and the development of tolerance to desiccation.

16.3.5 MANY SEEDS HAVE ADDITIONALREQUIREMENTS FORGERMINATION

Many seeds will not germinate even though the minimalenvironmental conditions have been met. These seedsare said to be dormant and will not germinate untiladditional conditions have been met.

The most common causes of seed dormancy arethe impermeability of the seed coat to water or oxygenor physiological immaturity of the embryo at the timethe seed is shed from the mother plant. Immature seedsmust undergo complex biochemical changes, collectivelyknown as after-ripening, before they will germinate.After-ripening is usually triggered by low temperature,a mechanism that appears to ensure that the seed willnot germinate precociously in the fall but will germinatewhen favorable weather returns in the spring.

Dormancy imposed by restricting the uptake ofwater and exchange of oxygen can be removed bymechanically disrupting or removing the seed coat, aprocess called scarification. In the laboratory, scarifi-cation may be accomplished with files or sandpaper. Innature, abrasion by sand, microbial action, or passage ofthe seed through the gut of an animal will accomplishthe same end. Seed coats can be very tough. Uniformityand rate of germination of morning glory (Pharbitis nil ),cotton, and some tropical legume seeds, for example,can be improved by soaking the dry seed in concen-trated sulphuric acid for up to an hour. Scarification bypassage through animal gut no doubt occurs as a resultof the acidic conditions in the gut.

There is a considerable body of evidence to suggestthat seed coats also interfere with gas exchange, oxygenuptake in particular. Removal of the seed coat often leadsto a significant increase in respiratory consumption ofoxygen. Measurements of the oxygen permeability ofseed coats have been made and there is general agree-ment that permeability is very low in those seeds tested.However, it is not always clear that limited oxygenpermeability is the primary cause of dormancy. Thecomplexity of the situation and problems of interpreta-tion are well illustrated by studies of the genus Xanthium,or cocklebur. A cocklebur contains two seeds: an upper,dormant seed and a lower, nondormant seed. Dormancyof the upper seed can be overcome either by removingthe seed coat or by subjecting the intact seed to highoxygen tension. The inference is that seed coat perme-ability in the dormant seed limits the supply of oxygento the embryo and thus prevents germination. How-ever, several other observations have cast doubt on thisconclusion. There are, for example, no measurable dif-ferences between the dormant and nondormant seedwith respect to the permeability of the seed coat to oxy-gen. Moreover, the rate of oxygen diffusion through theseed coats is more than sufficient to support measuredrates of oxygen consumption by the embryos inside.Clearly, dormancy of the upper seed in Xanthium can-not be due to limited permeability of the seed coat tooxygen. Why then, do the upper, dormant seeds requirea higher oxygen level to elicit germination? It appearsthat the seed coat is a barrier, not to the uptake of oxy-gen but to the removal of an inhibitor from the embryo.Aqueous extracts of Xanthium seeds have revealed thepresence of two unidentified inhibitors, based on testsof the extracts in a wheat coleoptile elongation assay.The same two inhibitors are found in diffusate collectedfrom isolated embryos placed on a moist medium, butnot in diffusates from seeds surrounded by an intact seedcoat. Thus germination in the dormant seed appearsto be prevented by the presence of these inhibitorsand the seed coat serves as a barrier that preventsthose inhibitors from being leached out. The apparent

16.4 From Embryo to Adult 285

oxygen requirement can be explained by the observa-tion that high oxygen tension reduces the quantity ofan extractable inhibitor, presumably by some oxidativedegradation.

The seed coat may contain inhibitors that pre-vent growth of the embryo but the role of inhibitorsin seed dormancy is not clear. Along with hormonessuch as auxins and gibberellins, a large number ofpotential inhibitors have been identified in seeds, fruits,and other dispersal units. These include the hormoneabscisic acid, unsaturated lactones (e.g., coumarin), phe-nolic compounds (e.g., ferulic acid), various aminoacids, and cyanogenic compounds (i.e., compounds thatrelease cyanide) characteristic of apple and other seedsin the family Rosaceae. The simple presence of aninhibitor does not, however, prove a role in dor-mancy. The inhibitors could be localized in tissues notdirectly involved in growth of the embryo or otherwisesequestered so as to preclude any role in preventinggermination. Evidence in support of a role for inhibitorsis generally limited to leaching experiments such as thatdescribed above for Xanthium. In some cases, dormancycan then be restored by exposing the leached seed tothe inhibitor. In order to clearly establish whether aninhibitor has an active role in regulating germination,it is necessary to establish whether inhibitor levels inthe seed correlate with the onset and termination ofdormancy. In spite of the voluminous literature relatinginhibitors to dormancy, there is very little critical sup-port for a direct role. For the present, evidence for theimposition and maintenance of dormancy by inhibitorsremains largely circumstantial.

The dormancy and germination of many seeds arealso influenced by light and hormones. These factorswill be discussed in later chapters.

16.4 FROM EMBRYO TO ADULT

The first structure to emerge when a seed germinatesis the radicle. The radicle, which is the nascent primaryroot, anchors the seed in the soil and begins the processof mining the soil for water and nutrients. As the primaryroots elongates, it gives rise to branch, or lateral, roots.Unlike the situation in the shoot apical meristem, lateralroots do not originate in the root apical meristem.Lateral root primordia originate in the pericycle, a ringof meristematic cells that surround the central vascularcore, or stele, of the primary root. The growing lateralroot works its way through the surrounding cortex,either by mechanically forcing its way through or bysecreting enzymes that digest the cortical cell walls.Lateral root primordia arise in close proximity to thenewly differentiated xylem tissue, which allows vascularelements developing behind the growing tip of thelateral root to maintain connections with the xylem andphloem of the primary root.

Emergence of the radicle is followed by elonga-tion of the shoot axis. In some dicot seedlings, such asthe bean (Phaseous vulgaris) the hypocotyl (hypo, belowthe cotyledons) is the first part of the axis to elongate.The hypocotyl is hooked so that it pulls rather thanpushes the cotyledons and the enclosed first foliageleaves and shoot tip, called the plumule, up throughthe soil. This pattern is known as epigeal germina-tion (Figure 16.11A). In other dicots, such as gardenpea, the hypocotyl remains short and compact and thecotyledons remain underground. Instead, the epicotyl(epi, above the cotyledons) is the part of the shoot thatelongates. The epicotyl in this case forms a hook so

Epicotyl

Hypocotyl

C.B.A.

Cotyledons

Primaryroot Cotyledons

Primaryroot

Epicotyl

Primaryroot

Adventitiousroot

Coleoptile

First foliage leaf

FIGURE 16.11 Germination and seedling development. (A) Stages in the germina-tion of bean (Phaseolus vulgaris), an example of epigeal germination. (B) Hypogealgermination of pea (Pisum sativum). (C) Germination of corn (maize). See text fordetails.

286 Chapter 16 / Development: An Overview

that the clasping leaves of the plumule are pulled ratherthan pushed through the soil. This pattern of germina-tion, where the cotyledons remain in the soil is knownas hypogeal (Figure 16.11B). In either case, once theseedling axis breaks through the soil surface and emergesinto the light, the hook straightens and the primaryleaves unfold begin to expand. The principal objective ofthe hypocotyl or epicotyl hook is to bring the first leavesand meristem (the plumule) safely above the soil linein order to establish active photosynthesis. Without thehook, the plumule would no doubt be subject to consid-erable abrasion and damage as it moves through the soil.

A third example of seedling development isprovided by corn (maize), oats, and other cereal grains(Figure 16.11C). In the seed, both the plumule, includ-ing the first leaves and meristem, and the radicle aresurrounded by sheath-like structures—the coleoptileand the coleorhiza, respectively (see Figure 16.9B).The coleorhiza is the first structure to emerge from thepericarp. (In cereals, each grain is actually a fruit and thesurrounding tissues are the mature ovary walls whichfunction as a seed coat. In corn this is called the peri-carp.) Growing rapidly, the radicle quickly penetratesthe coleorhiza and assumes the function of the primaryroot. Meanwhile, the coleoptile is pushed upward byelongation of the mesocotyl or first internode. Thecoleoptile serves the same purpose as the plumular hookin dicots. It encases and protects the fragile first leavesuntil they emerge from the soil. Then the coleoptilesplits open, allowing the leaves to grow into the sunlight.

Elongation of the shoot axis proceeds througha combination of cell division and enlargement ofthe cells laid down by the meristem. The rate andextent of elongation is subject to a variety of controls,including nutrition, hormones, and environmental fac-tors such as light and temperature. The final heightof a shoot is determined by the rate and extent towhich internodes—the sections of stem between leafnodes—elongate. In some plants, such as pea (Pisumsativum), elongation occurs primarily near the apicalend of the youngest internode. The older internodeseffectively complete their elongation before the nextinternode begins. In other plants, elongation may bespread through several internodes, which elongate andmature more or less simultaneously (Figure 16.12). Stillothers exhibit changing rates of elongation with suc-cessive internodes, usually increasing. In some plants,internodes fail to elongate, thus giving rise to the rosettehabit in which all the leaves appear to originate frommore or less the same point on the stem. This rosettehabit is common in biennial plants (those that flower inthe second year) such as cabbage and root crops suchas carrot (Daucus carota) and radish (Raphanus sativus)before they reach the flowering stage. Failure of intern-ode elongation is commonly related to low levels of the

FIGURE 16.12 Internode elongation in broad bean (Viciafaba) over 48 hours. The initial spacing between markswas 2 mm. This internode has elongated more or lessuniformly over its entire length.

plant hormone, gibberellin, since application of the hor-mone usually stimulates internode elongation in rosetteplants.

16.5 SENESCENCE ANDPROGRAMMED CELL DEATHARE THE FINAL STAGESOF DEVELOPMENT

The final stage in the development of cells, tissues, andorgans is senescence, an aging process characterizedby increased respiration, declining photosynthesis, andan orderly disassembly of macromolecules. Senescingcells and tissues are metabolically very active—anumber of metabolic pathways are turned off andnew pathways, principally catabolic in character, areactivated. Catabolism of proteins, for example, releasesorganic nitrogen and sulfur in the form of solubleamines, while nucleic acids release inorganic phosphate.Chlorophyll is broken down and lipids are converted tosoluble sugars via gluconeogenesis. The products ofthese pathways are all small, soluble molecules that arereadily exported from the senescing tissue. Senescencethus enables the plant to recover nutrients from cells ortissue that have reached the end of their useful life andreallocate them to other parts of the plant that surviveor for storage in the roots.

Many of the new pathways are encoded by senes-cence associated genes (SAGs), which show increasedtranscription during senescence. The more than 50SAGs that have been identified to date fall generallyinto one of two broad groups. One group includes genesthat encode components of the system, such as theenzymes of the largely catabolic pathways that are newlyactivated during senescence. These include productssuch as ubiquitin, a range of proteolytic enzymes, and

Chapter Review 287

metallothioneins—proteins that chelate metal ions.The second group of genes regulates the initiation ofsenescence or its rate of progress. Among this groupare the so-called ‘‘stay-green’’ mutants characterized bydelayed leaf senescence. An interesting consequence ofthe ‘‘stay-green’’ phenotype is the potential for highercrop yields. Because senescence of the leaf is delayed,the period of active photosynthetic output and, conse-quently, yield are extended.

Senescence is a normal consequence of the agingprocess and will occur even when the supply of waterand minerals is maintained. For example, older leavesat the base of a plant commonly senesce in favor ofyounger leaves near the top. Flower petals senesce oncethe ovule has been fertilized and seed development isunderway. The leaves of perennial plants senesce at theend of the growing season and export the nutrients tothe roots for storage over the winter.

Senescence is one form of programmed cell death(PCD), broadly defined as a process in which the orga-nism exerts a measure of genetic control over the deathof cells. PCD requires energy and is normally regulatedby a distinct set of genes.

PCD is essential for normal vegetative and repro-ductive development. One example is the developmentof xylem tracheary elements. In order to function effi-ciently as a conduit for water transport, the protoplastof the developing tracheary element must die and beremoved at maturity. PCD also operates in the forma-tion of aerenchyma, a loose parenchymal tissue withlarge air spaces. Aerenchyma normally forms in thestems and roots of water lilies and other aquatic plants.These air spaces, created by a cell death program, pro-vide channels for oxygen transport to the submergedportions of the plant. Even corn (Zea mays) and otherterrestrial plants can be induced to form aerenchymawhen subject to flooding.

In the development of unisexual flowers, primordiafor both the male and female flowers are present in theearly stages. One or the other then aborts via a cell deathprogram, leaving only one type of organ to completedevelopment.

PCD is also an important factor in plant responsesto invading pathogens and abiotic stress. When a plantrecognizes a pathogen, for example, host cells in theimmediate area of the infection undergo PCD. Thisdeprives the invading pathogen of living tissue andeither slows or prevents it spread.

SUMMARY

Plants, like any other multicellular organism, buildcomplexity by combining growth with differentiation.Growth is simply an irreversible increase in size that

reflects an increase in cell number as well as the size ofindividual cells. Differentiation refers to the qualitativechanges, or specialization in cell structure and function,that normally a accompany growth. The sum of growthand differentiation is development.

Growth in plants is limited to discrete regionscalled meristems. The principal meristems that giverise to the primary plant body are located at the apicesof stems and roots. Secondary meristems in stems androots are responsible for increases in girth.

The life of an individual plant begins with fertiliza-tion or the union of a haploid sperm nucleus (suppliedby a pollen grain) with a haploid egg nucleus in theovary, located at the base of the pistil. The resultingzygote goes through a series of stages to produce amature embryo consisting of one or more cotyledons, aplumule consisting of primary leaves and a shoot apicalmeristem, and a radicle that is destined to become aprimary root. The embryo develops within a nutritivetissue called the endosperm.

Seed germination is initiated when the seedtakes up water to re-hydrate the dry tissues. Respi-ration is one of the first metabolic activities to bedetected as nutrient reserves are mobilized from theendosperm or cotyledons and the embryo renews itsgrowth.

Genetic control over the death of cells, calledprogrammed cell death, is a normal component ofdevelopment and gives rise to structures as diverseas mature xylem vessels and unisexual flowers.

CHAPTER REVIEW

1. Distinguish between growth, differentiation, anddevelopment. Can you give examples of each?

2. Describe the significance of meristems.3. Where are the principle meristems located in

plants? What is their contribution to developmentof the plant body?

4. If you had nailed an object to a young tree ata height of four feet off the ground and thenreturned after the tree had grown for severalyears, the object would still be four feet offthe ground. Why has the object not movedfurther from the ground as the tree grew?

5. Where is the egg cell in a plant and how isthe sperm delivered to effect fertilization?

6. Describe the process of seed formation from a fer-tilized egg cell?

7. Compare and contrast imbibition and osmosis.8. What is a plumular hook and what function

does it serve in young seedling development?

288 Chapter 16 / Development: An Overview

FURTHER READING

Bewley, J. D., M. Black. 1994. Seeds: Physiology of Developmentand Germination. New York: Plenum Press.

Bewley, J. D., M. Black, P. Halmer (eds.). 2006. The Encyclo-pedia of Seeds: Science, Technology, and Uses. CABI Publish-ing.

Buchanan B. B., W. Gruissem, R. L. Jones.2000. Biochemistry and Molecular Biology of

Plants. Rockville, MD: American Society of PlantPhysiologists.

Esau, K. 1977. Anatomy of Seed Plants. New York: Wiley.Raven, P. H., R. F. Evert, S. E. Eichhorn. 2005. Biology

of Plants. 7th ed. New York: Bedford, Freeman &Worth.

Steeves, T. A., I. M. Sussex. 1989. Patterns in Plant Develop-ment. 2nd ed. Cambridge: Cambridge University Press.

∗∗∗

Elongation

Wall stress

Turgor Pressure

17Growth and Development of Cells

Cells are the basic unit of life. The development ofa whole plant, as intricate as it may seem, ultimatelydepends on the growth and coordinated developmentof individual cells and groups of cells. The growth anddevelopment of cells, both individually and collectively,is in turn directed by a variety of internal and externalsignals such as hormones, light, temperature, gravity,insect predation, disease, and even the position of acell with respect to other cells. This chapter providesa general introduction to selected aspects of cellulardevelopment, including

• the structure of plant cell walls,• the dynamics of cell division and cell growth, and• an overview of signal perception and the signal

chains that enable cells to respond to those signal.

17.1 GROWTH OF PLANT CELLSIS COMPLICATED BY THEPRESENCE OF A CELL WALL

Plant cells are characterized by a complex mixture ofmaterials, called the extracellular matrix (ECM), thatlie outside the plasma membrane. The ECM is dom-inated by the cell wall, which provides protection forthe underlying protoplast and is ultimately responsible

for maintaining cell shape and the structural integrityof the plant. Two types of cell walls are recognized:primary walls, which surround young, actively growingcells, and secondary walls that are laid down as the cellsmature and are no longer growing.

17.1.1 THE PRIMARY CELL WALL IS ANETWORK OF CELLULOSEMICROFIBRILS ANDCROSS-LINKING GLYCANS

The primary wall is very thin, measuring only a fewmicrometers in thickness. Its principal constituent islong, threadlike chains of α−1 → 4−linked glucoseunits, called cellulose. The individual cellulosemolecules are bundled in long, parallel arrays calledmicrofibrils (Figure 17.1). Each microfibril is approxi-mately 5 to 12 nm in diameter and contains, in higherplants, approximately 36 individual cellulose chainsin cross-section. A single cellulose chain may containas many as 3,000 or more glucose units but, becausethe chains begin and end at different places within themicrofibril, an individual microfibril may contain severalthousand cellulose chains and reach lengths of severalhundred micrometers. Adjacent cellulose chains withina microfibril are held together by hydrogen bondsbetween hydroxyl (—OH) groups on adjacent glucoseunits. This bonding arrangement makes a microfibril

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290 Chapter 17 / Growth and Development of Cells

OH

OH

HOHO

HO

HO

O

OO

OO

OH

OH

HOHO

HO

HO

O

OO

OO

OH

OH

HOHO

HO

HO

O

OO

O

OH

OH

HOHO

OH

OH

HO

O

O

OH

OH

HO

O

OHO

HO

O

OO

O Hydrogenbonds

Cell

1 4-�-Glucose chains

Primary cell wall

Cellulose microfibril

FIGURE 17.1 The principal structural components of cell walls are cellulose microfib-rils constructed of (1→4)-linked β-glucose chains. Adjacent cellulose chains withinthe microfibril are joined by intermolecular hydrogen bonds.

very strong. In fact, the tensile strength of a microfibril,or its ability to withstand tension without breaking, issimilar to that of a steel wire of the same size.

The orientation of microfibrils in the primary wallis more-or-less random (Figure 17.2), although in elon-gating cells they tend to orient parallel to the directionof growth. The cellulose microfibrils within the pri-mary wall are held in position by cross-linking glycans.1Cross-linking glycans are noncellulosic polysaccharidesthat bind to the cellulose microfibrils, but are also longenough to bridge the distance between neighboringmicrofibrils and link them into a semi-rigid network(Figure 17.3). In the primary cell wall of all dicotyledo-nous (dicot) species and about one-half of the monoco-tyledons (monocot) species, the principal cross-linkingglycans are xyloglycans (XyGs). Xyloglycans are linear(1→4)β-glucose chains (like cellulose), but with a 5-car-bon sugar, xylose, linked to the oxygen at the carbon-6position on many of the glucose units (Figure 17.4). Insome species, some of the xylose units may be replaced

1Cross-linking glycans are the principal constituent of whatwas formerly referred to as ‘‘hemicellulose,’’ a heterogeneousbut generally uncharacterized mixture of cell wallpolysaccharides characterized solely by their extractabilityinto strong alkali solution.

by other sugars, such as arabinose, fructose, galactose,or glucuronic acid. Some side chains may be composedof two or three of these sugars instead of one.

17.1.2 THE CELLULOSE–GLYCANLATTICE IS EMBEDDED IN AMATRIX OF PECTINAND PROTEIN

As much as 35 percent of the primary wall consistsof pectins, or pectic substances. Pectin is a complex,heterogeneous mixture of noncellulosic polysaccharidesespecially rich in galacturonic acid—the acid form ofthe sugar galactose. When the pectin chain is secretedinto the wall space, most of the free carboxylic acidgroups are esterified with methyl groups. Once in thewall, however, the enzyme pectin methylesterase cleavessome of the methyl groups, leaving the acid groups freeto bind with calcium. The divalent calcium ions formcross-links between pectin chains and contribute to thestability of the cell wall. It is significant that calciumconcentrations (and, consequently, calcium bridging)are kept low in the walls of actively growing cells.

Pectins are also the principal constituent of themiddle lamella. The middle lamella lies between theprimary walls of adjacent cells. It is the ‘‘cement’’ that

17.1 Growth of Plant Cells is Complicated by The Presence of a Cell Wall 291

FIGURE 17.2 The primary cell wall. An electron micrograph of the primary wall ofa parenchyma cell from the coleoptile of an oat (Avena) seedling. Note the poresthrough which plasmodesmata pass. (From Bohmer, H. 1958. Untersuchungen uberdas Wachstum und den Feinbau der Zellwande in der Avena-Koleoptile. Planta50:461–497, Figure 20. Copyright Springer-Verlag, Heidelberg.)

holds the cells together. The softening of fruit as itripens, for example, is due in part to the action of theenzyme polygalacturonase, which degrades the pecticsubstances in the middle lamella and loosens of thebonds between the cells.

Primary cell walls also contain approximately 10percent glycoprotein which have an unusually highcontent of the amino acid hydroxyproline. Thesehydroxyproline-rich glycoproteins are called extensin.Although the precise function of extensin is unknown,

Middle lamella pectin

Middle lamella

Primary wall

Plasma membrane

50 nm

Pectin

Cellulose

Xyloglucan

FIGURE 17.3 A simplified model to illustrate how cellulose microfibrils, cross-linkingxyloglucans, and pectic substances might be arranged in the cell wall. Not shownare extensin and other cell wall proteins. (From McCann, M. C., K. Roberts. 1991.Architecture of the primary cell wall. In: C. W. Lloyd (ed.), The Cytoskeletal Basisof Plant Growth and Form. Academic Press. Original figure courtesy of Dr. M. C.McCann.)

292 Chapter 17 / Growth and Development of Cells

Xylose

Glucose Glucose Glucose Glucose

Xylose

Xylose

FIGURE 17.4 A schematic diagram illustrating the struc-ture of a simple cross-linking xyloglycan.

it is thought to be one component of a structuralnetwork that adds strength to the wall and locks the wallinto shape once the cell stops growing. Several otherfamilies of cell wall proteins rich in proline, glycine, orthreonine have been described more recently, addingto the apparent complexity of cell wall structure.

17.1.3 CELLULOSE MICROFIBRILS AREASSEMBLED AT THE PLASMAMEMBRANE AS THEY AREEXTRUDED INTO THECELL WALL

Cellulose synthesis is catalyzed by cellulose synthase, amultimeric enzyme that is localized in the plasma mem-brane (Figure 17.5). Although active cellulose synthasehas proven difficult to isolate, the enzyme complex canbe visualized in electron micrographs of membranesprepared by a technique known as freeze-fracturing. Atleast in angiosperms, the enzyme appears to be orga-nized in the form of a rosette composed of six subunits.The rosette not only synthesizes the cellulose chain, butassembles the chains into microfibrils and extrudes themicrofibrils into the cell wall.

The immediate precursor for cellulose synthesis isuridine diphosphoglucose (UDP-Glu). UDP-Glu canbe formed directly from sucrose by certain forms ofthe enzyme sucrose synthase, and also appears to beassociated with the plasma membrane. It thus appearsthat sucrose synthase may deliver UDP-Glu directly tothe catalytic site of the cellulose synthase rosette, wherethe glucose is added to the terminus of the growingmicrofibril.

The details of cellulose biosynthesis are not wellunderstood, in part because of the difficulty in isolatingactive enzyme. Even the rosettes are absent from isolatedmembranes, leading to the suggestion that some form ofinteraction with the cytoskeleton is required to stabilizethe enzyme in situ. When this interaction is disrupted,the complex dissociates and its activity is lost.

17.2 CELL DIVISION

The division of plant cells, like any eukaryotic cell,occurs in two steps: mitosis or karyokinesis, which

Cytosol

Precursorglucose-UDPmolecules

Cell wallCellulosemicrofibril

Cellulosesynthase

Plasmamembrane

UDP

UDP

UDP

UDP

FIGURE 17.5 A model for cellulose synthesis fromprecursor molecules, uridine diphosphoglucose(UDP-glucose). The enzyme, formally known asUDP-glucose:(1→4)-β-glucan glucosyltransferase, isbelieved to be located in the plasma membrane where itsimultaneously synthesizes the cellulose chains, assem-bles them into microfilaments, and extrudes the micro-filaments into the cell wall.

results in the creation of two new nuclei, and cytokine-sis, the subsequent division of the cytoplasm to createtwo separate daughter cells. Between divisions, however,each of the daughter cells must increase the amount ofcytoplasm as it enlarges, replicate its DNA, and preparefor the next division. This sequence of events is knownas the cell cycle.

17.2.1 THE CELL CYCLE

The cell cycle can be described as four distinct phases:the mitotic phase (M), the DNA synthesis phase (S),and the two intervening phases or ‘‘gaps’’ (G1 andG2) (Figure 17.6). The M phase is the division phase inwhich chromosomes are condensed and their distinctivemorphology becomes visible with a light microscope.Throughout the other three phases, collectively referredto as interphase, the chromosomes are fully extended anduncoiled and are not visible with a light microscope.During S phase, DNA replication leads to the formationof two identical copies of the chromosomes. In the G2phase, the chromosomes begin to condense and the cellassembles the rest of the machinery necessary to movethe chromosomes apart during mitosis.

17.2 Cell Division 293

INTERPHASE

DIVISION

M

G2

G1

S

FIGURE 17.6 Phases of the cell cycle. M is the nucleardivision phase, or mitosis. During mitosis the chromo-somes condense and the sister chromatids are separated.S is the phase during which the chromosomes are diffuse,but DNA synthesis occurs. Mitosis and the synthesisphases are separated by two gaps in time. The progressof the cell cycle is normally controlled at either the G1 toS transition, or the G2 to M transition.

The time required to complete a cell cycle is highlyvariable, depending on the cell type and various devel-opmental cues, but the length of the S phase and theM phase remain roughly constant for a given cell type.G1 is the most variable phase and may account for themajor portion of a cell’s life span.

In order for development to proceed in an orderlymanner, the timing and rate of cell division and con-sequently entry into the cell cycle must be preciselycontrolled. The required control is achieved through acomplex interplay of various kinases, phosphatases, andproteases that respond to both intrinsic and externalsignals. Central to cell cycle control are the activi-ties of cyclin-dependent kinases (CDKs). CDKs arecomposed of two subunits—one subunit functions as acatalyst and the other serves to activate the complex.The activating subunit is a small, unstable protein calledcyclin, so named because when they were first discov-ered in Xenopus eggs, it was observed that during eachcell cycle the concentration rose steadily from zero andthen suddenly collapsed.

In plants, there are at least four classes of cyclins (A,B, D, and H) and the association of CDKs with specificcyclins is a key regulatory factor. Equally important isthe availability of cyclins as the levels of the activatingsubunit oscillate during the cell cycle due to regulatedsynthesis and degradation. Activation of CDKs also

requires phosphorylation of a specific threonine residue(Thr160) on the catalytic subunit by a CDK-activatingkinase (CAK), while phosphorylation of other threonineand tyrosine residues (Thr14, Tyr15) inhibits CDK acti-vity. Phosphatase enzymes are available to remove theinhibitory phosphate groups at the appropriate time.The two principal sites of CDK control appear to be atthe G1 to S or G2 to M transitions. It is assumed that thefunction of an active CDK complex is to phosphorylateand thus activate other proteins involved in the synthesisof DNA (S phase) or initiation of mitosis (M phase).

CDK inhibitors (CKI) also play a pivotal role byarresting cell division when necessary, such as at the endof a developmental program or in response to hormonalor environmental cues. Operating principally at the G1to S transition, a CKI will bind with an active CDKcomplex in order to prevent it from phosphorylatingsubstrate. When conditions are appropriate for the cellcycle to continue, the CKI can be removed by proteoly-sis. First, CKI is marked for degradation by attachmentof ubiquitin (See Box 17.2). The ubiquitinated CKI isthen degraded, the CDK complex reactivated, and theG1 to S transition is allowed to proceed.

17.2.2 CYTOKINESIS

Cytokinesis in animal cells is relatively simple, involvinga constriction of the cell membrane which advancestoward the center until the one cell becomes two cells.The existence of the cell wall prevents a similar patternof cell division in plant cells. Plant cells must insteadconstruct new membranes and a new extracellular matrix,including a middle lamella and two new primary walls,inside a living cell.

The formation of the new walls begins in the latemitotic or M phase of the cell cycle, after the two sets ofchromosomes have separated and moved toward oppo-site poles in the cell and the new nuclear membraneshave begun to form. At this point, the mitotic spindle dis-appears and the microtubules that had made up the spin-dle disassemble and then reassemble to form a cluster ofinterdigitating microtubules oriented perpendicular tothe plane of the new crosswall (Figure 17.7) (Box 17.1).Called the phragmoplast, these microtubules serve todirect the movement of small secretory vesicles, derivedfrom nearby Golgi complexes, into the equatorialregion of the cell. The vesicles begin collecting in thecenter of the cell where they align along the equatorialplane and begin to fuse with one another. The resultingaggregate of vesicles is called the cell plate.

As newly arrived vesicles continue to add to thecell plate, the plate grows outward in all directionsuntil its leading edge makes contact with the existingplasma membrane surrounding the parent cell. Fusionof the plate membranes with the parent cell membrane

294 Chapter 17 / Growth and Development of Cells

FIGURE 17.7 The phragmoplast and cell platein a dividing plant cell. Following nucleardivision, tubulin subunits originating fromremnants of the mitotic spindle reorganize asmicrotubules oriented perpendicular to theplane of cell division. These microtubules,collectively called the phragmoplast, serveto track small, Golgi-derived secretory vesi-cles into the equatorial plane of the cell. Thevesicles fuse to form the cell plate, whichgradually grows outward toward the lateralmembranes of the dividing cell. The grow-ing cell plate eventually fuses with the lat-eral membranes. The membranes of the cellplate form the new plasma membranes of thedaughter cells and the contents of the orig-inal Golgi vesicles form the middle lamellabetween them. Once membrane fusion iscomplete, each daughter cell completes a newcrosswall by depositing a new primary wall.

Golgi body

Nucleus

Nucleus

Plasma membrane

Cell plate

Primary cell wall

Middle lamella

Phragmoplastmicrotubules

effectively completes the separation of the two daugh-ter cells. The interior of the plate—between the twomembranes—is filled with the pectic substances deliv-ered by the Golgi vesicles and that now make up themiddle lamella between the daughter cells. Cytokinesis iscomplete when cellulose synthase complexes are insertedinto the new membranes and each daughter cell depositsa new primary wall adjacent to the middle lamella.

17.2.3 PLASMODESMATA ARECYTOPLASMIC CHANNELS THATEXTEND THROUGH THE WALLTO CONNECT THEPROTOPLASTS OFADJACENT CELLS

When the cell plate forms, membrane fusion is notcomplete. This leaves some locations where cytoplasmcontinuity is maintained between daughter cells. As thecellulose is laid down and the new wall increases inthickness, these connections form membrane-encasedchannels called plasmodesmata (sing. plasmodesma)(Figure 17.9). The membrane that encases the channel isa continuation of the plasma membranes from adjacentcells. Running through the center of the plasmodesmais a second membranous tube—a tube within a tube—called the desmotubule. The desmotubule is formedas an extension of the endoplasmic reticulum that wasentrapped during the formation of the cell plate. Asleeve of cytosol fills the space between the desmotubuleand plasmodesmata itself.

Plasmodesmata are not large—approximately60 nm in diameter—but there are often large numbersof them. Estimated frequencies are in the range of 0.1to 10.0 μm−2 of cell wall, although there is a tendencyfor plasmodesmata to be grouped in roughly oval areascalled primary pit fields. Plasmodesmata are smallenough to preclude the exchange of organelles betweencells but large enough to permit the diffusion of smallsolute molecules, including infectious viral RNA andplant transcription factors, through the cytosolic sleeve.Plasmodesmata thus provide a measure of membraneand cytosolic continuity between cells and allow forsupracellular control over developmental programs.

The connection of neighboring protoplasts throughplasmodesmata creates a continuous cytoplasmic net-work, referred to as the symplast, throughout the plant.In a similar manner, the apoplast consists of continu-ous extracellular or noncytoplasmic space. The apoplastconsists of interconnected cell walls, intercellular spaces,and mature, nonliving vascular tissue. The concept of asymplast and an apoplast is especially useful when con-sidering the movement of water and dissolved solutesthroughout the plant.

17.3 CELL WALLSAND CELL GROWTH

Although the primary wall is not very thick, its inter-locking network of cellulose microfibrils, cross-linkingxyloglycans, and structural proteins confers remarkable

17.3 Cell Walls and Cell Growth 295

BOX 17.1CYTOSKELETON

Virtually all eukaryotic cells, both animal and plant,contain a three-dimensional, interconnected network offibrous protein called the cytoskeleton. The cytoskele-ton plays vital roles in determining the organization ofcytoplasm and cell shape, and in cell division, growth,and differentiation.

The cytoskeleton of plant cells is composed of twodifferent elements: microtubules and microfilaments.Microtubules are long rods approximately 24 nm indiameter and a hollow core about 12 nm in diameter.They are assembled from subunits of a globular proteincalled tubulin, which has a molecular mass of approx-imately 100,000 daltons (100 kD) and is made up oftwo globular polypeptides (α-tubulin and β-tubulin).1Microtubules are formed when tubulin subunits sponta-neously self-assemble into long chains called protofila-ments. Protofilaments then line up laterally to form themicrotubule wall (see Figure 17.8). Microfilaments aresolid threads composed of actin, also a globular protein.Two chains of actin subunits self-assemble in a helicalfashion to form a microfilament approximately 6 nm indiameter.

The term cytoskeleton is an unfortunate choicebecause it implies a rigid structure with a staticfunction. Instead, the cytoskeleton is very dynamic.Microtubules in particular are constantly beingassembled, disassembled, and rearranged as the celldivides, enlarges, and differentiates. Microtubulesform the mitotic spindle, which plays a significantrole in the movement of chromosomes during celldivision. Microtubules also determine the orientationand location of the new cell wall between daughtercells, and the deposition of cellulose in growing cellwalls.

Microfilaments appear to control the directionof cytoplasmic streaming, the continuous flowof cytoplasmic particles and organelles around theperiphery of the cell. The microfilaments formaggregates or bundles oriented parallel to the directionof cytoplasmic flow. Microfilaments are also involved in

1The molecular mass of a molecule or particle is expressedin units of Daltons, defined as 1/12 the mass of a carbon atom.Molecular mass should not be confused with molecularweight, which is a dimensionless quantity expressing the ratioof particle mass to 1/12 the mass of a carbon atom. Molecularweight is symbolized by M, for relative molecular mass.

the growth of pollen tubes. When a pollen grain germi-nates, it develops a tubular extension that grows downthe stigma of the flower and serves to deliver the malenucleus to the egg. Growth of the tube is only at the tip,and vesicles that contain cell wall precursors are guidedthrough the cytoplasm to the growing tip by a networkof microtubules.

The importance of the cytoskeleton in organizingand coordinating the dynamic properties of growingcells is only beginning to be appreciated. Techniquesfor the study of plant cell cytoskeleton are rapidlyimproving, and we can expect exciting advances in thefuture.

45

6

7

8

91011

12

13

1

2

3

12

34

56

78

910

1112

13 Protofilament

24 nm 6 nm

α

β

α

β

α

β

FIGURE 17.8 Microtubules and microfilaments. Left: Dia-gram of a microtubule in cross-section and longitudinalview. The number of vertical protofilaments varies from11 to 15 but is usually 13, as shown here. Protofilamentsare offset to form a helix. Right: A microfilament is com-posed of two parallel strands of globular subunits twistedto form a helix.

296 Chapter 17 / Growth and Development of Cells

A.

B.

Desmotubule

ER

CELL B

Wall(Cell B)

Wall(Cell A)

CELL A

ER

Middle lamella

FIGURE 17.9 Plasmodesmata. (A) Electron micrographshowing plasmodesmata connecting adjacent cells. (B)Diagram of a plasmodesmata showing the relationshipbetween plasma membranes, ER, and desmotubules.

strength and rigidity to the wall. This strength andrigidity maintains the shape of the cell and its structuralrelationship with neighboring cells and, ultimately, thesupport of the entire plant. Thus a growing cell faces adelicate balancing act. It must, at the same time, main-tain the strength and structural integrity of the wallwhile remaining sufficiently pliant to provide space forthe expanding protoplast.

17.3.1 CELL GROWTH IS DRIVEN BYWATER UPTAKE AND LIMITEDBY THE STRENGTH ANDRIGIDITY OF THE CELL WALL

Since most of the volume of any cell is water, it followsthat for a cell to increase its volume it must take up water.If, for example, a cell is bathed in an isotonic solution of

mannitol or similar solute that cannot enter the cell, thecell will also not take up water and it will not grow. Thedriving force for cell enlargement is the uptake of water.

Recall that cells take up water by the process ofosmosis. The high solute concentration of the proto-plasm and vacuolar sap decreases the water potential ofthe cell to the point where water diffuses into the cell.With no means to compensate, a cell surrounded by purewater might continue to swell indefinitely or at least untilinternal pressures exceeded the tensile strength of themembrane. The consequences of such a situation arevividly demonstrated when mammalian red blood cellsare placed in water. The cells quickly swell until theplasma membrane bursts, releasing their contents intothe medium. Most animal cells avoid such osmotic dis-aster by using metabolic energy to excrete either soluteor water and thus maintain a favorable pressure bal-ance. Plant cells have found a different solution—theysurround the plasma membrane with a strong, more orless rigid cell wall. Turgor pressure (a positive pressure)developed as the expanding protoplast pushes againstthe wall rises until it balances the negative osmotic pres-sure of the protoplast. At that point the water potentialof the cell approaches zero and no further net wateruptake—or cell growth—will occur. It is the strengthand rigidity of the cell wall that imposes critical restric-tions on the capacity of plant cells to grow. Thus, fora cell to increase in size, the strength and rigidity ofthe cell wall must be modified in order to reduce thewater potential of the cell, permit water uptake, and,consequently, allow the cell to enlarge.

17.3.2 EXTENSION OF THE CELL WALLREQUIRES WALL-LOOSENINGEVENTS THAT ENABLELOAD-BEARING ELEMENTS INTHE WALL TO YIELD TOTURGOR PRESSURE

Any increase in volume of a cell requires a correspondingincrease in the surface area of the surrounding wall, orwall extension. Investigators know that wall extensionis related to turgor pressure—this has been demon-strated empirically. For example, when turgor pressureis experimentally reduced, the rate of cell expansion alsodeclines. Furthermore, wall extension and growth donot occur in cells at very low or zero turgor pressure,even though the cells remain metabolically active andappropriate growth stimuli are present. This interde-pendence of wall extension and turgor pressure can besummarized by the following simple relationship:

dV/dt = m(P − Y) (17.1)

The term dV/dt is the change (d) in volume (V) of acell over time (dt). dV/dt is thus a simple way to expressthe growth rate of a cell. Y is the yield threshold, or the

17.3 Cell Walls and Cell Growth 297

∗∗∗

2b

2a

1

Elongation

Wall stress

Turgor Pressure

PL.

Yielding

Relaxation

Creep

EL.EL.

FIGURE 17.10 A model for stress relaxation in the wall ofa growing cell. Both elastic (EL) and plastic (PL) compo-nents of the wall bear the stress of an expanding proto-plast. In 1, turgor causes stress in the wall and extensionof the elastic component, represented as springs. In 2a,yielding of the plastic components allows relaxation inthe elastic component, illustrated by contraction of thesprings. In 2b, turgor reestablishes wall stress and thewall expands to the same extent that the plastic compo-nent has lengthened. (From Cosgrove, D. I. 1987. PlantPhysiology 84:561–564. Copyright American Society ofPlant Physiologists.)

minimum turgor pressure necessary for cell expansion tooccur. The term m is wall extensibility, a proportionalityconstant between growth rate and turgor pressure (P)in excess of the yield threshold. Wall extensibility isa quantitative measure of the capacity of the wall toirreversibly increase its surface area. According to thisrelationship, the growth of a cell depends principally onthe amount by which turgor pressure exceeds the yieldthreshold.

It is apparent that a growing cell is faced with con-flicting roles of turgor pressure. On the one hand, turgorpressure opposes the continued uptake of water, whichis the driving force for cell expansion. At the same time,turgor pressure promotes irreversible wall extension andcell enlargement. How does the cell resolve this con-flict? The answer to this question is provided by theconcept of stress relaxation. Stress relaxation in the wallis central to the process of cell growth.

Turgor pressure develops because the cell wall—cellulose microfibrils cross-linked with xyloglucans—resists deformation as the protoplast attempts to expand.The force of the expanding protoplast pushing againstthe wall thus generates stress (defined as force/unitarea) within the wall. Cell growth appears to be initi-ated when these stresses are relaxed by wall-looseningevents that cause load-bearing elements in the wall to

yield (Figure 17.10). The most likely candidate for theload-bearing element is the xyloglucan linking two adja-cent cellulose microfibrils. If the xyloglucan preventsdisplacement of the microfibrils, then a loosening of thexyloglucan cross link would constitute a stress relaxationand allow the cellulose microfibrils to move apart. Theresult would be a simultaneous and proportionate reduc-tion in turgor pressure. A reduction in turgor pressureleads to a decrease in the water potential of the cell(�� becomes more negative), followed by the passiveuptake of water. The influx of water in turn increasescell volume, extends the cell wall, and tends to restoreboth wall stress and turgor pressure. The process of cellgrowth is thus seen as a continuous adjustment of turgorpressure through stress relaxation in order to balance itsconflicting roles in water uptake and cell wall extension.

17.3.3 WALL LOOSENING AND CELLEXPANSION IS STIMULATED BYLOW PH AND EXPANSINS

It has been known since the 1930s that plant tissueselongate faster when bathed in a medium with a low pH(Figure 17.11). In the 1970s, a large body of work estab-lished that this ‘‘acid-growth’’ phenomenon applied notonly to living tissues, but heat-killed tissues and iso-lated cell walls as well. These experiments gave rise tothe concept of a wall-loosening enzyme with a low pHoptimum.

In the early 1990s, two cell wall proteins wereisolated and found to stimulate the expansion of cucum-ber hypocotyl sections that had been heat-treated in

1.0

2.0

3.0

4.5 5.0 5.5 6.0 6.5 7.0

Rel

ativ

e el

onga

tion

pH

FIGURE 17.11 An experiment demonstrating theacid-growth response. Apical segments (5 mm) fromdark-grown oat coleoptiles (Avena sativa) were floatedon buffers at the indicated pH. The increase in length ofthe segments was measured after 18 hours. (From data ofHopkins & Hillman. 1965. Planta 65:157–166.)

298 Chapter 17 / Growth and Development of Cells

order to inactivate endogenous wall-loosening activ-ity. These proteins, called expansins, characteristi-cally induce stress relaxation and extension of isolatedcell walls at low pH. Expansins are small proteins,with a relative molecular mass of 26,000. In termsof dry mass—one part protein per 5,000 parts cellwall—they might be considered minor proteins. How-ever, expansins are very active proteins and will inducecell wall extension when added in amounts as low as 1part expansin per 10,000 parts of cell wall. The responseis also very rapid—extension can be detected withinseconds of adding expansins to the cell wall test sys-tem. In addition to stimulating extension of isolatedcell walls, expansins enhance the rate of growth whenapplied to living cells. Moreover, expression of expansingenes is highest in growing and differentiating tissues,adding further weight to the hypothesis that expansinsare significant wall-loosening agents.

Just how expansins achieve their unique effectson the physical properties of cell walls is not yetknown, although two possibilities present themselves.The first possibility is that expansin hydrolyzes thecross-linking glycans or other elements of the wallmatrix that hold the cellulose microfibrils in place. How-ever, no hydrolytic activity on the part of expansins hasbeen demonstrated to date. There are known enzymeswhose activity is to hydrolyze components of the wallmatrix, but these enzymes do not induce stress relaxationand extension. The second possibility is that expansinsattack and weaken the noncovalent bonds by which thecross-linking glycans attach to the cellulose microfib-rils. According to this proposal, expansin would migratealong the surface of the cellulose microfibril and weakenthe attachment of the xyloglucan. The partial detach-ment of a xyloglucan that was under tension would allowit to relax. The cellulose microfibrils would then be dis-placed in response to turgor until tension is reestablishedin the cross-link and new cross-links are formed. In thisway, expansin would not progressively weaken the cellwall, but would catalyze an inchworm-like movement ofthe wall polymers.

17.3.4 IN MATURING CELLS, ASECONDARY CELL WALL ISDEPOSITED ON THE INSIDE OFTHE PRIMARY WALL

When a cell stops enlarging and begins to mature, asecondary cellulose wall is laid down on the inside ofthe primary wall. Secondary walls are much thickerand more rigid than primary walls. They contain upto 45 percent cellulose, correspondingly lower amountsof cross-linking glycans, and relatively little pectic sub-stance. In thick-walled woody cells, the secondary wallfrequently consists of two distinct zones, characterizedby differing orientation of the microfibrils. In both zones

the microfibrils are oriented helically around the cell.In the outermost layer, adjacent to the primary wall, themicrofibrils are oriented at a large angle to the long axisof the cell. In the inner layer, the microfibrils are almostparallel to the long axis. In some cells, such as fibers, thesecondary wall may be so thick that only a very smalllumen, devoid of protoplasm, remains in the center atmaturity.

Most secondary walls also contain lignin, whichmay account for as much as 35 percent of the dry weightof woody tissues. Next to cellulose, lignin is probablyone of the most important biological substances in termsof structural importance. Lignin is a plastic-like poly-mer that has a high degree of strength, stronger eventhan cellulose microfibrils. It is extremely resistant toextraction without chemical degradation, which makesits chemistry difficult to study. It is known to consist ofa complex system of interlocking bonds between severalrelatively simple phenolic alcohols. The combination ofcross-linked cellulose microfibrils embedded in a matrixof pectic substances and lignin is responsible for theexceptional strength of wood. In engineering terms,secondary cell walls are a composite material, similar tomodern fiber-reinforced plastics and with many of thesame properties. The composite structure of cell walls iswhat enables tall trees to withstand the stresses of highwinds and that makes wood such a useful and importantbuilding material.

17.4 A CONTINUOUS STREAMOF SIGNALS PROVIDESINFORMATION THATPLANT CELLS USE TOMODIFY DEVELOPMENT

Plant cell development is highly regulated by a varietyof signals that operate at several levels. Hormones arechemical messengers that enable cells to communicatewith one another. Several classes of plant hormones,including auxin, gibberellins, cytokinins, abscisic acid,ethylene, and brassinosteroids, are known to promoteor inhibit various developmental responses, either singlyor in combination. Other intracellular factors such asturgor, mineral status, and internal clocks may also causea cell to modify its metabolism and development.

Plant cells are also bombarded with external sig-nals that provide information about their environmentand that are used to modulate development accordingly.Light, temperature, gravity, and insect predation havethe most obvious and dramatic impact. Factors such asmagnetic field, sound, and wind (a mechanical stimu-lus) may have more subtle effects, but these have beendifficult to establish experimentally. Other environmen-tal factors such as soil moisture, humidity, nutrition,

17.4 A Continuous Stream of Signals Provides Information That Plant Cells Use to Modify Development 299

and pathogens may also influence development in somecases. More recently it has become evident that airand water pollutants represent not only an importantenvironmental challenge to plants but may modify devel-opmental patterns as well.

17.4.1 SIGNAL PERCEPTIONAND TRANSDUCTION

A signal, regardless of whether it originates inside oroutside the plant, can have no effect unless there is areceptor that allows it to be detected or perceived bythe cell. The presence or absence of the appropriatereceptor determines which cells are able to respond to aparticular signal. There must then be a mechanism thatconverts, or transduces, the signal to some change in thechemistry or metabolism of the cell, or gene expression,that will ultimately give rise to the intended response.All developmental signals, regardless of their nature,share this sequence of signal perception, transduction, andresponse.

There are two principal mechanisms for signal per-ception and transduction. The first involves a receptorprotein associated with the plasma membrane. Thereceptor is both specific to the signal (e.g., hormonemolecule) and characteristic of the target cell. Theformation of a signal-receptor complex then sets intomotion a cascade of biochemical events that alters someaspect of cellular metabolism. In the second mechanism,the signal (e.g., a hormone molecule) is taken up by thecell and migrates into the nucleus where it reacts with anuclear-based receptor to either activate or repress geneexpression.

The two mechanisms are not mutually exclusive.The hormone auxin, for example, controls cell enlarge-ment through a plasma membrane-based receptor while

it controls more complex developmental responsesthrough a nuclear-based receptor.

17.4.2 THE G-PROTEIN SYSTEMIS A UBIQUITOUSRECEPTOR SYSTEM

One receptor system that is proving to be ubiquitous inplants and animals is the G protein-coupled receptor.G-proteins are a large family of guanosine triphos-phate (GTP) binding proteins that have long beenknown for their role in the response of animal cellsto hormones, neurotransmitters, and a variety of othersignals that operate across the cell membrane. However,recent genetic evidence from Arabidopsis and rice (Oryzasativa) has linked the presence of G proteins to signalperception in plants as well (Figure 17.12).

The two major players in the G protein system arethe G protein itself, which consists of three distinct sub-units (designated Gα, Gβ, and Gγ) and is located on thecytoplasmic side of the plasma membrane, and a trans-membrane protein known as the G-protein-coupledreceptor, or GPCR. GPCR is commonly referred toeither as a 7-trans-membrane (7TM) receptor or hep-tahelical protein because the protein consists of sevenα-helix domains that span the membrane. The sevenhelices are in turn connected by three hydrophilic loopsthat extend into the aqueous environment on each ofthe outer and inner surfaces of the membrane.

In effect, the G protein system functions as a bio-chemical on/off switch. In the absence of a hormoneor other regulatory signal, the three subunits of theG protein combine to form an inactive complex that islocated on the cytoplasmic surface of the membrane. Inthe inactive form (i.e., when the switch is ‘‘off ’’), theGα subunit carries a molecule of guanosine diphosphate

Ligand

Cytoplasm

GTP

Pi

GTPGDP Effector

Effector

GDP

GPCR

2

1

3 4 5

5

6

α

α

αβ

β

β

δ

δ

δ

FIGURE 17.12 A model for signal transduction by the Gprotein receptor system. In step 1, a signal molecule (theligand) binds to the transmembrane G protein coupledreceptor (GPCR), increasing its affinity for the G pro-tein (step 2). In step 3, the G protein exchanges its GDPfor GTP, which causes the dissociation of the Gα subunitfrom the Gβγ dimmer and the release of both subunitsfrom the receptor (step 4). Both the Gα subunit and theGβγ dimer may bind to an effector, thus activating ordeactivating the effector (step 5). In step 6, the inher-ent GTPase activity of Gα hydrolyses the GTP, whichdeactivates Gα and allows the two subunits to recombine.

300 Chapter 17 / Growth and Development of Cells

(GDP). Signal transduction begins when a ligand (ahormone or other small regulatory molecule) binds toGPCR on the outer cell surface. Ligand binding causesthe GPCR molecule to alter its conformation, or shape,such that it now has a higher affinity for the G pro-tein. The G protein binds to the cytoplasmic surfaceof the receptor and the Gα subunit exchanges its GDPfor a molecule of guanosine triphosphate (GTP). TheG protein is now active, or ‘‘on.’’

In the next step, the activated G protein dissociatesfrom the receptor and the Gα subunit dissociates fromthe combined Gβγ subunit. Each Gα and Gβγ subunitis now free to activate an effector protein. Effectorsare enzymes that in turn control the amount of a sec-ondary messenger produced or that regulate the flow ofions across membranes. Examples of effector moleculesinclude calmodulin, phospholipases, and protein andlipid kinases discussed in the following sections. Gα

subunits also possess GTPase activity, so when the Gα

subunit has completed its job, the GTP is hydrolyzed toGDP and inorganic phosphate. Following hydrolysis ofthe GTP, the Gα subunit dissociates from the effectorprotein and is free to recombine with a Gβγ subunitand thus reform the inactive heterotrimeric complex. Aslong as the GPCR itself remains activated, it can turnon multiple G proteins, thereby amplifying the streamof signals to the effector molecules. The study of G pro-teins in plants is still in its infancy, but they have thus farbeen implicated in signaling responses to gibberellins,brassinosteroids, abscisic acid, and auxin.

17.5 SIGNAL TRANSDUCTIONINCLUDES A DIVERSE ARRAYOF SECOND MESSENGERS

Signal perception is followed by a diverse array of bio-chemical events, referred to as signal transduction orsignaling, that ultimately determines the cell’s responseto that signal. This often complex web of interactingpathways usually involves a variety of small, mobilemolecules known as second messengers. The functionof a second messenger is to relay information from theprimary receptor to the biochemical machinery insidethe cell. Second messengers commonly amplify the orig-inal signal by initiating a cascade of biochemical events.The principal species of second messengers in plantsinclude protein kinase enzymes, calcium ions, and phos-pholipid derivatives.

17.5.1 PROTEIN KINASE-BASEDSIGNALING

Protein kinases are enzymes that activate other pro-teins by catalyzing their phosphorylation. The action ofprotein kinases is balanced by the action of phosphatase

enzymes that deactivate the protein by removing thephosphate group. Protein kinases are able to amplifyweak signals because one active kinase is able to phos-phorylate hundreds of target proteins. Often specificprotein kinases act in series, creating a protein kinasecascade. Protein kinases and kinase cascades have beenimplicated in a wide array of plant signal transductionpathways.

17.5.2 PHOSPHOLIPID-BASEDSIGNALING

Lipids are emerging as an important class of secondmessengers. These second messengers are generatedby the action of enzymes known as phospholipasesthat hydrolyze phospholipids. Recall that phospholipidsare a major constituent of cellular membranes. Fourdifferent phospholipases are known: phospholipase A1(PLA1), phospholipase A2 (PLA2), phospholipase C(PLC), and phospholipase D (PLD). Each of theseenzymes catalyzes the hydrolysis of a specific bond inthe phospholipid molecule as shown in Figure 17.13.Virtually all of the products of phospholipase activ-ity, including free fatty acids, appear to be involved infurther signaling chains.

Lipid-based signaling in plants can be illustrated bythe inositol triphosphate system. In this system, thesignal receptor-complex activates PLC, possibly involv-ing a G protein (Figure 17.14). PLC catalyzes the releaseof inositol triphosphate (IP3) and diacylglycerol (DAG)from the membrane phospholipid phosphatidylinositolbisphosphate (PIP2). Both IP3 and DAG are second mes-sengers. IP3 diffuses into the cytoplasm where it activatescalcium channels and stimulates the release of calciumfrom intracellular stores, most probably from the vac-uole. In plants, DAG remains within the membranewhere it is immediately phosphorylated to phosphatidic

PLDO-

O

OO

PLC

PLA2PLA1

RO = P

H2C

O = C O = C

CH2

HC

O

(CH2)n

CH3 CH3

(CH2)n

FIGURE 17.13 Structure of a ‘‘generic’’ phospholipid,showing the bonds that are subject to phospholipaseactivity. PLA1 and PLA2 remove the fatty acid chainsfrom the sn-1 and sn-2 positions, respectively. PLCremoves the phosphorylated head group (R). PLDremoves only the head group.

17.5 Signal Transduction Includes a Diverse Array of Second Messengers 301

IP3 PA+ DAG

Phospholipase C

Receptor

G protein ?

Signal

Lipid kinase

Membrane

Ion channels Ion channelsEnzyme activation

PIP2

FIGURE 17.14 (A) The generation of second mes-sengers by phospholipase C (PLC). PLC hydrolyzesphosphotidylinositol bisphosphate (PIP2), producinga molecule each of inositol triphosphate (IP3), anddiacylglycerol (DAG). R1 and R2 are fatty acid–basedacyl groups. (B) A model for signaling by the inositoltriphosphate system. The hormone or other externalsignal activates a plasma membrane enzyme phospho-lipase C (PLC), which catalyzes the breakdown of PIP2

to IP3 and DAG. IP3 diffuses into the cytoplasm whereit stimulates the release of calcium. DAG is immedi-ately phosphorylated to phosphatidic acid (PA) by theplasma membrane lipid kinase DAG kinase. PA thendiffuses into the cytoplasm where it may also activateion channels or various effector proteins.

acid (PA). PA then diffuses into the cytoplasm where itregulates ion channels or activates various enzymes.

17.5.3 CALCIUM-BASED SIGNALING

Calcium ions (Ca2) are involved in the regulation ofnumerous physiological processes in plants, includingcell elongation and division, protoplasmic streaming,the secretion and activity of various enzymes, hormoneaction, and tactic and tropic responses. In order for cal-cium to function effectively as a second messenger, thecytosolic Ca2+ concentration must be low and undermetabolic control. Large amounts of calcium are storedin the endoplasmic reticulum, the mitochondria, andthe large central vacuole but the cytosolic Ca2+ con-centration is kept low largely through the action ofmembrane-bound, calcium-dependent ATPases. Activ-ity of the ATPase and, consequently, the cytoplasmicCa2+ concentration, is presumably under control of var-ious stimuli such as light and hormones (Figure 17.15).

Calcium concentration throughout the cell is in partregulated by calcium channels located in the plasmamembrane, endoplasmic reticulum, and vacuolar mem-brane (tonoplast). Some channels, which control theflow of Ca2+ between compartments, are voltage-gated,which means that their opening is determined by aparticular value of membrane potential. Others areregulated either directly by signal receptors or by sec-ond messengers such as inositol triphosphates or cyclicnucleotides. Still others are able to sense tension in themembrane and open in response to turgor or mechanicalstimuli such as touch and wind. Free Ca2+ also diffusesvery slowly in the cytosol—much more slowly than infree solution. Because Ca2+ does not quickly disperse,Ca2+ gradients are easily established and maintained.

The principal calcium receptor in plant and animalcells is calmodulin, a highly conserved, ubiquitousprotein that can be isolated from a variety of higherplants, yeasts, fungi, and green algae. Calmodulin fromseveral plant sources, including spinach, peanut, barley,

corn, and zucchini, has been well characterized, andmany of its properties are similar to calmodulin isolatedfrom bovine brain tissue. Plant and bovine calmodulinhave similar molecular mass (17 to 19 kDa), amino acidcomposition, and calcium-binding properties. When itbinds with calcium, calmodulin undergoes a change inconformation that allows it to recognize and activatetarget proteins.

Several classes of enzymes, including NAD+kinases, protein kinases, and Ca2+-ATPases, are knownto be stimulated by calmodulin. NAD kinase catalyzesthe phosphorylation of NAD to NADP in the presenceof ATP. Because many redox enzymes are specific forone of these cofactors, regulating the balance betweenNAD and NADP is an effective way to regulate meta-bolism. Similarly, as noted above, many other enzymesare activated by protein kinase–catalyzed phospho-rylation. Several calcium-dependent and calcium/

Stimulus

Lighthormonesetc.

ER, Mitochondria, Vacuole

Ca2+

CaM + Ca2+

Ca2+

+ CaM

Ca2+

- ATPase

Ca2+

Enzyme + CaM − Ca2+

(inactive)CaM − Ca2+

+ Enzyme(inactive)

CaM − Ca2+

− Enzyme(active)

FIGURE 17.15 Calcium as a second messenger. Exchangeof calcium between the vacuole and the cytosol may beregulated by hormones or other factors such as light.Cytosolic Ca2+ forms an active complex with calmodulin(CaM) or other calcium-binding protein.

302 Chapter 17 / Growth and Development of Cells

BOX 17.2UBIQUITIN ANDPROTEASOMES—CLEANING UPUNWANTEDPROTEINS

At any given time, a plant cell may contain upward of10,000 or more individual proteins. Many of these pro-teins may no longer be required because the time of theirfunction has passed. Some may contain errors intro-duced during their synthesis or may have been damageddue to excessive heat or some other stress. Others, inparticular certain enzymes, may be short-lived by designbecause they catalyze the first or perhaps rate-limitingstep in a multistep metabolic sequence. Still others maybe key regulatory proteins, such as effectors of the cellcycle, which depend on rapid turnover for maximumeffectiveness. Finally, ubiquitin-mediated degradationof repressor proteins is now known to be a key step inthe regulation of gene transcription by hormones andother developmental signals.

For all of these reasons, proteins are continuallysubject to degradation by proteolytic enzymes calledproteases. Protein degradation is a form of cellularhousekeeping, in which unneeded or damaged proteinsare broken down and their component amino acidsrecycled. Indeed, typically about half the protein com-plement of a cell is replaced every 4 to 7 days. However,proteases cannot be allowed uncontrolled access to cel-lular protein or the result would be chaos. There mustbe some mechanism both for controlling the access ofproteases to protein and for marking or identifying theright proteins for degradation at the appropriate time.

Although protein is degraded in several cellu-lar compartments, including chloroplasts, nuclei,mitochondria, and vacuoles, the process as it occursin the cytosol is best understood. In the cytosol,protein degradation is accomplished by two principalcomponents—a small, highly conserved protein,ubiquitin, and a large, oligomeric enzyme complex,the proteasome (Figure 17.16). The role of ubiquitinis to mark a protein for degradation by forming aconjugate with the target protein and delivering it tothe proteosome where the protein is degraded. Thisis accomplished by the activity of three enzymes:E1, E2, and E3. E1, the ubiquitin-activating enzyme,activates the ubiquitin molecule with ATP. E2, theubiquitin-conjugating enzyme, recruits the ubiquitin toE3, the ubiquitin-protein ligase.

Cullin

Degraded targetprotein

+ free ubiquitin

RBX1SKF1

ATP

Target

Targe

t

Target

E2

ub

ubE1 E1

E2

E2 ub

ub

ub ub

ubub

SCFF-Box

F-Box

FIGURE 17.16 Ubiquitin-mediated protein degradation.Through the action of a large, multienzyme complex,a targeted protein is marked for degradation by tag-ging the protein with ubiquitin. The complex consistsof three enzymes: (1) a ubiquitin-activating enzyme (E1)that forms a covalent bond with ubiquitin through a cys-teine residue; (2) a ubiquitin-conjugating enzyme (E2)that transfers the covalently bound ubiquitin; and (3) aubiquitin-ligating enzyme (E3) that recognizes both thetarget protein and the E2-ubiquitin complex. E3, alsoknown as the SCF complex, is composed of four subunits:SKP1, cullin, RBX1, and an F-box protein. The F-boxprotein recruits the target protein to the SCF complexwhich then catalyzes the transfer of ubiquitin to the tar-get protein. The action of E3 is repeated several times,marking the target protein with a chain of four or moreubiquitin molecules. The ubiquitin-protein complex isthen delivered to a 26S proteasome for degradation. Theproteasome consists of four stacked disks that form ahollow cylinder. Not shown are two 19S regulatory pro-teins that bind to the ends of the stack and direct theubiquitin-protein complex into the hollow, where theprotein-degrading active site is located.

There are several different kinds of ubiquitin-protein ligases. In plants the most common kindappears to be a multimeric protein called the SCFcomplex. The complex is named for the first threesubunits that were discovered: Skp1, cullin, and F-boxprotein. Later a fourth subunit, RBX1, was identified.Skp1, culllin, and Rbx 1 are common to all SCFcomplexes. They, in effect, form a scaffold onto whichdifferent F-box proteins can be assembled, each witha unique substrate-specificity. The Arabidopsis genome,for example contains nearly 700 putative F-box proteins.An F-box protein typically contains a highly conservedrecognition site for the scaffold (the ‘‘F-box’’) and a

17.6 There is Extensive Crosstalk Among Signal Pathways 303

recognition site for a specific target protein which itrecruits to the SCF complex for ubiquitination. At leastsome F-box proteins also contain a recognition site forhormones or other developmental control factors.

Once the F-box protein has delivered the targetprotein to the scaffold, the now complete SCF complexthen facilitates the transfer of the ubiquitin from E2to the target protein. Additional ubiquitin moleculesare added to form a ubiquitin chain. Normally, fourubiqitins are sufficient, at which point the ubiquitinatedtarget protein is delivered to the proteasome.

The actual degradation of the protein is carried outby the proteasome, a complex with a molecular massof more than 1.5 megadaltons (Mda). The proteasome

consists of two parts: a 20S core proteasome and two19Sregulatory complexes.1 Together the three parts makeup the active 26S proteasome. The core proteosome ismade up of four stacked rings with a central channelthat contains the active sites for proteolysis. The regu-latory complex governs access to the channel, probablyunfolding the target protein and injecting it into thechannel for degradation. When a ubiquitinated proteinis delivered to the proteasome, the ubiquitin is releasedto be reused as the target protein is inserted into theproteasome for degradation.1S values refer to the behavior of a complex with respect tosedimentation in a centrifugal field. It is dependent on theshape and density of the complex.

calmodulin-dependent NAD and protein kinases havebeen isolated from both soluble and membrane fractionsfrom a large number of plants.

17.5.4 TRANSCRIPTIONAL-BASEDSIGNALING

There is increasing evidence that some plant signals,such as the light-sensitive pigment phytochrome andsome hormones, may by-pass plasma membrane recep-tors in favor of nuclear receptors and direct interventionin gene expression. They do this by diffusing into thenucleus where they interact with specific transcriptionfactors.

Transcription factors are small, DNA-binding pro-teins that control the transcription of messenger RNAby binding to specific regulatory (i.e., noncoding) DNAsequences in particular genes. Transcription factorsmay either activate (or, up-regulate) the transcriptionof that particular gene, or it may act to repress (or,down-regulate) transcription. Some transcription fac-tors have two binding sites: a DNA-binding site and abinding site for a regulatory molecule. Binding of thesignal molecule, such as a hormone, to the transcrip-tion factor induces changes in the expression of thetarget gene. Exactly how these changes are broughtabout has not yet been determined with any cer-tainty. However, in the case of at least three signalmolecules—phytochrome and the two hormones auxinand gibberellins—the evidence indicates the presenceof a transcription factor that, in the absence of the signal,represses expression of the target gene. When this is thecase, it appears that binding of the signal molecule tothe transcription factor flags the transcription factor fordegradation by the ubiquitin-26S proteasome system

(see Box 17.2). Degrading the repressor allows RNApolymerase to bind with the promoter, thus enablingfull expression of the gene. These pathways will bediscussed more fully in later chapters.

17.6 THERE IS EXTENSIVECROSSTALK AMONGSIGNAL PATHWAYS

Traditionally, signaling pathways were considered asthough each operated as an independent chain of events.For example, the inositol triphosphate pathway wasthought to represent one signal pathway while anotherused protein kinases and yet others used the G-proteinor calcium pathways. As more becomes known aboutsignal transduction pathways, it is increasingly appar-ent that this view of segregated pathways is far toosimplistic. Signaling pathways more closely resemblean interconnected web and established pathways arelinked by many connections, a situation commonlyreferred to as crosstalk. The Oxford English Dictio-nary defines crosstalk as ‘‘unwanted transfer of signalsbetween communication channels’’ but to the biolo-gist crosstalk refers to interactions between and withinvarious classes of signals, the branching and mergingof transduction pathways, and common use of secondmessengers—all of which help to coordinate develop-mental signals. Thus three classes of hormones (auxin,gibberellin, and brassinosteroids) stimulate hypocotylelongation, while two hormones (cytokinin and ethy-lene) and light have an inhibitory effect. The extent ofthese connections is not surprising when one recognizesthat there is a large number of rapidly changing envi-ronmental signals that must be integrated with various

304 Chapter 17 / Growth and Development of Cells

intrinsic control systems in order to execute a finelytuned developmental program.

SUMMARY

The cell cycle describes in effect describes the historyof DNA and chromosomes during cell division. Theactual division of the cell into two daughter cells, orcytokinesis, begins during the final stages of mitosiswhen the microtubules from the mitotic spindle reor-ganize as the phragmoplast. The phragmoplast directsGolgi-derived secretory vesicles to the equatorial planeof the cell where new plasma membranes form and thenew crosswalls are laid down.

The driving force for cell enlargement is wateruptake. However, in order to prevent excessive wateruptake and to avoid rupturing the plasma membranedue to high turgor pressure, plant cells are surroundedby a very strong and relatively rigid wall. In orderfor a cell to enlarge, the strength and rigidity of thewall must be modified. In a turgid cell, the forceof water pressing against the wall generates stresswithin the extensively cross-linked wall components.Growth is initiated when these stresses are relieved bywall-loosening events, which causes the load-bearingcross-links between wall polymers to yield. Relieved ofstress, the wall expands, turgor is reduced, and morewater moves in until both turgor and wall stresses arerestored.

The orderly development of a complex multi-cellular organism is coordinated by a combinationof intrinsic and extrinsic controls. Intrinsic controlsare expressed at both the intracellular and intercellu-lar levels. Intracellular controls are primarily genetic,requiring a programmed sequence of gene expression.Intercellular controls are primarily hormonal, chemicalmessengers that allow cells to communicate with oneanother. Extrinsic controls are environmental cues suchas light, temperature, and gravity. Most environmen-tal cues appear to operate at least in part by modifyinggene expression or hormonal activities.

All developmental stimuli are characterized bya sequence of signal perception, signal transduction,and response. Signal perception requires a recep-tor molecule, which is normally a protein. Receptorsare now known for red and blue light, and most hor-mones. Signal perception involves a diverse array ofinteracting metabolic pathways. Prominent in thesepathways are small, mobile second messengers, suchas the G-proteins, lipid kinases, protein kinases, cyclicnucleotides, calcium ion, and lipids. There is a highdegree of interaction between second messengers,forming a complex web of signal transduction path-ways.

CHAPTER REVIEW

1. Describe the process of cell division in plants.2. What structural characteristics contribute strength

and rigidity to a cell wall?3. What limitations does the cell wall place on the

growth of plant cells?4. Describe the conflicting roles of turgor in the

growth of plant cells.5. Describe the importance of signal perception?

What kinds of signals contribute to the develop-ment of plant cells?

6. What is a receptor protein and what is its role indevelopment?

7. Describe second messengers. What is their role insignal transduction?

8. What is a ‘‘signal cascade’’? How does a signal cas-cade amplify a signal?

9. How does ubiquitin assist in the removal ofunwanted proteins?

10. What is meant by crosstalk (with respectto signal transduction) and what functionmight it serve in plant development?

FURTHER READING

Buchanan, B. B., W. Gruissem, R. L. Jones. 2000.Biochemistry and Molecular Biology of Plants. Rockville,MD: American Society of Plant Physiologists

Chory, J., D. Wu. 2001. Weaving the complex web of signaltransduction. Plant Physiology 125:77–80.

Cosgrove, D. J. 2000. Loosening of plant cell walls byexpansins. Nature 407:321–326.

Cosgrove, D. J. 2001. Wall structure and wall loosening:A look backwards and forwards. Plant Physiology125:131–134.

Karp, G. 2008. Cell and Molecular Biology. 5th ed. New York:John Wiley & Sons.

Moon, J. G. Parry, M. Estelle. 2004. The Ubiqitin-ProteasomePathway and plant development. The Plant Cell16:3181–3195.

Pandy S., J. G. Chen, A. M. Jones, S. M. Assman. 2006.G-protein complex mutants are hypersensitive to abscisicacid regulation of germination and postgerminationdevelopment. Plant Physiology 141:243–256.

Raven, P. H., R. F. Evert, S. E. Eichhorn. 2005. Biology ofPlants. 7th ed. New York: Bedford, Freeman & Worth.

Ryu, S. B. 2004. Phospholipid-derived signaling mediatedby phospholipase A in plants. Trends in Plant Science9:229–235.

Van Leeuwen, W., L. Okresz, L. Bogre, T. Munnik. 2004.Learning the language of plant signaling. Trends in PlantScience 9:378–384.

1 2 3 4

α

18Hormones I: Auxins

Multicellular plants are complex organisms and theirorderly development requires an extraordinary measureof coordination between cells. In order to coordinatetheir activities, cells must be able to communicate witheach other. The principal means of intercellular com-munication within plants are the hormones. Hormonesare signal molecules that individually or cooperativelydirect the development of individual cells or carry infor-mation between cells and thus coordinate growth anddevelopment. Plant hormones have been the subject ofintensive investigation since auxin was first discoveredalmost a century ago.

The discussion of each hormone in this and subse-quent chapters will begin with a review of biosynthesisand metabolism. An understanding of hormone bio-chemistry makes it easier to understand what kinds ofmolecules they are and how they may function. In addi-tion, a lot of what is known about what these moleculesdo and how they do it is based on studies of mutantsthat interfere with their biosynthesis or metabolism.The metabolic turnover of hormone molecules is also asignificant factor in the regulation of cellular activities.

This first of four chapters on plant hormones isdevoted to auxin. The following chapters will covergibberellins, cytokinins, abscisic acid, ethylene, andbrassinosteroids. In the case of each hormone, we willaddress the same three basic questions: what is it, whatdoes it do, and how does it do it?

Because this is the first chapter on hormones, wewill begin with an introduction to the hormone conceptin plants. The balance of the chapter includes

• the biochemistry and metabolism of auxins,• a review of auxin’s principal effects on growth and

development,• how auxin controls cell enlargement,• auxin transport in the plant, and• auxin control of genetic expression.

18.1 THE HORMONE CONCEPTIN PLANTS

The concept of hormones, the chemical messengers thatenable cells to communicate with one another, arose inthe study of mammalian physiology. The latter halfof the nineteenth century witnessed exciting advancesin physiology and medicine. By 1850, it was knownthat blood-borne substances originating in the testisconditioned sexual characteristics. At the same time,physicians pursuing clinical studies had become inter-ested in the effect of glandular extracts and secretionson the course of various diseases. By the turn of the cen-tury, a number of substances that elicited specific effectson the growth and physiology of mammals had been

305

306 Chapter 18 / Hormones I: Auxins

demonstrated and the concept that bodily functionswere coordinated by the production and circulation ofchemical substances was gaining wide acceptance. In1905, the British physician E. H. Starling introducedthe term hormone (Gr., to excite or arouse) to describethese chemical messengers.

Application of the hormone concept to plants maybe traced as far back as the observations of Duhameldu Monceau in 1758. Du Monceau observed the forma-tion of roots on the swellings that occur above girdlewounds that interrupted the phloem tissues around thestems of woody plants. In order to explain these andsimilar phenomena, German botanist Julius Sachs (ca.1860) postulated specific organ-forming substances inplants. Sachs postulated that root-forming substances,for example, produced in the leaves and migrating downthe stem, would account for the initiation of rootsabove the wound. The real beginning of plant hormoneresearch, however, is found in a series of simple butelegant experiments conducted by Charles Darwin (seeBox 18.1). It was Darwin’s observations and experi-ments that ultimately led F. W. Went, almost half acentury later, to describe a hormonal-like substance asthe causative agent when plants grew toward the light.At about the same time, H. Fitting introduced the termhormone into the plant physiology literature.

What are hormones? Hormones are naturallyoccurring, organic molecules that, at low concen-tration, exert a profound influence on physiologicalprocesses. In addition, hormones, as defined by animalphysiologists, are (1) synthesized in a discrete organ ortissue, and (2) transported in the bloodstream to a specifictarget tissue where they (3) control a physiologicalresponse in a concentration-dependent manner. Whilethere are many parallels between animal and planthormones, there are also some significant differences.Like animal hormones, plant hormones are naturallyoccurring organic substances that profoundly influencephysiological processes at low concentration. The siteof synthesis and mode of transport for plant hormones,however, is not always so clearly localized. Althoughsome tissues or parts of tissues may be characterized byhigher hormone levels than others, synthesis of planthormones appears to be much more diffuse and cannotalways be localized to discrete organs.

A hormone can serve effectively as a regulatorysignal only if the molecule has a limited lifetime withinthe target cell. Any molecule sufficiently long-lived to beused repeatedly would sacrifice its dynamic, regulatoryfunction. This means that the amount of a hormone ina cellular pool must be closely regulated and exhibit arate of metabolic turnover that is rapid relative to theresponse that it controls.

The amount of hormone available to a target cellwill be governed primarily by the rates at which activehormone molecules enter (input) and exit (output)

the hormone pool. Hormones may enter the pool by(1) de novo synthesis of the hormone, (2) retrieval ofactive hormone from an inactive storage form, suchas a chemical conjugate, and (3) transport of hor-mone into the pool from a site elsewhere in the plant.Principal means for removing hormone from the poolonce it has acted include: (1) oxidation or some otherform of chemical degradation that renders the moleculeinactive or (2) synthesis of an irreversibly deactivatedconjugate. Clearly, in order to understand the dynamicregulation of hormone activity in plants, it is essentialto know something of these inputs and outputs. Nounderstanding of hormone function can be completewithout a working knowledge of hormone biosynthesisand metabolism.

18.2 AUXIN IS DISTRIBUTEDTHROUGHOUT THE PLANT

Auxin (fr. G. auxein, to increase) is the quintessentialplant hormone. Auxin was the first plant hormone tobe discovered and it has a principal role in the mostfundamental of plant responses—the enlargement ofplant cells. Auxin is synthesized in meristematic regionsand other actively growing organs such as coleoptile

Root

Seed

Coleoptile

0.50 1.0

Relative auxin activity

FIGURE 18.1 Auxin distribution in an oat seedling (Avenasativa), showing higher concentrations of hormone inthe actively growing coleoptile and root apices. (Basedon data from Thimann, K. V. 1934. Journal of GeneralPhysiology 18:23–34.)

18.3 The Principal Auxin in Plants is Indole-3-Acetic Acid (IAA) 307

NH

BOX 18.1DISCOVERINGAUXIN

The experimental beginnings of plant hormone researchin general and auxins in particular can be traced to thework of Charles Darwin. Although Darwin is best knownfor his work on evolution, later in his career he developedan interest in certain aspects of plant physiology. Someof these studies were summarized in the book The Powerof Movement in Plants, co-authored by his son, Francis.One of several ‘‘movements’’ studied by the Darwinswas the tendency of canary grass (Phalaris canariensis)seedlings to bend toward the light coming from a win-dow, a phenomenon we now know as phototropism.The primary leaves of grass seedlings are enclosed ina hollow, sheath-like structure, called the coleoptile,which encloses and protects the leaves as they grow upthrough the soil. Darwin observed that coleoptiles, likestems, respond to unilateral illumination by growingtoward the light source. However, curvature would notoccur if the tip of the coleoptile were either removedor covered in order to exclude light. Since the bend-ing response was observed over the entire coleoptile,Darwin concluded that the phototropic signal was per-ceived by the tip and ‘‘that when the seedlings are freelyexposed to lateral light, some influence is transmittedfrom the upper to the lower part, causing the latter tobend.’’ It was the implications of Darwin’s ‘‘transmis-sible influence’’ that captured the imagination of plantphysiologists and set into motion a series of experimentsthat culminated in the discovery of the plant hormone,auxin—the first plant hormone to be discovered.

Following the publication of Darwin’s book, a num-ber of scientists confirmed and extended their observa-tions. In 1910, Boysen-Jensen demonstrated that thestimulus would pass through an agar block and wastherefore chemical in nature. In 1918, Paal showedthat if the apex were removed and replaced asymmet-rically, curvature would occur even in darkness. In theclimate of the time—Baylis and Starling’s character-ization of animal hormones had appeared only a few

years earlier—plant physiologists were quick to inter-pret these observations as strong support for a planthormone.

The active substance was first successfully isolatedin 1928 by F. W. Went, then a graduate student work-ing in his father’s laboratory in Holland. Following upon the earlier work of Boysen-Jensen and Paal, Wentremoved the apex of oat (Avena sativa) coleoptiles andstood the apical pieces on small blocks of agar. Allowinga period of time for the substance to diffuse from thetissue into the agar block, he then placed each agarblock asymmetrically on a freshly decapitated coleop-tile. The substance then diffused from the block intothe coleoptile, preferentially stimulating elongation ofthe cells on the side of the coleoptile below the agarblock. Curvature of the coleoptile was due to differ-ential cell elongation on the two sides. Moreover, thecurvature proved to be proportional to the amount ofactive substance in the agar. Went’s work was partic-ularly significant in two respects: first, he confirmedthe existence of regulatory substances in the coleop-tile apex, and second, he developed a means for isolationand quantitative analysis of the active substance. BecauseWent used coleoptiles from Avena seedlings, his quanti-tative test became known as the Avena curvature test.Substances active in this test were called auxin, from theGreek auxein (to increase).

The results of Went’s studies naturally stim-ulated intensive efforts to isolate and identify theactive substance. One particularly active compound,indole-3-acetic acid (IAA), was isolated from humanurine in 1934. This peculiar source was selected becauseit was suspected that female sex hormones, secretedin urine, might have some plant growth activity. Ina beautiful piece of scientific serendipity, the impureurine preparation initially assayed was highly active,while subsequently purified hormone preparationswere inactive. This led the investigators back to thematerial from which the female sex hormones wereinitially extracted—the urine of pregnant women—andthe identification of IAA. At the same time, IAA wasisolated from yeast extracts, and the following year,from cultures of Rhizopus suinus. IAA was isolated fromimmature corn kernels in 1946 and since then has beenfound to be ubiquitous in higher plants.

apices, root tips, germinating seeds, and the apical budsof growing stems (Figure 18.1). Young, rapidly growingleaves, developing inflorescences, and embryos follow-ing pollination and fertilization are also significant sitesof auxin synthesis. Auxin, more than any other growthsubstance, appears to be actively distributed throughoutthe entire plant.

18.3 THE PRINCIPAL AUXIN INPLANTS IS INDOLE-3-ACETIC ACID (IAA)

Although a large number of compounds have beendiscovered with auxin activity, indole-3-acetic acid

308 Chapter 18 / Hormones I: Auxins

(IAA) is the most widely distributed natural auxin(Figure 18.2). In addition to IAA, several other natu-rally occurring indole derivatives are known to expressauxin activity, including indole-3-ethanol, indole-3-acetaldehyde, and indole-3-acetonitrile. However,these compounds all serve as precursors to IAA andtheir activity is due to conversion to IAA in the tissue.

The initial discovery of IAA in plants and recogni-tion of its role in growth and development stimulatedthe search for other chemicals with similar activity. Theresult has been an array of synthetic chemicals thatexpress auxin-like activity. One of these chemicals wasindole-3-butyric acid (IBA) (IV, Figure 18.2). Morerecently, IBA has been isolated from seeds and leaves

of maize and several other species. A chlorinated analogof IAA (4-chloroindoleacetic acid, or 4-chloroIAA; II,Figure 18.2) has also been reported in extracts of legumeseeds and a closely related, naturally occurring aromaticacid, phenyl acetic acid (PAA) (III, Figure 18.2) hasrecently been reported to have auxin activity. BecauseIBA, 4-chloroIAA, and PAA have now been isolated fromplants, are structurally similar to IAA, and elicit many ofthe same responses as IAA, there is a strong argument forconsidering them natural hormones. However, it is notyet clear whether they are active on their own or whetherthey are first converted to IAA. Chemically, the singleunifying character of molecules that express auxin activ-ity appears to be an acidic side chain on an aromatic ring.

FIGURE 18.2 The chemical struc-tures of some naturally occurringand synthetic auxins. Indole-3-aceticacid (I) is believed to be the activeauxin in all plants. Phenylacetic acid(III) is widespread and two oth-ers, 4-chlorindole-3-acetic acid andindole-3-butyric acid, have been iden-tified in plant extracts. The latter threeinduce auxin responses when appliedexogenously, but probably act via con-version to IAA. Structures VI, VII, andVIII are active herbicides.

NH

NH

III. Phenylacetic acidNH

Synthetic Auxins

COOH

Cl

O CH3Cl

OCl

Cl

O

ClCl

Cl

Cl

Naturally Occurring Auxins

CH2COOH CH2COOH

I. Indole-3-acetic acid(IAA)

II. 4-Chloroindole-3-acetic acid

CH2COOH

CH2COOH

(CH2)3 COOH

IV. Indole-3-butyric acid (IBA)

V. Naphthalene acetic acid(NAA)

VI. 2-Methoxy-3,6-dichloro- benzoic acid (dicamba)

CH2COOH CH2COOH

VII. 2,4-Dichlorophenoxyacetic acid (2,4-D)

VIII. 2,4,5-Trichlorophenoxy- acetic acid (2,4,5-T)

18.4 IAA is Synthesized from the Amino Acid l-Tryptophan 309

The amount of IAA present will depend on a num-ber of factors, such as the type and age of tissue and itsstate of growth. In vegetative tissues, for example, theamount of IAA generally falls in the range between 1 μgand 100 μg (5.7 to 570 nanomoles) kg−1 fresh weight,but in seeds it appears to be much higher. In one study,it was estimated that the endosperm of a single maizeseed four days after germination contains 308 picomoles(pmole = 10−12 mole) of IAA. At the same time, themaize shoot contained 27 pmoles of IAA and requiredan estimated input of approximately 10 pmoles of IAAhr−1 in order to support its growth. The high level ofIAA in the seed apparently serves to support the rapidgrowth of the young seedling when the seed germinates.

18.4 IAA IS SYNTHESIZED FROMTHE AMINO ACIDl-TRYPTOPHAN

Since the 1930s, when K. V. Thimann first observedthe synthesis of IAA in the mold Rhizopus suinus, whichhad been fed the amino acid tryptophan, the conversionof tryptophan to IAA has been studied in vivo in morethan 20 different plant species and in vitro with atleast 10 different cell-free enzyme preparations. Thesynthesis of IAA is normally studied by feeding plantstryptophan carrying a radioactive label, usually carbon(14C) or tritium (3H), and examining the radioactivity ofsubsequently isolated IAA or its intermediates.

Feeding experiments are complicated by severalfactors and the results must always be approachedwith caution. For example, radiolabeled tryptophancan apparently undergo radiochemical decomposition,thus giving rise to IAA by nonenzymatic reactions. Inaddition, the pool size of tryptophan (also a precursorfor protein synthesis) is very large relative to that ofIAA and there is little data on the actual quantity ofIAA synthesized. Finally, care must be taken to ensurethat experiments are conducted under sterile conditions,since many microorganisms readily convert tryptophanto IAA. While these complications make it difficult toascertain the exact pathway that functions in vivo, theavailable evidence clearly establishes that plants are ableto synthesize IAA from tryptophan.

In most plants, synthesis of IAA occurs inthree steps, beginning with the removal of aminogroup on the tryptophan side chain. The product isindole-3-pyruvic acid (IPA) (Figure 18.3). This reactionis catalyzed by tryptophanamino transferase, a widelydistributed multispecific enzyme that appears to act aswell to remove amino groups from structural analogsof tryptophan such as phenylalanine and tyrosine.The second step is the decarboxylation of IPA toform indole-3-acetaldehyde (IAAld). The enzyme thatcatalyzes this step, indole-3-pyruvate decarboxylase,

NH2

O

COOH

NH

COOH

NH

COOH

NH

ONH

L-Tryptophan

Indole-3-pyruvic acid

Indole-3-acetaldehyde

Indole-3-acetic acid (IAA)

1

2

3

FIGURE 18.3 Pathway for tryptophan-dependentbiosynthesis of indole-3-acetic acid. The enzymesinvolved are (1) tryptophan aminotransferase;(2) indole-3-pyruvate decarboxylase;(3) indole-3-acetaldehyde oxidase.

has been described in several plant tissues and cell-freeextracts. Finally, IAAld is oxidized to IAA by aNAD-dependent indole-3-acetaldehyde oxidase.The presence of this enzyme has been demonstratedin a number of tissues, including oat coleoptile. IAAldmay also be reversibly reduced to indole-3-ethanol.

310 Chapter 18 / Hormones I: Auxins

Indole-3-ethanol is active in bioassays using stemsections, but this is probably due to its conversionto IAA in the tissue. Finally, IAA can be reversiblyconverted to IBA by the enzyme indole-3-butyric acidsynthase.

There is some evidence for alternate biosyntheticpathways involving intermediates other than IPA, butthe burden of biochemical evidence indicates that theIPA pathway is the principal pathway for the synthesisof IAA from tryptophan in higher plants. AlthoughIAA-deficient mutants might be expected to providefurther useful information, none have been identified todate. This is perhaps because an IAA deficiency wouldprobably be lethal.

18.5 SOME PLANTS DO NOTREQUIRE TRYPTOPHANFOR IAA BIOSYNTHESIS

Evidence for the biosynthesis of IAA via atryptophan-independent pathway has been obtainedfrom mutants of both maize and Arabidopsis. Seedlingsof the orange pericarp (orp) mutant of Zea mays lack theenzyme tryptophan synthase, which catalyzes the finalstep in tryptophan synthesis (see Figure 18.3). Althoughseeds carrying the orp mutation germinate normally,they do not survive because of their diminished capacityfor tryptophan synthesis. The IAA content of mutantseedlings, however, is as much as 50-fold higher thanthat of wildtype seedlings. Several tryptophan-requiringmutants have also been isolated from Arabidopsis. Twoof these mutants, trp2 and trp3, also lack tryptophansynthase and are unable to convert indole-3-glycerolphosphate to tryptophan. The trp2 and trp3 seedlings,unlike orp, do not accumulate free IAA but they docontain elevated levels of conjugated IAA (see below).Apparently, trp2 and trp3 store excess IAA in theconjugated form. Radioisotope-labeling experimentsin both maize and Arabidopsis have confirmed that theIAA is synthesized from some precursor other thantryptophan.

The precise pathway for tryptophan-independentIAA synthesis is not known. However, the trp2 and trp3Arabidopsis mutants do accumulate indole-3-acetonitrile.Arabidopsis also contains the nitrilase enzymes necessaryfor converting indole-3-acetonitrile to IAA, thusimplicating indole-3-acetonitrile as an intermediate.The source of indole-3-acetonitrile is not known,although its accumulation in tryptophan mutantssuggests a tryptophan-independent pathway for thebiosynthesis of indole-3-acetonitrile as well. It isknown that indole-3-acetonitrile can be derived fromglucobrassicin, the principal glucosinolate presentin members of the family Cruciferae. Details of thetryptophan-independent indole-3-acetonitrile pathway

for auxin biosynthesis and whether it is limited toArabidopsis or the brassicas, or is more widespread,remain to be determined.

18.6 IAA MAY BE STORED ASINACTIVE CONJUGATES

Very early in the study of auxins, two populations ofthe hormone were recognized—one was free-movingand could be obtained by diffusion into agar; the otherappeared to be bound in the cell and could be isolatedonly by extraction with solvents or by hydrolysis underalkaline conditions. This latter population, referred toas ‘‘bound auxin,’’ is now recognized as IAA that has

N

Indole-3-acetyl-L-aspartate

C N C

O

COOH

COOHH

CH2

CH2

B. Irreversible Deactivation

N

Glucobrassicin

C S

HOOH

O

HO

NOSO3–

CH2OHCH2

N

C O

O

OHHO

OH

OHOH

Indole-3-acetyl-myo-inositol

CH2

A. Reversible Deactivation

FIGURE 18.4 Example of IAA conjugates. Conjugationties up the side-chain carboxyl group, which is essen-tial for auxin activity. Normally, conjugation with asugar reversibly deactivates the auxin molecule while de-activation by conjugation with an amino acid is irre-versible.

18.8 Auxin is Involved in Virtually Every Stage of Plant Development 311

formed chemical conjugates with sugars to form glyco-syl esters. Conjugates are formed by esterification ofa glucose or inositol molecule to the acid group of theside chain (Figure 18.4). IAA-glycosyl conjugates arethemselves inactive but they do release free, biologicallyactive IAA upon solvent extraction, alkaline hydrolysis,or enzymatic hydrolysis in vivo.

Although quantitative data are lacking for mostplants, large pools of IAA glycosyl esters have beendemonstrated in seeds of Zea mays. These pools ofIAA conjugates are formed in the milky endospermas the seed develops and appear to be an importantsource of active hormone for the embryo during thefirst few days of germination. It has been estimated,for example, that as much as 60 percent of the IAArequirement of a germinating maize shoot may be metby hydrolysis of IAA conjugates initially supplied bythe endosperm. Since most of our knowledge of IAArelease by hydrolysis of conjugates comes from studieswith germinating seeds, it is not yet known whetherconjugate hydrolysis is equally important in the growthof mature plants.

18.7 IAA IS DEACTIVATED BYOXIDATION ANDCONJUGATION WITHAMINO ACIDS

IAA in aqueous solution is relatively unstable and isreadily degraded by a variety of agents, including acids,ultraviolet and ionizing radiation, and visible light, thelatter especially in the presence of sensitizing pigmentssuch as riboflavin. IAA degradation in situ, however,appears primarily due to oxygen and peroxide, eitherseparately or in combination, in the presence of a suitableredox system.

Inactivation of the Avena growth-promoting sub-stance by aqueous extracts of leaves was first reportedin the 1930s, even before the active principle was iden-tified as IAA. An enzyme responsible for inactivatingIAA was first isolated from plant extracts in the 1940sand was called IAA oxidase. Later, the enzyme per-oxidase, in concert with a flavoprotein, was shownto catalyze the oxidation of IAA while at the sametime releasing CO2. The oxidative decarboxylation ofIAA by peroxidase is now known to be synonymouswith IAA oxidase. In vitro oxidative decarboxylationof IAA has been studied most extensively with purifiedhorseradish peroxidase. Because the end products of IAAoxidation are physiologically inactive, IAA oxidation isan effective way of removing the hormone moleculeonce it has accomplished its purpose. More recentstudies with green tomato fruits, Vicia faba, and otherspecies have shown that conjugation of IAA with amino

acids such as alanine or aspartic acid also leads to irre-versible deactivation (Figure 18.4).

18.8 AUXIN IS INVOLVED INVIRTUALLY EVERY STAGEOF PLANT DEVELOPMENT

Auxins are characterized principally by their capacity tostimulate cell elongation in excised stem and coleoptilesections, but they are also involved in a host of otherdevelopmental responses, including secondary root ini-tiation, vascular differentiation, and the developmentof axillary buds, flowers, and fruits. Auxins are also animportant component in the signal chain that enablesroots and shoots to respond to gravity and unilaterallight. In fact, auxin is involved in virtually every stage ofplant growth and development from the organization ofthe early embryo to flowering and fruit development.

18.8.1 THE PRINCIPAL TEST FORAUXINS IS THE STIMULATIONOF CELL ENLARGEMENT INEXCISED TISSUES

Regulation of cell enlargement in Avena coleoptiles wasthe basis for its discovery and this action has beendemonstrated repeatedly with excised plant tissues suchas subapical coleoptile tissues and stem segments cutfrom dark-grown pea seedlings.

Auxin concentration-response curves typically showan increasing response with increasing concentrationsof auxin until an optimum concentration is reached(Figure 18.5). Concentrations exceeding the optimumcharacteristically result in reduced growth. If the auxinconcentration is high enough, growth may be inhibitedcompared with controls.

Another characteristic feature of auxin physiologyis that intact stems and coleoptiles do not show a signifi-cant response to exogenous application of the hormone.Apparently the endogenous auxin content of intact tis-sues is high enough to support maximum elongation andadded auxin has little or no additional effect. Thus, itis a general rule that the effect of exogenously suppliedauxin on cell enlargement can be demonstrated only intissues that have been removed from the normal auxinsupply. These include excised segments of stems andcoleoptiles or tissues cultured on artificial media.

18.8.2 AUXIN REGULATES VASCULARDIFFERENTIATION

In addition to stimulating cell enlargement, auxin alsohas a role in regulating cellular differentiation. The mostextensively studied system is the induction of vasculardifferentiation in shoots, which is under control of auxin

312 Chapter 18 / Hormones I: Auxins

IAA Concentration (mg L−1)

Gro

wth

(m

m)

01.0

1.5

2.0

2.5

3.0

3.5

4.0

Float segmentson test solution

1 2 3 4

α

+IAA

−IAA

0.01 0.1 1.0 10

D.

IAA Concentration (mg L−1)

Cur

vatu

re (

degr

ees)

0

5

10

15

20

0.1 0.2 0.3 0.4

C.

B.A.

L

L

FIGURE 18.5 Concentration response curves for two classic auxin-regulatedresponses. (A) Went’s Avena curvature test. A small cube of agar containing auxinis placed on the cut surface of a decapitated oat coleoptile. The auxin diffuses intothe coleoptile, stimulating growth of the cells below the agar cube. The differen-tial growth causes the coleoptile to curve away from the block. (B) Curvature inthe Avena test is linearly related to auxin concentration. (Redrawn from the dataof Went, F. W., K. V. Thimann. 1937. Phytohormones. By permission of K. V. Thi-mann.) (C) Pea stem segment test. Stem sections from dark-grown pea seedlings arefloated on a medium with or without auxin. (D) Typical concentration-response in apea stem section test. Note auxin concentration is expressed on a logarithmic scale.(Redrawn from the data of Galston, A. W., M. E. Hand. 1949. American Journal ofBotany 36:85–94. With permission of the American Journal of Botany.)

produced in the young, rapidly developing leaves. Theproduction of xylem strands at the base of a Coleuspetiole, for example, is directly proportional to thestream of diffusible IAA moving through the petiole.Defoliation of Coleus epicotyls strongly reduces xylemdifferentiation in the petiole, but this effect can bereversed by applying equivalent amounts of IAA inlanolin paste.

A favorite system for the study of vascular dif-ferentiation is the regeneration of vessels and phloemsieve tubes around wounds in Coleus stems, which isalso under the control of auxin (Figure 18.6). Coleus,like other members of the mint family (Lamiaceae), hascharacteristic square stems with a vascular bundle at each

corner. If a wedge-shaped incision is made that inter-rupts one of these vascular bundles, parenchyma cells inthe region of the wound will differentiate into new vas-cular elements. These vascular elements will eventuallyreestablish continuity with the original bundle.

The differentiation of both xylem elements andphloem sieve tubes around the wound is limited and con-trolled by auxin supply. This can be shown by removalof leaves (a source of auxin) above the wound, forexample, which reduces vascular regeneration. On theother hand, because auxin moves preferentially downthe stem, removal of leaves below the wound has lit-tle or no effect. Furthermore, the extent of vascularregeneration is directly proportional to the auxin supply

18.8 Auxin is Involved in Virtually Every Stage of Plant Development 313

FIGURE 18.6 IAA-induced xylem regeneration. A longitu-dinal view of regenerated xylem vessel elements arounda wound (W) in a decapitated internode of cucumber(Cucumis sativus). Lanolin containing 0.1 percent IAAwas applied to the upper side of the internode immedi-ately after wounding. Polar regeneration is indicated bythe dense appearance of many xylem tracheary elements(arrow) in the region of the damaged vascular bundleabove the wound. This is the region where the basipetallyflowing IAA would initially accumulate because it wasinterrupted by the wound and forced to find a new path-way around the obstacle. (Magnification: × 60) (Photo-graph courtesy of Prof. R. Aloni, Tel Aviv University).

when exogenous auxin is substituted for the leaves. Ingeneral, differentiation of phloem sieve tubes is favoredby low auxin concentrations (0.1% IAA w/w in lanolin)while xylem differentiation is favored by higher auxinconcentrations (1.0% IAA w/w in lanolin).

Auxin is also required for vascular differentiationin plant tissue culture. When buds, which are a sourceof auxin, are implanted into clumps of undifferenti-ated callus tissue in culture, differentiation of callus

parenchyma into vascular tissue occurs in regions adja-cent to the implant. The same effect is achieved whenagar wedges containing IAA and sugars are substitutedfor the implanted bud.

18.8.3 AUXIN CONTROLS THEGROWTH OF AXILLARY BUDS

As a shoot continues to grow and the apical meristemlays down new leaf primordia, small groups of cellsin the axil (the angle between the stem and the leafprimordium) of the primordia become isolated from theapical meristem and produce an axillary bud. In somecases, such as the bean (Phaseolus), the bud continues togrow, although at a much slower rate than the apicalbud. In many plants, however, mitosis and cell expansionin the axillary bud is arrested at an early stage and thebud fails to grow. It has been known for some timethat removal of the shoot apex, a common horticulturaltechnique for producing bushy plants, stimulates theaxillary bud to resume growth (Figure 18.7). Apparentlythe apical bud is able to exert a dominant influence thatsuppresses cell division and enlargement in the axillarybud. For this reason, the phenomenon of coordinatedbud development is known as apical dominance.

Shortly after auxin was first discovered, K. V.Thimann and F. Skoog questioned whether there mightbe a relationship between the capacity of the shoot tipto release auxin and its capacity to suppress axillary buddevelopment—in other words, is apical dominance con-trolled by auxin? Thimann and Skoog tested this ideaby decapitating broad bean (Vicia faba) plants and apply-ing auxin to the cut stump. Axillary bud developmentremained suppressed in the presence of auxin. Since thisinitial demonstration, the capacity of auxin to substitute

FIGURE 18.7 Apical dominance in broadbean (Vicia faba).(Left) Control plants. (Center) Removal of the stemapex, a source of auxin, promotes axillary bud growthat the base of the young stem. (Right) Dominance can berestored by applying auxin (in lanolin paste) to the cutstem surface.

314 Chapter 18 / Hormones I: Auxins

OCl

Cl

CH2COOHBOX 18.2COMMERCIALAPPLICATIONSOF AUXINS

Hormones and other regulatory chemicals are now usedin a variety of applications where it is desirable forcommercial reasons to control some aspect of plantdevelopment.

The synthetic auxins are used in commercial appli-cations largely because they are resistant to oxida-tion by enzymes that degrade IAA. In addition totheir greater stability, the synthetic auxins are oftenmore effective than IAA in specific applications. Oneof the most widespread uses of auxin encounteredby the consumer is the use of 2,4-D in weed con-trol. 2,4-D and other synthetic compounds, such as2,4,5-T and dicamba, express auxin activity at low con-centrations, but at higher concentrations are effectiveherbicides.

The introduction of 2,4-D and 4-chlorophen-oxyacetic acid (4-CPA) as herbicides in 1946 revo-lutionized our approach to agriculture. For reasonsthat are not clear, chlorinated phenoxyacetic acids areselectively toxic to broadleaf species. 2,4-D remainsthe principal component of ‘‘weed-and-feed’’ mixturesfor home lawn care as well as for control of broadleafweeds in cereal crops. The synthetic auxins are favoredin commercial applications because of their low costand greater chemical stability.

Indolebutyric acid and naphthaleneacetic acid areboth widely used in vegetative propagation—the prop-agation of plants from stem and leaf cuttings. Thisapplication can be traced to the propensity for auxin tostimulate adventitious root formation. Generally mar-keted as ‘‘rooting hormone’’ preparations, the auxins,usually a synthetic auxin such as NAA or IBA, are mixedwith an inert ingredient such as talcum powder. Stemcuttings are dipped in the powder prior to planting in amoist sand bed in order to encourage root formation.

4-CPA may be sprayed on tomatoes to increaseflowering and fruit set while NAA is commonly used toinduce flowering in pineapples. This latter effect is actu-ally due to auxin-induced ethylene production. NAA isalso used both to thin fruit set and prevent preharvestfruit drop in apples and pears. These seemingly oppositeeffects are dependent on timing the auxin applicationwith the appropriate stage of flower and fruit develop-ment. Spraying in early fruit set, shortly after the flowersbloom, enhances abscission of the young fruits (again,due to auxin-induced ethylene production). Thinning isnecessary in order to reduce the number of fruits andprevent too many small fruits from developing. Sprayingas the fruit matures has the opposite effect, preventingpremature fruit drop and keeping the fruit on the treeuntil it is fully mature and ready for harvest.

The use of synthetic auxins, especially the chlori-nated forms, as herbicides has come under close scrutinyby environmental groups because of potential healthhazards. 2,4,5-T, for example, has been banned inmany jurisdictions because commercial preparationscontain significant levels of dioxin, a highly carcinogenicchemical.

for the shoot tip in maintaining apical dominance hasbeen confirmed repeatedly.

How does auxin from the shoot apex suppress axil-lary bud development? The most widely accepted theoryholds that the optimum auxin concentration for axillarybud growth is much lower than it is for the elongationof stems. The stream of auxin flowing out of the shootapex toward the base of the plant is thought to maintainan inhibitory concentration of auxin at the axillary bud.Removal of this auxin supply by decapitation reducesthe supply of auxin in the region of the axillary budand thereby relieves the bud of inhibition. More directevidence for the role of auxin transport is offered bythe observation that inhibitors of auxin transport (TIBAand NPA) stimulate release of buds from dominancewhen applied to the stem between the shoot apex andthe bud. In addition, lines of tomato that exhibit prolificbranching (that is, the absence of apical dominance)also fail to export radioactively labeled IAA from theshoot apex.

18.9 THE ACID-GROWTHHYPOTHESIS EXPLAINSAUXIN CONTROL OF CELLENLARGEMENT

Whatever its primary action, auxin can alter the rateof cell expansion only by ultimately influencing oneor more of the parameters previously identified inequation 17.1 (Chapter 17). An increase in growth rate,for example, would require an increase in wall exten-sibility (m), an increase in turgor pressure (P), or adecrease in yield threshold (Y). (Hydraulic conductance,L, of the plasma membrane depends on the presenceof aquaporins and is not normally a limiting parame-ter.) Direct measurements of P, using a micropressureprobe, have indicated that turgor pressure does notchange significantly during auxin-stimulated increase inthe growth rate of pea stem sections. Although Y cannotbe measured directly, the results of indirect tests indicate

18.9 The Acid-Growth Hypothesis Explains Auxin Control of Cell Enlargement 315

that yield threshold does not change either. That leavesextensibility, m. Extensibility is difficult to assess. It is onthe one hand a rate coefficient, but it is also a measure ofthe capacity of cell walls to undergo irreversible (plastic)deformation. A number of tests to measure extensibilityhave been devised. Whichever the method, however, theanswer is invariably the same—the induction of rapidcell enlargement by auxin is accompanied by a large andrapid increase in wall extensibility.

The role of low pH in cell enlargement was intro-duced in Chapter 17. At the same time that thisrelationship between acid pH and cell enlargement wasbecoming clear, it was also discovered that auxin wouldcause growing cells to excrete protons. Several linesof evidence indicate that proton secretion is centralto auxin-enhanced cell enlargement. (1) With Avenacoleoptiles the pH of the apoplastic, or cell wall, solu-tion drops from 5.7 to 4.7 within 8 to 10 minutes ofauxin application. This lag period is consistent withthe lag period observed between auxin addition and thebeginning of the growth response. (2) Auxin-stimulatedproton secretion is an energy-dependent process inhib-ited by both metabolic inhibitors and inhibitors ofauxin-induced growth. (3) If the wall space of coleoptilesections is infiltrated with neutral buffers to preventpH change, auxin-induced growth is almost completely

prevented. (4) Agents other than auxin that cause pro-ton excretion have an effect similar to auxin on thepromotion of growth. One such agent is fusicoccin, aphytotoxin from the fungus Fusicoccum amygdali, whichcauses cells to excrete protons at a great rate.

In 1970, R. Cleland and D. Rayle proposed asimple but rather provocative theory to explain auxin-stimulated increases in cell wall extensibility. They sug-gested that auxin causes acidification of the cell wall envi-ronment by stimulating cells to excrete protons. Therethe lower pH activates one or more wall-looseningenzymes, which have an acidic pH optimum. Atabout the same time, A. Hager, working in Germany,published a similar proposal but went further to suggestthat auxin stimulated proton excretion by activating aplasma membrane–bound ATPase proton pump. Thecombined Cleland-Hager proposals are known as theacid-growth hypothesis. Although the acid-growthhypothesis has been tested in relatively few tissues (it hasbeen tested thoroughly only in Avena coleoptiles), theevidence is generally supportive. In its present form, theacid-growth hypothesis proposes that auxin activatesATP-proton pumps located in the plasma membrane(Figure 18.8A). The resulting acidification of the cellwall space lowers the pH toward the optimum range forexpansin activity. Increased expansin activity, in turn,

(1) (2) (3)

CytoplasmTurgor

Membrane

Cytoplasm

A.

B.

(active)

MembraneDockingprotein PLA PK

phospholipid

ABP1

Dockingprotein

ABP1

IAA

Cell wall

H+

AUXIN

ATP ADP + Pi

H+

ATP ADP + Pi

FA + LPC

FIGURE 18.8 A schematic demonstrating the role of auxin in the acid-growth hypoth-esis for cell enlargement. (A) Cell wall polymers (cellulose microfibrils) are exten-sively cross-linked with load-bearing xyloglycans (1), which limits the capacity ofthe cell to expand. An auxin-activated ATPase-proton pump located in the plasmamembrane acidifies the cell wall space by pumping protons from the cytoplasm.The lower pH activates wall-loosening enzymes, such as extensins, that loosen theload-bearing bonds (2). The forces of turgor acting on the membrane and cell wallcause the polymers to displace (3) and allow the cell to enlarge. (B) A hypothetical sig-nal transduction chain linking auxin with activation of the ATPase-proton pump. Seetext for details. Abbreviations: ABP1, auxin-binding protein 1; PLA, phospholipaseA2; FA, fatty acids; LPC, lysophospholipid; PK, protein kinase.

316 Chapter 18 / Hormones I: Auxins

increases wall extensibility and allows for turgor-inducedcell expansion as described earlier in Chapter 17.

Although auxin does enhance the activity ofATPase-proton pumps in the plasma membrane,auxin itself does not bind to the ATPase. Therefore,there must be an auxin receptor that initiates a signaltransduction chain linking the presence of auxin withincreased ATPase activity. A putative auxin receptorhas been isolated from maize (Zea mays), but details ofthe signal transduction chain itself remain obscure.

The maize auxin receptor is membrane-associatedprotein designated ABP1 (Auxin-Binding Protein 1).ABP1 is a 43 kDa glycoprotein dimer of 22 kDa subunitsthat has a high affinity for IAA. ABP1 has been local-ized primarily in the endoplasmic reticulum, but smallpopulations are also found associated with the plasmamembrane and in the cell wall. ABP1 is a prime candi-date for the auxin receptor that mediates cell elongation,although the evidence for this role is indirect. Perhapsthe most compelling evidence comes from experimentswith antibodies. Antibodies are proteins produced bythe immune system of an animal in response to the pres-ence of antigens. Antibodies will bind with the antigen,usually a ‘‘foreign’’ protein, to render that protein inac-tive. Antibodies (designated IgG) can be raised againstplant proteins by injecting purified protein into an ani-mal such as a mouse or rabbit. Antibodies are a usefultool because of the specificity of the antibody–antigenreaction. Antibodies can also be ‘‘tagged’’ with fluores-cent chemicals or other markers so that their locationcan be readily visualized by microscopy. Antibodiesraised against the auxin-binding protein (designatedIgG-antiABP) specifically inhibit both auxin-inducedcoleoptile elongation and auxin-induced hyperpolar-ization of the plasma membrane. Also, IgG-antiABPapplied to coleoptile sections was localized in the outerepidermal cells, which are believed to be the mostauxin-responsive cells in the coleoptile.

The suggestion that ABP1 is the auxin-receptorhas attracted some controversy. The principal diffi-culty has to do with the location of ABP1 in thecell. ABP1 is found predominantly in the lumen ofthe endoplasmic reticulum (ER) and some investiga-tors have been unable to detect any ABP1 at theplasma membrane. ABP1 even contains amino acidsequences at either end of the molecule that are typ-ical of proteins normally retained within the lumenof the ER. However, more sensitive immunolocaliza-tion techniques have now confirmed a small population(perhaps 1000 molecules) on the plasma membrane ofmaize protoplasts. A second problem is that, based onamino acid sequence, the ABP1 protein appears to haveno lipophilic membrane-spanning domain. To recon-cile these observations, it has been proposed that ABP1forms a complex with a transmembrane docking protein.According to this model, the docking protein provides

the necessary lipid solubility to anchor ABP1 to themembrane. The ABP1-docking protein complex is thenexported from the ER to the plasma membrane where itis inserted with ABP1 facing the outside (Figure 18.8B).It has been proposed that the ABP1-docking proteincomplex is itself inactive, but attachment of an auxinmolecule activates the complex and initiates the signaltransduction pathway. The proposed docking proteinhas yet to be identified, but there is some suggestionthat it might be a GCPR receptor in the family ofG-proteins (Chapter 17).

Auxin also activates the enzyme phospholipase A2(PLA2) and several experiments have implicated PLA2in the signal transduction chain. For example, activa-tion of PLA2 can be blocked by IgG-antiABP. Also,both lysophospholipids and fatty acids (the productsof PLA2) stimulate proton secretion and elongation.These effects are inhibited by vanadate, which specif-ically blocks the plasma membrane proton-ATPase.These data suggest that PLA2 follows ABP1 in thechain and that lysophospholipids and fatty acids appearfurther along. Finally, both the IAA and lysophospho-lipids effects on proton secretion and elongation canbe blocked by protein kinase inhibitors, suggesting thatthe lipids activate the proton-ATPase with the involve-ment of a protein kinase cascade. A model illustratinghow these components might interact is presented inFigure 18.8B.

18.10 MAINTENANCE OFAUXIN-INDUCED GROWTHAND OTHER AUXINEFFECTS REQUIRES GENEACTIVATION

The acid-growth hypothesis does not alone resolve thequestion of how auxin regulates cell growth, let alonemore complex developmental problems such as cellmaturation and differentiation. One difficulty is thatgreen stem sections, which respond to auxins, do notrespond well (if at all) to acids. Another difficulty isthat exogenous acid induces only a transitory growthstimulation of coleoptiles. Neither acid nor fusicoccin iseffective after the first 30 to 60 minutes. Auxin-inducedgrowth kinetics show an initial rapid increase in thegrowth rate that reaches a maximum within 30 to60 minutes. This initial burst is followed by a steadyor gradually declining rate over the next 16 hours(Figure 18.9). The most plausible explanation for sucha two-phase response curve is that the acid-growthresponse is limited primarily to the rapid initial growthresponse. Additional auxin-regulated factors must thenbe required for the maintenance of growth over thelonger term, including the well-defined progressionof cells through the sequence division → expansion

18.11 Many Aspects of Plant Development are Linked to the Polar Transport of Auxin 317

Gro

wth

rat

e (μ

m m

in–1

)

125

100

75

50

25

0

Time (min.)

0 20 40 60 80 100 120

80 min.

5 min.IAA

FIGURE 18.9 Kinetics of auxin-induced elongation ofmaize (Zea mays) coleoptiles. The two curves differ inthe duration of auxin action. In each case, auxin (10−5 MIAA) was added at time = 0 and removed after the indi-cated period (5 or 80 min). (From Dela, Fuente, R. K.,A. C. Leopold. 1970. Plant Physiology 46:186. CopyrightAmerican Society of Plant Physiologists.)

→ maturation → differentiation. These additional fac-tors involve the transcription of genes and synthesis ofgrowth-promoting proteins.

Auxin rapidly and specifically stimulates the tran-scription of a set of genes known as primary auxinresponsive genes. These include both SAUR (smallauxin upregulated RNAs) and AUX/IAA. SAUR genesencode short, relatively unstable RNA transcripts. Insoybean hypocotyls, the expression of SAUR genesappears to be localized in tissues that normally respondto auxin and the RNA transcripts can be detectedwithin 2 to 3 minutes of auxin application—even beforeauxin-induced elongation can be observed. Further-more, an asymmetrical distribution of SAUR transcriptshas been detected in gravity-stimulated seedlings. Theasymmetry correlates with the differential cell elonga-tion in responding seedlings, but can be detected evenbefore any visible signs of curvature. Finally, severalauxin-resistant mutants in Arabidopsis show low levels ofSAUR expression in response to auxin treatment.

AUX/IAA genes are induced over a period of 4 to30 minutes following application of auxin. This is alarge family of genes—there are at least 29 differ-ent AUX/IAA genes in the Arabidopsis genome—thatfunction as transcriptional regulators. The AUX/IAAproteins do not bind directly with DNA, but exerttheir regulatory effect by interacting with other proteinscalled the auxin response factor (ARF). ARFs bind tothe promoter region of auxin-responsive genes and mayact either to activate or to repress gene expression. Since

AUX/IAA proteins repress the activity of ARFs, theymay act as either positive or negative regulators.

Early studies indicated that many of these respon-sive genes could also be induced by the protein synthesisinhibitor cycloheximide. This observation suggests thatthese genes may be controlled by short-lived repressorproteins that normally prevent transcription. Accord-ing to one model, auxin was thought to initiate theubiquitin-mediated degradation of these repressor pro-teins. This model was confirmed with the discovery ofthe TIR1 gene in Arabidopsis. Originally identified in agenetic screen for auxin transport inhibitors (hence itsname Transport Inhibitor Response 1), it was soon shownthat TIR1 is a soluble, nuclear-located auxin-receptorprotein that works in conjunction with auxin to derepressthe transcription of auxin-responsive genes.

TIR1 is an F-box protein (see Chapter 17, Box 17.3,for the role of F-box proteins). However, in additionto having a recognition site that allows it to bind withthe SCF scaffold, TIR1 also has a recognition site forauxin. A recent study of the crystal structure of TIR1 hasshown that on the surface of the protein there is a pocketthat accommodates the AUX/IAA peptide. However, theaffinity of TIR1 for AUX/IAA is very low unless an auxinmolecule is also present. The auxin molecule sits at thebottom of the pocket where it simultaneously interactswith both proteins. Auxin thus serves as a ‘‘molecu-lar glue’’ that enhances TIR1-AUX/IAA binding. Onceboth the auxin and AUX/IAA proteins are in place, TIR1is then able to link with the SCF complex for subse-quent ubiquitination and degradation of the repressor bythe ubiquitin-26S proteasome pathway (Figure 18.10).Removal of the AUX/IAA repressor protein derepressesthe auxin-responsive gene, allowing the gene to proceedwith the transcription of messenger RNA and, as a con-sequence, translation of auxin-induced proteins. Auxinappears to modulate development through depression ofauxin-responsive genes, not through a simple activation.

As a side note, it is interesting that the crystal-lographic studies now make it easier to answer thelong-standing question—‘‘What makes an auxin?’’Essentially any molecule that fits into the TIR1binding pocket to enhance TIR1-AUX/IAA interactionwill qualify as an auxin. The relative effectiveness ofdifferent auxin molecules depends on how well they fitin the pocket.

18.11 MANY ASPECTS OF PLANTDEVELOPMENT ARELINKED TO THE POLARTRANSPORT OF AUXIN

Auxin transport has naturally been studied almost exclu-sively in young seedlings, where synthesis takes placein the actively proliferating tissues. From these regions,

318 Chapter 18 / Hormones I: Auxins

Cullin

(High auxin levels)Derepression

Repression(low auxin levels)

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AA

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A

Aux/IAA

Aux/IAA

Aux-IAA

Rbx1

FIGURE 18.10 A model for auxin-induced gene derepres-sion. (1) Auxin response factor protein (ARF) binds tothe DNA in the promoter region of an auxin-responsivegene, but gene transcription is prevented by the presenceof AUX/IAA repressor protein. When auxin levels areelevated, the auxin (A) combines with a nuclear-locatedauxin receptor, TRI1, to form an auxin-TRI1 complex(2). Auxin increases the affinity of TRI1 for AUX/IAA andfacilitates the dissociation of AUX/IAA from the ARF (3).Removal of the AUX/IAA proteins from the ARF dere-presses the gene (4), allowing transcription of mRNAand the translation of auxin-induced proteins, includingAUX/IAA (5). Meanwhile, TRI1 recruits AUX/IAA to theE3 ubiquitin-ligating enzyme, or SCF complex (6), where(7) AUX/IAA is polyubiquitinated. The ubiqitinated pro-tein is then recruited to the 26S proteasome (8), whereit is degraded. The result is that when auxin levels arehigh, TIR1 facilitates active transcription of mRNA bycontinuously removing repressor protein. When auxinlevels are low, TIR1 is unable to bind with the repressor,the repressor protein accumulates, and transcription isshut down.

there appears to be a steady stream of auxin flowingdown the shoot into the root. At least in Arabidopsisseedlings, some of this stream apparently moves downa concentration gradient in the phloem. A significantportion, however, moves through a complex, highlyregulated polar transport mechanism.

Polar transport was originally described based onpreferential movement either up or down in grasscoleoptiles, stems, and roots (Figure 18.11). Whenmovement is away from the morphological apex towardthe morphological base of the transporting tissue, the

Donorblock

Receiverblock

Donorblock

Receiverblock

Donorblock

Receiverblock

Donorblock

Receiverblock

A

B

A

14C - IAA

14C - IAA

B

14C - IAA

14C - IAA

A

AB

Inverted sectionsNormal orientation

B

A

B

FIGURE 18.11 Polarity in auxin transport in an oatcoleoptile segment. The donor block contains 14C-IAA.Regardless of the orientation of the segment, transloca-tion of the radio-labeled IAA is always from the morpho-logically apical end (A) to the morphologically basal end(B) of the segment.

direction of movement is described as basipetal. Move-ment in the opposite direction, toward the morpholog-ical apex, is referred to as acropetal. When a stem orcoleoptile section is inverted, as shown in Figure 18.11,the original direction of movement is maintained. How-ever, as more is learned about auxin transport, the moreevident it becomes that directed auxin transport may belateral as well as up and down.

Polar transport of auxin in shoots tends to be pre-dominantly basipetal at a velocity somewhere between5 and 20 mm hr−1. Acropetal transport in shoots isminimal. In roots, on the other hand, there appear tobe two transport streams. An acropetal stream, arrivingfrom the shoot, flows through xylem parenchyma cellsin the central cylinder of the root and directs auxintoward the root tip. A basipetal stream then reverses thedirection of flow, moving auxin away from the root tip,or basipetally, through the outer epidermal and corticalcell files.

The phenomenon of polar auxin transport hasattracted widespread attention because of the assump-tion that auxin concentration is an important variablein several developmental responses. Auxin gradients dueto polar transport have been invoked to explain, at leastin part, developmental phenomena such as apical dom-inance, adventitious and secondary root formation, anddifferential growth responses to light and gravity. Theflow of auxin in the root, for example, is intimatelyinvolved in the root response to gravity and will becovered in a later chapter.

Several observations indicate that polar trans-port involves a carrier-mediated, active transport

18.11 Many Aspects of Plant Development are Linked to the Polar Transport of Auxin 319

mechanism in both shoots and roots. First, itcan be shown that polar transport is inhibitedby anaerobiosis or by respiratory poisons such ascyanide and 2,4-dinitrophenol. This is consideredevidence that polar transport is an energy-requiringprocess dependent on oxidative metabolism in themitochondrion. Second, certain chemicals, calledphytotropins, have been known for some time to bespecific, noncompetitive inhibitors of polar transport.These include TIBA (2,3,5-triiodobenzoic acid),morphactin (9-hydroxyfluorine-9-carboxylic acid), andNPA (N-1-naphthylphthalamic acid) (Figure 18.12). Itis presumed that such inhibitors block auxin transportby binding to discrete carrier molecules that areinvolved in polar transport system. Third, the uptakeof radioactive IAA is at least partially inhibited bynonradioactive IAA. This latter observation indicatesthat the labeled IAA and unlabeled IAA compete witheach other for a finite number of carrier sites.

These observations formed the basis for thechemiosmotic model for auxin transport, proposed byP. H. Rubery, A. R. Sheldrake, and J. A. Raven in themid-1970s. In its present form, the chemiosmotic modelcontains three essential features: (1) a pH gradientor proton motive force across the plasma membranethat provides the driving force for IAA uptake, (2) anIAA influx carrier, and (3) an IAA efflux carrier that ispreferentially located at the base of auxin-transportingcells (Figure 18.13). The principles of the chemiosmoticmodel may be summarized as follows. IAA is a weaklyacidic, lipophilic molecule. Depending on the pH, IAAmay exist in either the protonated form (IAAH) orthe unprotonated, anionic form (IAA−). The cell wallspace is moderately acidic with a pH of about 5.5. Atthat pH, approximately 20 percent of the IAA will beprotonated (IAAH). Consequently, the cell wall spacewill contain both anionic and protonated IAA. Basedon its lipid solubility, a small proportion of unchargedIAAH molecule would be expected to diffuse slowlyacross the plasma membrane from the cell wall space

COOH

I

2,3,5-Triiodobenzoic acid(TIBA)

II

C NH

C OH

O

O

Naphthylphthalamic acid(NPA)

FIGURE 18.12 Phytotropins. Two examples of inhibitorsof polar IAA transport.

H+

H+

H+

H+

IAAH IAA-

IAAH IAA-

PH5.5

PH7

FIGURE 18.13 The chemiosmotic-polar diffusion modelfor polar transport of IAA. In the acidic cell wall space(pH 5.5) approximately 20% of the IAA is protonated.Protonated IAA (IAAH) may enter cells by diffusionacross the cell membrane (dashed arrows) while theanionic form (IAA−) may be taken up through AUX1 (cir-cles), a proton/IAA symport carrier located randomly inthe plasma membrane. Inside the cell (pH 7.0) the depro-tonated form IAA− will dominate. IAA− can exit the cellonly through efflux carriers of the PIN family (squares)which are located preferentially at the base of the cell.Membrane-bound ATPase-proton pumps help to main-tain the appropriate pH differential across the membraneand provide protons for IAA/H+ symport. The uniquelybasal location of the efflux carriers is the key to polartransport.

into the cell. The bulk of the IAA, however, will enterthe cell as IAA− through an H+/auxin symport carrier(the influx carrier) that is uniformly distributed aroundthe cell.

Once in the cytoplasm, where the pH is closer to7.0, IAAH will dissociate to IAA− and H+. The auxinis now trapped inside the cell because IAA− can notreadily diffuse across the membrane. The key to thechemiosmotic model, however, is the existence of acarrier, located only in the basal membranes of the cell,which mediates the efflux of IAA− from the cell. It isthe unique location of this efflux carrier, more than anyother single factor, which establishes polarity in auxintransport.

The first direct evidence for the existence of a basalefflux carrier took advantage of the fact that the puta-tive IAA-transport protein binds the phytotropin NPA.The NPA-binding protein was isolated, antibodies wereraised against it, and the antibodies subsequently labeledwith the fluorescent dye fluorescein in order to make

320 Chapter 18 / Hormones I: Auxins

the antibodies visible under the microscope. When peastem sections were treated with the labeled antibodies,fluorescein was found to be localized on the basal plasmamembranes of the stem cells.

More recently, largely due to studies of auxinmutants of Arabidopsis, two good candidates for auxininflux and efflux carriers have been identified. Theputative influx carrier is a membrane protein, AUX1.The AUX1 gene has been linked to auxin metabolismand transport because mutations at that locus exhibitIAA-resistant root growth, reduced lateral root initia-tion, and reduced response of the root to gravity. Sucha phenotype is consistent with a reduced capacity totake up IAA. The AUX1 gene has been cloned and thepolypeptide sequence of the protein is similar to that ofknown amino acid permeases. Amino acid permeases aremembrane proteins that function as amino acid/protonsymport carriers. The protein homologies together withthe structural similarities between IAA and its precursoramino acid, tryptophan, have led to the suggestion thatthe AUX1 protein functions as an auxin/proton sym-porter. Further support for this model is offered by theobservation that the synthetic auxin NAA restores thegravitropic response to mutant (aux1) seedlings. NAAuptake by cells is not carrier-mediated, so the loss ofAUX1 does not interfere with the response.

A family of genes, the PIN genes, that encode puta-tive auxin efflux carriers have also been identified. (Atotal of eight PIN genes have now been identified inArabidopsis.) One of the first to be discovered is the PIN1gene that controls flower development in Arabidopsis.The pin1 mutant is characterized by influorescences thatterminate in pinlike structures and show little or no evi-dence of floral bud development. Polar auxin transportis significantly reduced in mutant pin1 influorescencesand the characteristics of the mutant can be mimickedby blocking polar transport with phytotropins. As pre-dicted by the chemiosmotic model, fluorescent-labeledantibodies have demonstrated that PIN1 protein is local-ized in the basal membranes of xylem parenchyma cells.Moreover, in keeping with the chemiosmotic model,AUX1 and PIN1 proteins are located at opposite endsof young root phloem cells (protophloem).

A second gene is variously called PIN2, EIR1,WAV6, or AGR1. The several names are due to thefact that the gene was identified independently in dif-ferent laboratories, all studying mutants with impairedroot response to gravity. PIN2 was so named becauseit encodes a protein that is very similar to the proteinencoded by PIN1. As with the PIN1 protein, immunolo-calization experiments have shown that the PIN2 pro-tein is localized in the basal membranes (i.e., furthestfrom the root tip) of cell files in the root cortex and epi-dermis. Moreover, like AUX1, the structure of the PINproteins resembles the bacterial amino acid permeasesand hence is a likely candidate for an IAA transporter.

Polarity in auxin transport is fundamental to plantdevelopment and PIN proteins direct this transport bymoving from one cell surface to the other, in keepingwith changing demands for auxin asymmetry. One prob-lem, for example, that has long puzzled developmentalbiologists is how the apical-basal axis is established inyoung embryos. PIN proteins appear to part of the key.Immediately after the first division of the zygote, PINproteins located acropetally in the basal cell direct theflow of auxin into the apical cell, specifying that cell asthe founder of the proembryo. As the apical cells pro-liferate to form the globular embryo stage, they beginto synthesize auxin themselves. The PIN proteins thenshift to a basipetal location and the direction of auxinflow reverses, thereby establishing the position of thedeveloping root pole. Similar changing patterns of PINdistribution and the resulting changes in the polarity ofauxin transport are equally important in other responsessuch as secondary root initiation and the responses ofshoot and roots to gravity and unilateral illumination,which will be covered in detail in a later chapter.

SUMMARY

Hormones are numerous naturally occurring chemicalsubstances that profoundly influence, at micromo-lar concentrations, the growth and differentiation ofplant cells and organs. The effectiveness of a hormonedepends on maintaining a closely regulated pool size,which is accomplished by a balance of biosynthesis,storage as inactive conjugates, and catabolic degrada-tion of the molecule.

Auxins are characterized by their capacity to stim-ulate elongation in coleoptile and stem segment but areinvolved in virtually every aspect of plant development,including seed germination, vascular differentiation,lateral bud development, secondary root initiation, theresponse of roots and shoots to gravity, and flower andfruit development.

A large number of synthetic compounds expressauxin activity, but indole-3-acetic acid (IAA) is thoughtto be the only naturally occurring auxin. In most plantsIAA is synthesized from the amino acid tryptophanalthough the study of tryptophan-requiring mutantshas established that in some plants, such as Arabidop-sis, IAA is synthesized via a tryptophan-independentpathway. IAA can be stored as chemical conjugatessuch as glycosyl esters, which will release active IAA onenzymatic hydrolysis. Glycosyl esters are an importantsource of IAA during seed germination. Once IAA hasaccomplished its purpose, it can be removed by peroxi-dation to inactive products or conversion to amino acidconjugates. Auxins may be transported in the phloemor cell-to-cell by polar transport. The key to polar

Further Reading 321

transport is the location of efflux carriers on specificcell walls.

The role of auxin in cell enlargement is bestdescribed by the acid-growth hypothesis. Central tothis hypothesis is the activity of expansins; enzymesthat weaken cross-links between cellulose molecules,increase wall extensibility, and allow for turgor-inducedcell expansion. Auxin also acts to derepress genes bytargeting repressor proteins for degradation by the26S proteasome pathway, a process that accounts forauxin-induced developmental responses.

CHAPTER REVIEW

1. Why is it necessary for a hormone to be rapidlyturned over? Describe how the size of theactive hormone pool is regulated for auxins.

2. Some seeds appear to accumulate auxin conjugates.Can you suggest a physiological advantage for this?

3. Review the synthesis of IAA from tryptophan.Do all plants synthesize IAA from tryptophan?What is the evidence for alternative pathways?

4. The auxin needs of a germinating seed are initiallymet by stored auxin conjugates. Are all hor-mone-conjugates a form of hormone ‘‘storage’’?

5. Describe the auxin-signaling pathway for cellenlargement.

6. Describe the auxin-signaling pathway for control-ling gene expression.

7. Auxin transport is uniquely polar in character.How is this directional transport accomplished?

FURTHER READING

Badescu, G. O., R. M. Napier. 2006. Receptors for auxin:Will it end in TIRs? Trends in Plant Science 11:217–223.

Benjamins, R., N. Malencia, C. Luschnig. 2005. Regulatingthe regulator: The control of auxin transport. BioEssays27:1246–1255.

Blakeslee, J. J., W. A. Peer, A. S. Murphy. 2005. Auxin trans-port. Current Opinion in Plant Biology 8:494–500.

Buchanan, B. B., W. Gruissem, R. L. Jones. 2000.Biochemistry and Molecular Biology of Plants. Rockville,MD: American Society of Plant Physiologists.

Davies, P. J. 2004. Plant Hormones: Biosynthesis, Signal Trans-duction, Action. Dordrecht: Kluwer Academic Publishers.

Kramer, E. M., M. J. Bennett. 2006. Auxin transport: A fieldin flux. Trends in Plant Science 11:382–386.

Napier, R. M. 2005. TIRs of joy: New receptors for auxin.BioEssays 27:1213–1217.

Tan, X. et al. 2007. Mechanism of auxin perception by theTIR1 ubiquitin ligase. Nature 446:640–645.

Woodward, A. W., B. Bartel. 2005. Auxin: Regulation, action,and interaction. Annals of Botany 95:707–735.

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19Hormones II: Gibberellins

Gibberellins are members of a large and varied familyof plant constituents known as the terpenoids. Terpenesare normally recognized on the basis of their chemicalstructure, which may be dissected into one or more5-carbon isoprene units. Gibberellins are thus biosyn-thetically related to the carotenoid pigments, sterols,latex, and other isoprene derivatives that will be dis-cussed later. Gibberellins are noted for their capacityto stimulate hyper-elongation of intact stems, especiallyin dwarf plants and rosettes. They are also promi-nently involved in seed germination and mobilizationof endosperm reserves during early embryo growth, aswell as leaf expansion, pollen maturation, flowering, andfruit development. This chapter describes

• the biochemistry and metabolism of gibberellins,• a review of gibberellins’ principal effects on growth

and development, and• gibberellin receptors and the gibberellin signaling

chain.

19.1 THERE ARE A LARGENUMBER OF GIBBERELLINS

Gibberellins are a large class of molecules. In fact, morethan 135 have now been identified in higher plants andfungi and additional members are added almost every

year. Only a few of these are biologically active in theirown right. The others are either intermediates in thebiosynthetic pathway or products of inactivation. It isworth noting, however, that the number of gibberellinsfound in any one species or organ may be very smalland the number of active gibberellins smaller yet. It isbelieved, for example, that GA1 and GA4 are the prin-cipal naturally occurring, active gibberellins in higherplants.

All gibberellins are diterpenes based on the20-carbon ent-gibberellane structure (Figure 19.1).A little more than one-third of the gibberellinscharacterized to date have retained the full complementof 20 carbon atoms and are known as C20-gibberellins.The others have lost carbon atom number 20 andare consequently known as C19-gibberellins. With acomplex ring structure and the number of possiblesubstitutions on 19 or 20 carbon atoms, it is not difficultto see how there could be such a large number ofgibberellins.

Gibberellins that are demonstrated to be naturallyoccurring and that have been chemically characterizedare assigned an ‘‘A’’ number. This number does notimply chemical relationships; it is assigned roughly inorder of discovery. A C20-gibberellin commonly knownas gibberellic acid was one of the first to be isolatedand characterized. Because GA3 is readily extractedfrom fungal cultures, it is also the most common

323

324 Chapter 19 / Hormones II: Gibberellins

1

2

3

1112

13

15

19

8

71918

CH3

20

H

HO H

CO

O

COOH

OH

HO

AH

C

D H

HO H

CO

O

COOH

OH

ent-Gibberellaneskeleton GA3 Gibberellic acid

C19-GA (Active) C19-GA (Inactive)

H3C CH3

CH3

CH3

CH3

CH2

CH2

CH3

GA1 GA8

H

HO H

CO

O

COOH

OH

CH2

CH3

FIGURE 19.1 The ent-gibberellane skeleton and chemical structures of selected activeand inactive gibberellins. GA8 is inactive because of the addition of the hydroxylgroup in the 2 position.

commercially available form. GA1, GA3, and GA4, all ofwhich promote vegetative growth, are the most activegibberellins and, consequently, the most widely used ingibberellin research.

There are certain structural requirements for GAactivity. A carboxyl group at carbon-7 is a feature ofall GAs and is required for biological activity, andC19-GAs are more biologically active than C20-GAs.In addition, those GAs with 3-β-hydroxylation, 3-β,13-dihydroxylation, or 1,2-unsaturation are generally moreactive; those with both 3-β-OH and 1,2-unsaturationexhibit the highest activity.

19.2 THERE ARE THREEPRINCIPAL SITES FORGIBBERELLINBIOSYNTHESIS

It is generally accepted that there are three principalsites of gibberellin biosynthesis: (1) developing seedsand fruits, (2) the young leaves of developing apicalbuds and elongating shoots, and (3) the apical regionsof roots. Immature seeds and fruits are prominent sitesof gibberellin biosynthesis. This is based on the obser-vation that young fruits, seeds, and seed parts contain

large amounts of gibberellins, particularly during stagesof rapid increase in size. In addition, cell-free prepara-tions from many seeds, such as wild cucumber (Marahmacrocarpus) and pea (Pisum sativum), are able to activelysynthesize gibberellins. The site of gibberellin biosyn-thesis may be the developing endosperm (as it is inthe cucurbits), the young cotyledons of legumes, orthe scutellum of cereal grains. As the seed matures,metabolism appears to shift in favor of gibberellin-sugarconjugates.

It is not as easy to obtain clear evidence that gib-berellin biosynthesis occurs in shoots and roots. Thisis partly because gibberellin levels are much lower invegetative tissues. Vegetative tissues also yield cell-freepreparations that are less active, suggesting that enzymelevels for gibberellin metabolism are also lower thanfor reproductive tissues. Gibberellin synthesis in vege-tative tissues is generally supported by the occurrenceof gibberellins in tissue exudates and the effects ofinhibitors of gibberellin biosynthesis. Gibberellins areapparently derived from fundamental precursors thatare synthesized in chloroplasts (Section 19.3), which isconsistent with the synthesis of gibberellins in greenleaves. On the other hand, application of the inhibitor(2-chloroethyl) trimethylammonium chloride (CCC,an ‘‘antigibberellin’’) to roots rapidly decreases the

19.3 Gibberellins are Terpenes, Sharing a Core Pathway 325

BOX 19.1DISCOVERY OFGIBBERELLINS

During the late nineteenth and early twentieth cen-turies, Japanese rice farmers grew concerned about adisease that seriously reduced the yield of their crops.Plants infected with the bakanae (‘‘foolish seedling’’)disease exhibited weak, elongated stems and producedlittle or no grain. Japanese plant pathologists, interestedin developing means for controlling the disease, soonestablished a connection with the presence of a fungus,Gibberella fujikuroi. In 1926, E. Kurosawa reported theappearance of symptoms of the disease in uninfectedrice plants that had been treated with sterile filtratesfrom cultures of this fungus. By 1938, Japanese investi-

gators had isolated and crystallized the active material,which they called gibberellin after the genus name for thefungus.

Gibberellin did not come to the attention of West-ern plant physiologists until after the 1939–1945 war,when two groups—one headed by Cross in Englandand one by Stodola in the United States— isolated andchemically characterized gibberellic acid from fungal cul-ture filtrates. At the same time, Japanese workers isolatedthree gibberellins, which they named gibberellin A1, gib-berellin A2, and gibberellin A3. Gibberellin A3 provedto be identical with gibberellic acid. The known effectof gibberellins on rice and several other plant systemsindicated that similar substances might be present inhigher plants as well. The first higher-plant gibberellinto be characterized was isolated from immature seeds ofrunner bean (Phaseolus coccineus) and found to be iden-tical with gibberellin A1. Since then, gibberellins havebeen shown to be ubiquitous in higher plants.

amount of gibberellin appearing in exudates. Further-more, removal of the root apices from seedlings of scarletrunner bean (Phaseolus coccineus) results in decreasedshoot growth and the disappearance of GA1. It remainspossible that the synthesis of gibberellin precursors isnot limited to chloroplasts, but occurs in other forms ofplastids as well.

19.3 GIBBERELLINS ARETERPENES, SHARING ACORE PATHWAY WITHSEVERAL OTHERHORMONES AND A WIDERANGE OF SECONDARYPRODUCTS

Terpenes are a functionally and chemically diversegroup of molecules. With nearly 15,000 structuresknown, terpenes are probably the largest and mostdiverse class of organic compounds found in plants.This large diversity arises from the number of basicunits in the chain and the various ways in which theyare assembled. Formation of cyclic structures, addi-tion of oxygen-containing functions, and conjugationwith sugars or other molecules all add to the possiblecomplexity.

The terpene family includes, in addition tothe gibberellins, the hormones abscisic acid andbrassinosteroids, the carotenoid pigments (caroteneand xanthophyll), sterols (e.g., ergosterol, sitosterol,cholesterol) and sterol derivatives (e.g., cardiac

glycosides), latex (the basis for natural rubber), andmany of the essential oils that give plants theirdistinctive odors and flavors. Cytokinin hormones andchlorophyll, although not terpenes per se, do containterpenoid side chains.

The distinguishing feature of terpenes and terpenederivatives is that they may be viewed as polymersof the simple 5-carbon unit 2-methyl-1,3-butadiene,or isoprene (Figure 19.2). Consequently, terpenes areoften referred to as isoprenoid compounds. The actualbuilding blocks for terpenoids, however, are not iso-prene itself, but two phosphorylated derivatives knownas iso-pentenyl pyrophosphate (IPP) and its iso-mer dimethylallyl pyrophosphate (DMAP). IPP isa product of two different biosynthetic pathways, onelocated in the cytoplasm and one located in chloroplasts(Figure 19.3).

The cytoplasmic pathway for IPP synthesis beginswith acetyl-coenzyme A (acetyl-CoA), an interme-diate in the respiratory breakdown of carbohydrateand fatty acid metabolism. Three molecules ofacetyl-CoA condense in a two-step reaction to formhydroxymethylglutaryl-CoA, which is then reducedto mevalonic acid (MVA). Because the reductionof hydroxymethylglutaryl-CoA to mevalonic acid isvirtually irreversible (at least in vitro), mevalonic acid

C CH2CHH2C

CH3

FIGURE 19.2 The 5-carbon isoprene unit is the basicbuilding block of terpenes and terpene derivatives.

326 Chapter 19 / Hormones II: Gibberellins

Isopentenyl Pyrophosphate(IPP)

PC

CH3

CH2H2C CH2 O P

Dimethylallyl Pyrophosphate(DMAP)

PC

CH3

CHH3C CH2 O P

Mevalonic Acid (MVA) (6 carbons)

(3) Pyruvate (6 carbons)

(3) Acetyl-CoA (6 carbons)

Hydroxymethylglutaryl-CoA (6 carbons)

2 NADPH

2 NADP++ CoA

CO−O

CCH3

CH2H2C CH2OH

A.

ATP

ATP

ADP+Pi+CO2

ATP

Mevalonic Acid Pyrophosphate (6 carbons)

FIGURE 19.3 There are two pathways for the synthesis of isopentenylpyrophos-phate (IPP) and dimethylalylpyrophosphate (DMAP). (A) The mevalonic acid (MVA)pathway is located in the cytoplasm and is the source of cytoplasmic IPP. (B) Themethylerythritol-4-phosphate (MEP) pathway is located in the chloroplast. Thepathways are named after the first committed, or irreversible, intermediate.

is the first committed precursor for terpenes and thispathway is therefore known as the mevalonic acidpathway. Mevalonic acid then undergoes a two-stepphosphorylation, at the expense of two moleculesof ATP, to form C6 mevalonic acid pyrophosphate.Removal of a carboxyl group converts mevalonic acidpyrophosphate to the C5 compound IPP, which isreversibly isomerized to DMAP.

The chloroplast pathway begins with the conden-sation of glyceraldehyde-3-phosphate (G3P) and pyru-

vic acid to form deoxyxylulose phosphate, a 5-carbonmolecule. Deoxyxylulose phosphate is then reduced tomethylerythrltol-4-phosphate (MEP). MEP is sub-sequently phosphorylated and oxidized to form IPP.

Like the synthesis of IPP, the subsequent synthesisof terpenes is highly compartmentalized. In general,sesquiterpenes (C15), triterpenes (C30), and polyter-penes such as latex and other large molecules areproduced in the cytosolic compartment while isoprene (a5-carbon volatile organic hydrocarbon), monoterpenes

19.4 Gibberellins are Synthesized from Geranylgeranyl Pyrophosphate (GGPP) 327

G3P + Pyruvate (6 carbons)

P

OH

COH

OHOCH2C

H3CCH2

H

Methyerythritol-4-Phosphate (5 carbons)

Isopentenyl Pyrophosphate(IPP)

PC

CH3

CH2H2C CH2 O P

Dimethylallyl Pyrophosphate(DMAP)

PC

CH3

CHH3C CH2 O P

OHO

H3C C CH

COH

HCH2O P

1-Deoxyxyulose-5-Phosphate (5 carbons)

C02

NADPH

NADP

3ATP

3ADP

+

B.

FIGURE 19.3 (Continued)

(C10), diterpenes (C20), and tetraterpenes (C40) origi-nate in plastids (Figure 19.4). In either case, the synthesisbegins with the isomerization of IPP to DMAP. DMAPis, in effect, a reactive primer molecule that initiateschain formation by condensation with a molecule ofIPP. Elongation of the chain then continues with therepetitive addition of IPP catalyzed by prenyltransferaseenzymes. The distinction between cytosolic and plastidproducts is largely dependent on the compartmental-ization of specific prenyltransferase enzymes. FPP is abranch point that can give rise to other C15 sesquiter-penes and, by head-to-head condensation of two C15units, the C30 triterpenes (e.g., sterols). GGPP is a sec-ond branch point, giving rise to linear C20 diterpenes,cyclic diterpenes (including the gibberellins), and, byhead-to-head condensation of two C20 units, the C40tetraterpenes (e.g., carotenoid pigments).

19.4 GIBBERELLINS ARESYNTHESIZED FROMGERANYLGERANYLPYROPHOSPHATE (GGPP)

The synthesis of gibberellins begins with the C10isoprenoid geranylgeranyl phyrophosphate GGPP.The first two reactions involve the cyclization ofGGPP to form copalylpyrophosphate and thenkaurene (Figure 19.5). These two cyclization stepsare inhibited by the antigibberellin or dwarfingagents, AMO-1618, CCC, and phophon-D, thusleading to a deficiency of gibberellin in the plantand reduced growth. Following cyclization, thecarbon at position 19 on the kaurene moleculeundergoes three successive oxidations in the sequence

328 Chapter 19 / Hormones II: Gibberellins

Geranylgeranyl Pyrophosphate (C20)

Farnesyl Pyrophosphate (C15)

Geranyl Pyrophosphate

OtherSesquiterpenes (C15)

Phytosterols

Brassinosteroids

Plastoquinones

AbscisicAcid

CYTOPLASM CHLOROPLAST

Cytokinins (X3)

CH2OPP

CH2OPP

IPP IPP

(GPP, C10)

(X2)Monoterpenes

Carotenoids(C40)

Chlorophyll(phytol tail)

(C20)

Gibberellins(C20)

(X2)

FIGURE 19.4 Cytoplasmic and plastidic terpene biosynthesis, showing the structuresof the key intermediates farnesyl pyrophosphate and geranylgeranyl pyrophosphate.Terpenes of increasing carbon number are formed by sequential addition of isopen-tenyl pyrophosphate units. Odd numbered terpenes (C15, C30) are synthesized in thecytoplasmic compartment while even numbered terpenes (C10, C20, C40) are synthe-sized in the chloroplast.

CH3→CH2OH→CHO→COOH to form kaurenoicacid. The oxidation of kaurene to kaurenoic acid isinhibited by ancymidol, another dwarfing agent. Thefinal two steps involve a hydroxylation at carbon-7 andcontraction of the B ring with extrusion of carbon-7 toform GA12-7-aldehyde.

The enzymes that convert ent-kaurene to GA12-7-aldehyde are all NADPH-dependent, membrane-boundcytochrome P450 monooxygenases. The cytochromeP450 family is an interesting and important groupof enzymes. They are generally noted for their lackof substrate specificity and are involved in numerousbiosynthetic as well as degradative pathways in plantsand animals. In mammals they are used to detoxify theliver by converting hydrophobic compounds to morehydrophilic derivatives that are more readily excreted.

As noted above, GA12-7-aldehyde is the first com-pound with the true gibberellane skeleton and is the

precursor to all other gibberellins. Oxidation of thealdehyde group on carbon-7 to a carboxyl group givesGA12. This carboxyl group is a feature of all GAs and isrequired for biological activity. C19-GAs arise by sub-sequent oxidative elimination of carbon-20. While thebiosynthetic pathway up to GA12-7-aldehyde is the samein all plants, subsequent pathways can vary substantiallyfrom genus to genus or even in different tissues in thesame plant. A brief summary of demonstrated intercon-versions among gibberellins in pea seed and seedlings ispresented in Figure 19.6 simply as one example. Simi-lar pathways for biosynthesis of gibberellins have beendemonstrated in cell-free endosperm and embryo prepa-rations from pumpkin (Cucurbita maxima) or inferredfrom the knowledge of native gibberellins in Arabidopsis.The 13-hydroxylation pathway (bold arrows) leading toGA20 and GA1 is probably of widespread occurrence inhigher plants.

19.6 Growth Retardants Block the Synthesis of Gibberellins 329

CH3H3C

CH2

CH3

CH2 OPP

(AMO-1618, CCC, Phosphon-D)GGPP

(Phosphon-D)

CH2

CH3H3C

CH3

(Ancymidol)

ent-Kaurene

Copalylpyrophosphate

pyrophosphate (PPi)

CH2

COOHH3C

CH3

Kaurenoic acid

CH2

H3C

CH3

GA12 -7-aldehydeCOOHCHO

CH3

FIGURE 19.5 Gibberellin biosynthesis from geranyl-geranyl pyrophosphate (GGPP) to GA12-7-aldehyde.GA12-7-aldehyde is inactive, but serves as the precur-sor to all other gibberellins. The positions at which someantigibberellins (growth retardants) block gibberellinbiosynthesis are indicated.

19.5 GIBBERELLINS AREDEACTIVATED BY2β-HYDROXYLATION

There are several mechanisms for deactivating gib-berellins and removing them from the active hormonepool. The principal mechanism, however, appears to bethe introduction of a hydroxyl group at the 2 position,which renders the gibberellin inactive. Other inactiva-tion products are possible, depending on the species. In

(2β-OH)

(3β-OH)

(3β-OH)

GA8

(inactive)

∗

(2β-OH)

GA29

(inactive)

∗

CO2

GA20∗ GA1

∗

GA19∗ GA17

∗

GA53

GA12

GA12 - 7 - aldehyde

(13-OH) hydroxylation

(2β-OH)

GA51

(inactive)

∗

CO2

GA9∗

GA24

FIGURE 19.6 Proposed pathway for gibberellin biosyn-thesis in pea (Pisum sativum). The major pathway (boldarrows) occurs in seeds and shoots. The pathway shownin light arrows occurs only in shoots. The asterisk (*)indicates known endogenous forms.

immature, actively developing seed of Phaseolus vulgaris,the principal free gibberellins are the active GA1 andits inactive 2-β-hydroxyl analog GA8, although smallamounts of GA4, GA5, GA6 (all C19 GAs), and GA37 andGA38 (C20 GAs) are also found. Mature seeds, however,contain mainly GA8-glucoside, with smaller amounts ofglucosyl esters of GA1, GA4, GA37, and GA38. In maize(Zea mays) the biologically inactive GA8 is further con-verted to GA8-glucoside, which is also inactive. In pea(Pisum sativum), however, GA8 is oxidized to a 2-ketoderivative referred to as a GA8 catabolite.

19.6 GROWTH RETARDANTSBLOCK THE SYNTHESISOF GIBBERELLINS

Since the 1950s, a number of synthetic compoundsknown as growth retardants have been developed. Thesecompounds, also known as antigibberellins, have foundcommercial use, particularly in the production of orna-mental plants. Growth retardants may be applied topotted plants either as a foliar spray or soil drench. Theprincipal effect of growth retardants is to reduce stemelongation, resulting in plants that are shorter and morecompact, with darker green foliage. Flower size, how-ever, is unaffected. Because individual growth retardantsblock specific steps in gibberellin biosynthesis, they havealso found use as tools in physiological research.

330 Chapter 19 / Hormones II: Gibberellins

AMO-1618 and phosphon are antigibberellins thatinhibit enzymes involved in the synthesis of kaurenewhile ancymidol blocks the subsequent oxidationof kaurene to kaurenoic acid. The effects of theseinhibitors can be reversed by the application ofgibberellins, such as GA20 or GA3. Another inhibitor,known as BX-112, blocks the 3β-hydroxylation ofGA20 to GA1. In those plants where GA1 is the activegibberellin, the effects of BX-112 can be reversed onlyby the application of GA1 itself.

19.7 GIBBERELLIN TRANSPORTIS POORLY UNDERSTOOD

Gibberellin transport studies have been conductedlargely by application of radioactively labeled GAsto either stem or coleoptile sections. Gibberellinshave been detected in both the phloem and xylemsaps. Transport of gibberellins does not appear to bepolar, as it is with auxin, but moves along with otherphloem-translocated organic materials according toa source-sink relationship. Whether gibberellins areactually transported in the xylem is not clear; theycould end up there simply by lateral translocation fromthe phloem. On the other hand, it is likely that anygibberellins synthesized in the root tip are distributed

to the aerial portions of the plant through the xylemstream. It is not known whether gibberellins aretransported as free hormones or in conjugated form.

19.8 GIBBERELLINS AFFECTMANY ASPECTS OF PLANTGROWTH ANDDEVELOPMENT

Since the discovery of gibberellins, it has been clearthat this hormone regulates stem elongation and themobilization of endosperm reserves during the earlystages of seed germination. In addition, largely throughstudies of GA-deficient mutants, gibberellins have beenimplicated in a wide range of other developmentalresponses such as flowering and flower development,root and fruit growth, the development of seeds in thefruit, de-etiolation, and the initiation of leaf primordiain meristems.

19.8.1 GIBBERELLINS STIMULATEHYPER-ELONGATION OFINTACT STEMS, ESPECIALLY INDWARF AND ROSETTE PLANTS

It was excessive stem elongation in infected riceplants that led to the discovery of gibberellins, and

BOX 19.2COMMERCIALAPPLICATIONS OFGIBBERELLINS

The principal commercial use of gibberellins is in theproduction of table grapes, such as the ‘‘ThompsonSeedless.’’ A gibberellin spray in the early stages offlowering thins the cluster by stimulating elongation ofthe floral stems. This spreads the flowers out, allowingfor the development of larger fruit. Larger fruit size isencouraged by a second spray at time of pollination andfruit set. Gibberellins have also been used to enhancegermination and stimulate early seedling emergence andgrowth in species such as grape, citrus, apples, peach, andcherry. Treatment of cucumber plants with gibberellinwill promote formation of male flowers, which is usefulin the production of hybrid seed. The GA-inducedα-amylase in barley aleurone has led to widespread useof GA in the malting industry where it is used to speedup malt production.

The inhibition of gibberellin biosynthesis also hascommercial applications. The growth of many stems

can be reduced or inhibited by synthetic growth retar-dants or antigibberellins. These include AMO-1618,cycocel (or, CCC), Phosphon-D, ancymidol (knowncommercially as A-REST), and alar (or, B-nine). Growthretardants mimic the dwarfing genes by blocking spe-cific steps in gibberellin biosynthesis, thus reducingendogenous gibberellin levels and suppressing internodeelongation. These compounds have found significantcommercial use, particularly in the production of orna-mental plants. Growth retardants may be applied topotted plants either as a foliar spray or soil drench.Their principal effect is to reduce stem elongation,resulting in plants that are shorter and more com-pact, with darker green foliage. Flower size, however,is unaffected. Commercial flower growers have foundthese inhibitors useful in producing shorter, more com-pact poinsettias, lilies, and chrysanthemums, and otherhorticultural species.

In some areas of the world, wheat tends to ‘‘lodge’’near harvest time, that is, the plants become top-heavywith grain and fall over. Spraying the plants with anti-gibberellins produces a shorter, stiffer stem and thusprevents lodging. Antigiberellins also have been used toreduce the need for pruning of vegetation under powerlines.

19.8 Gibberellins Affect Many Aspects of Plant Growth and Development 331

hyper-elongation of stem tissue remains one ofthe more dramatic effects of gibberellins on higherplants. Unlike auxins, gibberellins promote elongationalmost exclusively in intact plants rather than excisedtissues. Nowhere is this more evident than in thecontrol of internode elongation in genetic dwarfs. Therelationship between dwarfing or internode-lengthgenes and gibberellins was pioneered by the work ofB. O. Phinney on maize (Zea mays) and P. W. Brian andcoworkers on garden pea (Pisum sativum). Since thesepioneering studies, experiments have been conductedwith dwarf mutants of rice (Oryza sativa), bean (Phaseolusvulgaris), Arabidopsis thaliana, and several others. In allcases, application of exogenous gibberellin to the dwarfmutant restores a normal, tall phenotype (Figure 19.7).Exogenous gibberellin has no appreciable effect on thegenetically normal plant.

In maize, more than 30 mutants that influenceplant height have been described. Maize plants express-ing these mutations have shortened internodes, dueto reduced cell division and cell elongation, and atmaturity reach only 20 to 25 percent of the height ofnormal plants. At least five of these mutants (d1, d2,d3, d5, an1) exhibit the normal phenotype when treatedwith GA3, but show no response to other hormonesor growth regulators. Activity assays such as these havebeen supported by other biochemical and radiotracerexperiments, demonstrating conclusively that internodeelongation in maize is under control of gibberellins.Specifically, each mutation blocks a different step in the

FIGURE 19.7 The effect of gibberellic acid on dwarfpea seedlings. Left: Control, showing reduced intern-ode elongation characteristic of the dwarf growthhabit. Right: Gibberellin treated with a 5 × 10−4 Mfoliar-drench of GA3. Note that gibberellin treatmentincreased stem elongation by simulating elongation ofthe internodes.

biosynthetic pathway toward GA1, which is the activeform of gibberellin in maize. Similar experiments withthe Le allele in garden pea (the same allele studied byGregor Mendel in his pioneering genetic studies) havedemonstrated that this dwarf genotype (the homozygousrecessive le/le) also blocks the synthesis of GA1. In bothmaize and pea, it has been shown that the dwarf genotypeleads to a significant reduction in the gibberellin levels.While studies with dwarf plants have been instrumen-tal in linking gibberellins with stem elongation, thereare other dwarf mutants known that do not respondto application of gibberellin. These mutants may beunrelated to gibberellin-controlled growth and subjectto other, as yet unknown, regulating factors.

Additional support for the role of gibberellins instem elongation comes from the study of rosette plants.A rosette is essentially an extreme case of dwarfism inwhich the absence of any significant internode elon-gation results in a compact growth habit characterizedby closely spaced leaves. The failure of internode toelongate may result from a genetic mutation, or maybe environmentally induced. Regardless of the cause,hyper-elongation of stems in rosette plants is invariablybrought about by the application of small amounts ofgibberellin (Figure 19.8).

Environmentally limited rosette plants such asspinach (Spinacea oleraceae) and cabbage (Brassica sp.)generally do not flower in the rosette form. Just beforeflowering, these plants will undergo extensive internodeelongation, a phenomenon known as bolting. Boltingis normally triggered by an environmental signal,either photoperiod (as in spinach) or a combinationof low temperature and photoperiod (as in cabbage).We will return to the phenomena of photoperiod andcold requirement in later chapters. It is sufficient tonote here that, under conditions normally conduciveto the rosette habit, spinach, cabbage, and many other

FIGURE 19.8 Gibberellin-stimulated stem growth in arosette genotype of Brassica napus. Treatments were(from left): 0, 0.5, 1.0, 10.0 ng GA3 per plant, applied tothe meristem.

332 Chapter 19 / Hormones II: Gibberellins

rosette plants can be induced to bolt by an exogenousapplication of gibberellic acid.

The above results suggest that (1) gibberellins are alimiting factor in the stem growth of rosette plants and(2) the effect of long days or cold treatment is to removethat limitation. These possibilities have been confirmedin spinach and Silene armeria, both photoperiodic plantsrequiring long days to flower, by the extensive inves-tigations of J. A. D. Zeevaart and coworkers. Spinachcontains six gibberellins, including GA19 and GA20.GA20 will cause bolting in spinach under short day con-ditions while GA19 is biologically inactive. Zeevaart andcoworkers found that rosette plants of spinach containhigh levels of the inactive form GA19 and low levels ofthe active GA20. Upon transfer to long day conditions,however, the level of GA19 declined while the level ofGA20 increased (Figure 19.9). The reciprocal changesin GA19 and GA20 levels suggests a precursor-productrelationship, which was confirmed in whole plants byfeeding deuterium (2H)-labeled precursors. In otherexperiments, 14C-labeled GA19 was converted to GA20by cell-free extracts from spinach plants maintainedunder long days, but not in extracts from plants main-tained under short days. On the basis of these studies,it may be concluded that gibberellins have a significantrole in the control of stem elongation in rosette plants.

The relationship between gibberellins and stemelongation in cold-requiring plants has not been studiedas thoroughly as it has for plants that are sensitive to pho-toperiod. As mentioned above, exogenous application ofGA3 will substitute for the cold requirement in manyplants and there is some evidence, based on bioassays,

Number of long days

Rel

ativ

e G

A c

onte

nt

100

80

60

40

20

0 2 4 6 8 10 12 14

Ste

m le

ngth

(m

m)

50

40

30

20

10

Stemlength

GA19

GA20

FIGURE 19.9 Spinach plants (Spinacea oleraceae) exhibitextensive stem elongation when transferred from shortdays to long days. Stem elongation is accompanied bychanges in gibberellin content as an inactive form is con-verted to an active form. A decrease in the level of theinactive GA19 is matched by a corresponding increasein the active form G20. (Redrawn from Metzger andZeevaart 1980. Plant Physiology 66:844–846. CopyrightAmerican Society of Plant Physiologists.)

that gibberellin-like activity increases in plants followingcold treatment. It is reasonable to expect on the basis ofthese studies that changes in gibberellin biosynthesis ormetabolism are involved, but a more thorough study isrequired.

19.8.2 GIBBERELLINS STIMULATEMOBILIZATION OF NUTRIENTRESERVES DURINGGERMINATION OF CEREALGRAINS

Gibberellins initiate the mobilization of nutrientreserves stored in the endosperm, while auxins promoteelongation of the embryonic axis. The auxins thatsupport early embryo growth are largely derived fromthe breakdown of stored conjugates to free, activeIAA while the gibberellins, at least in cereal grains,appear to be released by the hydrated embryo from apreformed GA pool. In Arabidopsis, on the other hand,seeds carrying mutations such as ga1, ga2, and ga3, thatact early in gibberellin biosynthesis, fail to germinatebut germination can be rescued by applying exogenousgibberellin. More recently, it has been suggested thatbrassinosteroids might also have a role in germinationof Arabidopsis seed (Chapter 21).

A role for gibberellins in mobilization of reservesduring seed germination was first suggested by exper-iments on germinating cereal grains in the late 1950s.Cereal grains such as rye, barley, and wheat have aprotein-rich layer of cells called the aleurone whichsurrounds the starchy endosperm tissue. During germi-nation, cells in the aluerone secrete a range of hydrolyticenzymes, including α-amylase and proteases, which areinvolved in the hydrolysis of carbohydrate and proteinstored in the endosperm.

The involvement of gibberellins in enzyme secre-tion can be shown by a relatively simple experiment.Seeds of cereals such as barley are transected to producetwo half-seeds (Figure 19.10). One half-seed containsthe embryo and the other half-seed does not. Whenimbibed, the embryo-containing half-seed will proceedto secrete α-amylase and other hydrolytic enzymesin order to digest the starchy endosperm, mobilizethe resulting nutrients, and initiate germination. Thehalf-seed without the embryo cannot, of course, ger-minate but neither does it produce elevated levels ofα-amylase or any of the other hydrolytic enzymesrequired for germination. Treatment of the embryo-lesshalf-seed with gibberellic acid, however, will stimu-late the half-seed to produce high levels of α-amylase.Experiments of this general nature have shown that thegerminating embryo sends a signal, probably gibberellin,to the aleurone cells. There the gibberellin either acti-vates or derepresses transcription of genes encoding thenecessary hydrolytic enzymes. These enzymes are then

19.9 Gibberellins Act by Regulating Gene Expression 333

FIGURE 19.10 Gibberellin-stimulated secretion of α-amylase from barley half-seeds.Embryo-less half-seeds were incubated on the surface of a starch-agar gel. After48 hours, the gel was washed with iodine-potassium iodide (IKI), a reagent thatreacts with starch to form a blue-black color. A clear circle, or halo, surround-ing the half-seed indicates the digestion of starch by α-amylase. The control plate(left) contains four half-seeds but no added gibberellin. Two half-seeds producedno α-amylase while the other two exhibited low activity. The plate on the right con-tained 10 nanomoles gibberellic acid. Each of the gibberellin-treated half-seeds issurrounded by a large halo, indicating active α-amylase secretion. (From a studentexperiment.)

released into the endosperm where they break downthe starches and proteins to provide nutrients for thegrowing embryo (Figure 19.11).

It should be noted that much of this work hasbeen conducted on a single cultivar (cv. ‘‘Himalaya’’)of barley and the responsiveness of seeds to gibberellincan be significantly affected by environment during seeddevelopment and maturation. Seed from barley grownat high temperatures, for example, produce high levelsof α-amylase in the absence of added GA. Nonetheless,the principles that have emerged from the barley systemappear to be widely applicable to wheat, rye, oats, andother cereal grains.

19.9 GIBBERELLINS ACT BYREGULATING GENEEXPRESSION

Early physiological studies found that GA-stimulatedα-amylase secretion could be blocked by inhibitors suchas actinomycin D and cycloheximide, which inhibit RNAand protein synthesis, respectively. Results like this indi-cated that gibberellin stimulated de novo synthesis ofα-amylase by the aleurone layer.

Does gibberellin regulate transcription of α-amylase mRNA? An unequivocal answer to thisquestion was possible only after techniques for isolationof protoplasts from barley aleurone cells were perfected.Aleurone protoplasts provide a useful experimentalsystem. They respond normally to GA by exhibiting thesame ultrastructural changes and producing the sameisozymes with the same efficiency as intact aleuronecells. Most importantly, high yields of transcriptionallyactive nuclei can be readily isolated from protoplasts.

Based on evidence from several lines of investiga-tion, it is clear that gibberellin dose regulates transcrip-tion of α-amylase mRNA. Both in vivo pulse labelingof protein and in vitro translation of protein from totalaleurone RNA, followed by electrophoretic and autora-diographic analysis, show significant increases in theamount of α-amylase translated following the appli-cation of gibberellin (Figure 19.12). α-Amylase mayconstitute as much as 50 to 60 percent of the totaltranslated aleurone protein.

The gibberellin response is apparently not limitedto α-amylase alone, as a small number of other pep-tides either increase or decrease following gibberellintreatment. Furthermore, Northern blot hybridizationof α-amylase cDNA clones with RNA from aleuronelayers has confirmed that gibberellin causes an increase

334 Chapter 19 / Hormones II: Gibberellins

α-amylase

23

2

1

Aleurone

Endosperm

β-amylase(inactive)

β-amylase(active)

Starch

Glucose

Protease

GA

Scutellum

Coleoptile

Plumule(with embryonic leaves)

Radicle(embryonic root)

Coleorhiza

EM

BR

YO

FIGURE 19.11 A schematic illustratinggibberellin-induced release of enzymes and carbo-hydrate mobilization during germination of barley(Hordeum vulgare) seed. Gibberellin moves from theembryo (1) to the aleurone where it stimulates thesynthesis of α-amylase and protease enzymes. (2) Theprotease converts an inactive β-amylase to the activeform. α- and β-amylase together digest starch to glucose,(3) which is mobilized to meet the metabolic demands ofthe growing embryo.

in the physical abundance of α-amylase mRNA. Fol-lowing gibberellin treatment, α-amylase mRNA maycomprise as much as 20 percent of the total translatablemRNA. Finally, time course studies have shown thatthe rate of α-amylase synthesis following gibberellintreatment closely correlates with the rate of mRNAaccumulation. These studies pretty well established thata primary action of gibberellin, at least with respectto seed germination, is to regulate in some way genetranscription.

The challenge over the past ten years or so hasbeen to identify the genes and proteins that are involvedin the gibberellin signaling pathway. This has beenaccomplished primarily by screening for mutations thateither enhance or lower gibberellin sensitivity. One suchmutant is the rice mutant slender rice1 (slr1). The slr1mutation results in what is called a constitutive gibberellin

phenotype, which means that it behaves as if it weresaturated with gibberellin. Mutant seedlings elongaterapidly and excessively compared with the wildtype andα-amylase is produced by embryo-less half-seeds with-out added gibberellin. Excessive gibberellin productioncan be ruled out as the cause because the endogenousGA content of the mutant is lower than that of the wild-type and the response is not sensitive to inhibitors thatblock gibberellin synthesis. Similar ‘‘slender’’ mutantshave been described in barley (Hordeum vulgare) and pea(Pisum sativum).

Another example is the gibberellic acid insensitive(gai) mutant in Arabidopsis. gai mutants are dwarfs thatdo not respond to applied gibberellin. Similar mutations

68,000

43,000

25,000

12,400

SECRETEDPROTEINSAT 12 hr

�-AMYLASE

ABA GA C A+G

FIGURE 19.12 Hormonal control of α-amylase biosynthe-sis by barley aleurone layers. Polypeptides synthesized byisolated aleurone layers were labeled with the radioactiveamino acid [35S]methionine incubated in the presence ofABA, GA3, or ABA + GA3 (A + G). Polypeptides secretedinto the incubation medium were separated by elec-trophoresis on polyacrylamide gels. Polypeptides incor-porating the radioactive label were detected by exposingthe gel to X-ray film. Note the pronounced stimulationof α-amylase biosynthesis in the presence of GA3, a stim-ulation almost completely abolished in the presence ofABA. C represents untreated controls. Numbers indicateapproximate molecular mass. (From Higgins et al. 1982.Plant Molecular Biology 1:191. Reprinted by permissionof Kluwer Academic Publishers. Original photographkindly provided by Dr. J. V. Jacobsen.)

19.9 Gibberellins Act by Regulating Gene Expression 335

BOX 19.3DELLA PROTEINSAND THE GREENREVOLUTION

Throughout the 1950s and 1960s, the world witnessedadvances in plant breeding and agronomy that led to sig-nificant increases in agricultural production worldwide.These transformations, collectively referred to as the‘‘green revolution,’’ were supported in part by the Rock-efeller Foundation, other philanthropic organizations,and universities in an effort to help food productionkeep pace with worldwide population growth.

Some of the most significant advances were seen inwheat and maize breeding. Wheat yields, for example,were substantially increased by breeding new varieties

that were shorter and thus more resistant to lodgingdamage by wind and rain. The new, shorter varietiesalso increased grain production at the expense of strawbiomass. The significance of these efforts was recognizedwhen Norman Borlaug of the International Maize andWheat Improvement Center (CIMMYT) in Mexicowas awarded the 1970 Nobel Peace Prize for his workin wheat breeding.

The new varieties were shorter because the breedershad introduced mutant dwarfing alleles, Reduced height-1or Rht-B1in wheat and dwarf-8 or d8 in maize, thatconferred a reduced response to gibberellins. In 1999,J. Peng et al. (Nature 400:256) showed that the Rhtand d8 loci encode Della transcription factors that areanalogous to the gibberellin insensitive (GAI) protein inArabidopsis. Like the mutant gai protein in Arabidopsis,the rht-b1 and d8 proteins have an altered amino acidsequence at the N-terminal end that interferes withgibberellin signaling.

at the same genetic locus are responsible for dwarfingin wheat (Triticum aestivum) and maize (Zea mays). slr1and gai mutant plants respond the way they do becausethe wild-type SLR1 and GAI genes encode transcriptionregulators referred to as DELLA proteins. DELLAproteins are a class of nuclear proteins that appear tofunction as repressors in gibberellin signaling. This con-clusion is based on earlier observations that degradationof DELLA proteins in planta triggers gibberellin-typeresponses. Although SLR1 is the only DELLA genedescribed for rice, a total of five, including GAI, havebeen described for Arabidopsis.

The discovery of DELLA proteins has been fol-lowed by the discovery of a soluble GA receptor protein,GA INSENSITIVE DWARF 1 (GID1), in rice. GID1encodes a nuclear protein that binds with gibberel-lic acid both in vitro and in vivo. Subsequently itwas found that Arabidopsis carries three homologousgenes, designated AtGID1a, AtGID1b, and AtGID1c,which encode similar proteins. Each of the Arabidop-sis genes apparently has a specific, but overlapping,role in growth and development. For example, individ-ually the mutant genes atgid1a, atgid1b, and atgid1cshow no clear phenotype and even when any twomutant genes were present, germination was normal.The double mutant atgid1a-atgid1c exhibited a dwarfphenotype while other double mutant combinationswere of normal height. However, the triple mutantatgid1a-atgid1b-atgid1c failed to germinate without peel-ing off the seed coat and the seedlings were severedwarfs, achieving a height of only a few millime-ters after a month’s growth. On the other hand, the

expression of any one of the Arabidopsis wildtype genesin gid1 mutant rice plants produced normal rice plantswith normal GA sensitivity.

All of this has been brought together recently bythe demonstration that the GID1 receptor binds notonly with gibberellin, but also with the DELLA proteinSLR1 to form a GA-GID1-SLR1 complex. Accordingto the current model, this hormone-receptor-repressorcomplex then results in the degradation of the DELLArepressor protein by the 26S proteasome pathway,thus relieving a DELLA-imposed repression and allow-ing expression of the gibberellin-responsive gene. Inother words, the function of gibberellin—to derepressgenes by facilitating the removal and ubiquitin-mediateddegradation of a repressor protein— is similar to that ofauxin described earlier in this chapter. This model wouldalso explain why the gai mutants are dwarfs that do notrespond to GA. Cloning of the Arabidopsis GAI gene hasshown that the mutant gai protein is missing a group ofamino acids at its N-terminal end. These missing aminoacids presumably render the protein insensitive to thegibberellin-receptor complex without interfering withits role as a repressor (Figure 19.13).

GA elicits multiple responses in plants and it may yetbe demonstrated that some of these responses involvemembrane-bound receptors and a cascade of secondarymessengers. However, it is interesting to note that, in1957, P. W. Brian suggested that plants contained anendogenous inhibitor and gibberellin acted not by directstimulation of a response but by relieving this inhibition.Fifty years later, we have come full circle and Brian’sproposal now seems remarkably prescient.

336 Chapter 19 / Hormones II: Gibberellins

A. Wild-type minus GID

B. Wild-type plus GID

C. Mutant

Repression

Repression

GA+

GID

DerepressionGrowth

Growth

GA

GAI

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GID

gai

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FIGURE 19.13 A model for the roles of GA receptors and DELLA proteins in the gib-berellin signaling pathway. (A) GAI represses growth-dependent gene expression.GA alone cannot bind with GAI to reverse the repression. (B) The GA/GID-receptorcomplex is able to bind with GAI, targeting the repressor for degradation and dere-pressing the gene. (C) The mutant DELLA protein gai is unable to bind with theGA/GID-receptor complex, but continues to repress gene expression. (After Har-berd, N. P. 1998. BioEssays 20:1001–1008.)

SUMMARY

Gibberellins are diterpenes, noted for their capacity tostimulate hyper-elongation of intact stems, especiallyin dwarf and rosette plants and stimulate mobilizationof endosperm reserves during seed germination.

Gibberellins are related biosynthetically tocarotenes and other isoprene derivatives. Principalintermediates are geranyl-geranyl-pyrophosphate andGA12-7-aldehyde.Gibberellins can be inactivated byhydroxylation at the C-2 position or by conversionto inactive conjugates. Gibberellins appear to betransported primarily in the phloem in responseto source-sink relationships. Chemicals that block

gibberellin biosynthesis are used commercially toproduce shorter, more compact plants.

Gibberellin appears to act primarily by target-ing the repressor proteins for degradation by the 26Sproteasome pathway.

CHAPTER REVIEW

1. Describe how the size of the active hor-mone pool is regulated for gibberellins.

2. What is the significance of the ‘‘G’’ numberassigned to each gibberellin?

Further Reading 337

3. Identify structural characteristics that determinewhether a gibberellin is active or inactive.

4. Describe hormonal involvement in nutrientmobilization in germinating cereal grains.

5. Compare and contrast the auxin and gib-berellin signal chains for gene derepression.

FURTHER READING

Brian, P. W. 1957. The effects of some microbial metabolicproducts on plant growth. Symposium of the Society forExperimental Biology 11:168–182.

Buchanan, B. B., W. Gruissem, R. L. Jones. 2000.Biochemistry and Molecular Biology of Plants. Rockville,MD: American Society of Plant Physiologists.

Davies, P. J. 2004. Plant Hormones: Biosynthesis, Signal Trans-duction, Action. Dordrecht: Kluwer Academic Publishers.

Eckardt, N. A. 2007. GA perception and signal transduction:Molecular interactions of the GA receptor GID1 with

GA and the DELLA protein SLR1 in Rice. The PlantCell. 19:2095–2097.

Harberd, N. P. et. al. 1998. Gibberellin: Inhibitor of aninhibitor of . . . ? BioEssays 20:1001–1008.

Olszewski, N. T-p. Sun, F. Gubler 2002. Gibberellin signal-ing: Biosynthesis, catabolism, and response pathways.The Plant Cell. Supplement S61–S80.

Razem, F. A., K. Baron, R. D. Hill. 2006. Turning on gib-berellin and abscisic acid signaling. Current Opinionin Plant Biology 9:454–459.

Schechheimer, C. 2008. Understanding gibberellic acidsignaling—are we there yet? Current Opinion in PlantBiology 11:9–15.

Swain, S. M., D. P. Singh. 2005. Tall tales from sly dwarfs:Novel functions of gibberellins in plant development.Trends in Plant Science 10:123–129.

Weiss, D., N. Ori. 2007. Mechanisms of cross talk betweengibberellins and other hormones. Plant Physiology144:1240–1246.

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20Hormones III: Cytokinins

Cytokinins (CK) are derivatives of the nitrogenous baseadenine and are noted primarily for their capacity tostimulate cell division in tissue culture. Cytokinins alsoinfluence a number of other developmental responses,including shoot and root differentiation in tissue culture,the growth of lateral buds and leaf expansion, chloroplastdevelopment, and delay of senescence. Most recently ithas been learned that cytokinins play a fundamentalrole in maintaining the indeterminate property of shootapical meristems.

This chapter includes

• the structure, synthesis, and metabolism of cyto-kinins,

• the role of cytokinins in regulating cell division,• the control of shoot meristem function by cyto-

kinins,• the role of cytokinins in apical dominance, senes-

cence, and other developmental events, and• cytokinin receptors and signal transduction.

20.1 CYTOKININS ARE ADENINEDERIVATIVES

Naturally occurring cytokinins are all adenine deriva-tives with either an isoprene-related side chain or an

aromatic (cyclic) side chain. The former are called iso-prenoid cytokinins and the latter are called aromaticcytokinins. Although there is some variation dependingon species and developmental stage, the most commonisoprenoid cytokinins are N6-(�2-isopentenyl)- ade-nine (iP), trans-zeatin (tZ), and dihydrozeatin (DZ)(Figure 20.1). The aromatic cytokinins, such as benzyl-adenine (BA) are less common and are found in only afew species.

The original cytokinin, kinetin, is a synthetic de-rivative that has not yet been identified in plants(Box 20.1). Curiously, however, there is a recent reportthat kinetin has been identified in human urine.

20.1.1 CYTOKININ BIOSYNTHESISBEGINS WITH THECONDENSATION OF ANISOPENTENYL GROUP WITHTHE AMINO GROUP OFADENOSINE MONOPHOSPHATE

Enzymes that direct the synthesis of cytokinins havebeen isolated from the slime mold Dictyostelium dis-coideum, tobacco callus tissue, and crown gall tissue (seeFigure 20.7). The key reaction is the addition of adimethylallyl diphosphate (DMAP) group to the nitro-gen at the 6-position of adenosine−5′-monophosphate

339

340 Chapter 20 / Hormones III: Cytokinins

Kinetin (N6-furfuryl adenine)

CH2HN

NHN

N N O6

iP (N6-(Δ2-isopentenyl) adenine)

CH2HN

NHN

N N6

CH3

CH3

CH2 N

NHN

N N6

CH2OH

CH3

BAP (N6-(benzyl) adenine)

CH2HN

NHN

N N6

Z (trans-Zeatin)

FIGURE 20.1 The chemical structures of four representa-tive cytokinins. Kinetin, the first compound found withcytokinin activity, is a synthetic cytokinin prepared byheating DNA. Isopentenyl adenine (iP) and trans-Zeatin,both isoprenoid-type cytokinins, are the most commonnaturally occurring cytokinins. Benzyladenine (BAP) isan aromatic cytokinin. The N6 position of adenine isindicated and the side chains are highlighted.

(AMP) (Figure 20.2). DMAP, introduced previously inChapter 19, is also known as �2-isopentenyl diphos-phate or �2iPP. This addition reaction is catalyzed bythe enzyme adenosine phosphate-isopentenyl trans-ferase (IPT). The product is N6-(�2-isopentenyl)-adenosine−5′-monophosphate or iPRMP. The IPT-catalyzed reaction is also the rate limiting reaction incytokinin biosynthesis, a factor that has enabled manyinvestigators to manipulate the cytokinin content of

tissues by transforming plants with genes that cause anoverexpression of IPT.

In the next two steps, the phosphate group andthe ribose group removed from [9R−5′P]iP to formthe active cytokinin N6-(�2-isopentenyl)-adenine (iP).Alternatively, the isopentenyl side chain of [9R−5′P]iPmay be hydroxylated before the phosphate and ribosegroups are removed to form zeatin (Z). Zeatin and iPare thought to be the most biologically active cytokininsin most plants. Reduction of the double bond in the sidechain of zeatin would give the dihydrozeatin derivative,which is particularly active in some species of legumes.

Cytokinins are known to undergo extensive inter-conversions between the free base (or, nucleobase),ribosides, and ribotides when experimentally suppliedto tissues. Enzymes have been identified in wheat germthat catalyze the conversion of iP to its riboside ([9R]iP)or to its ribotide ([9R−5′P]iP) as well as enzymes thatcatalyze the hydrolysis of the ribotides and ribosides tothe free base (iP). Naturally, these rapid interconversionsmake it very difficult to ascertain which form is the truly‘‘active’’ form of the hormone. However, this confusionhas been at least partially resolved with the identificationof cytokinin receptors, discussed later in this chapter.Individual receptors from different species appear tohave different affinities for cytokinin nucleobases, ribo-sides and ribotides. This may be simply one way ofconferring specificity through the cytokinin-receptorinteraction.

20.1.2 CYTOKININS MAY BEDEACTIVATED BYCONJUGATION OR OXIDATION

There are two principal routes for regulating cytokininactivity levels by removal of cytokinins from the activepool: (1) conjugation with either glucose or aminoacids, and (2) oxidation. Examples of glucosylation ofthe side chain hydroxyl group (O-glucosylation) ofzeatin or dihydrozeatin, for example, are abundant inplants (Figure 20.3). O-glucosides are not themselvesactive, but are readily hydrolyzed to active cytokininby glucosidase enzymes. O-glucosylation appears to bea mechanism for storing excess cytokinin for retrievalwhen physiological conditions warrant. O-glucosidesare also resistant to oxidation and thus may serve toprotect the hormone from oxidation while being trans-ported to a target tissue. In some plants, such as radish,maize, or tobacco, cytokinins may also be glucosylatedat one of the nitrogen positions on the purine ring.Both the 7- and 9-glucosides are biologically inactive.N-glucosides are also very stable and do not appear to behydrolyzed readily to give the active free base. Their for-mation thus appears to be more a means for permanentinactivation of cytokinins rather than storage.

20.2 Cytokinins are Synthesized Primarily in the Root and Translocated in the Xylem 341

CH2HN

NHN

N N O6

BOX 20.1THE DISCOVERYOF CYTOKININS

The discovery of cytokinins came about because plantcells in culture would not divide. The first experimentalevidence for chemical control of plant cell division wasprovided by Haberlandt in 1913, when he demonstratedthat phloem sap could cause nondividing, parenchyma-tous potato tuber tissue to revert to an actively dividingmeristematic state. Other cell-division factors were laterdemonstrated in wounded bean pod tissue, extracts ofDatura ovules, and the liquid (milky) endosperm ofcoconut.

In the 1940s and 1950s, plant tissue culture wasattracting the attention of physiologists as a tool forstudy of cell division and development. One group,under the direction of F. Skoog at the University ofWisconsin, was studying the nutritional requirementsof tissue cultures derived from tobacco stem segments.Skoog and coworkers found that stem tissue explantscontaining vascular tissue would proliferate on a de-fined medium containing auxin. On the sameauxin-containing medium, however, tissue explantsfreed of vascular tissue would exhibit cell enlargement,but failed to divide. They soon found that extractsof vascular tissue, coconut milk, and yeast would allstimulate cell division in the presence of auxin.

C. O. Miller, then working as a postdoctoral studentin Skoog’s laboratory, took on the task of isolating theactive material. Miller was able to provisionally identifythe active material as a adenine, one of the nitrogenous

bases found in nucleic acids. This led to a search foractive material in nucleic acid preparations, a source highin adenine. In a beautiful piece of serendipity, Millersampled a bottle of herring sperm DNA which had beensitting on the laboratory shelf. The sample proved to behighly active, so a fresh supply of herring sperm DNAwas ordered. Unfortunately, the fresh sample proved tobe completely inactive. It turns out that the active princi-ple in the original DNA sample had slowly accumulatedas the DNA aged on the laboratory shelf. Activity couldbe generated in fresh DNA samples simply by artificially‘‘aging’’ the sample with heat and acid.

In 1956, Miller and his colleagues reported theisolation and crystallization of a highly active substance,identified as the adenine derivative N6-furfurylamino-purine, from autoclaved herring sperm DNA. Becausethe compound elicited cell division, or cytokinesis, intissue culture, Miller and his colleagues named thesubstance kinetin. In 1965, Skoog and his colleaguesproposed the term cytokinin.

Even though kinetin remains one of the mostbiologically active cytokinins, it is an artifact of iso-lation from DNA and has not been found in plants.However, the discovery of kinetin and its dramaticeffect on cell division stimulated physiologists tolook for naturally occurring cytokinins. In the early1960s, Miller, then at Indiana University, and D.S. Letham, working in Australia, independentlyreported the isolation of a purine with kinetinlikeproperties from young, developing maize seed andplum fruitlets. This substance was characterized as6-(4-hydroxy-3-methyl-trans-2-butenylamino) purine,which was given the trivial name zeatin. Since the dis-covery of zeatin, a number of other naturally occurringcytokinins have been isolated and characterized.

Cytokinins also form conjugates with the aminoacid alanine (Figure 20.3). 9-Alanyl conjugates of zeatinand dihydrozeatin have been identified in tissues oflupine (Lupinus) fruit and root nodules, immature appleseeds, and bean (Phaseolus) seedlings. These are also verystable conjugates that probably serve to inactivate thecytokinin in the same manner as N-glucosides.

Another mechanism—possibly the major mecha-nism—for removing cytokinins from the hormonepool is their irreversible degradation by the enzymecytokinin oxidase/dehydrogenase (CKX). CKXcleaves the isopentenyl side chain from either zeatinor iP or their ribosyl derivatives. In order to function,however, CKX must recognize the double bond of theisoprenoid side chain. Consequently, dihydrozeatin andaromatic cytokinins are resistant to degradation byCKX. In the same way that the endogenous cytokinin

content can be increased by overexpression of thegene Isopentenyl transferase (IPT) in transgenicplants, cytokinin content can be reduced in trans-genic plants that overexpress cytokinin oxidase/dehydrogenase.

20.2 CYTOKININS ARESYNTHESIZED PRIMARILYIN THE ROOT ANDTRANSLOCATEDIN THE XYLEM

A major site of cytokinin biosynthesis in higher plants isthe root. High cytokinin levels have been found in roots,especially the mitotically active root tip, and in the xylem

342 Chapter 20 / Hormones III: Cytokinins

Adenosine Monophosphate (AMP) Δ2-isopentenyl pyrophosphate

N6(Δ2-isopentenyl)-adenosine-5'-phosphate [9R-5'P]iP

N6(Δ2-isopentenyl) adenine(iP)

CH3

CH3

NH2

OH2C

NN

N N

O

OH OH

POH2C

Zeatin

HN

NHN

N N

CH2OH

CH3CH2

P P+

HN

OH2C

NN

N

O

OH

1 2

OH

P

P

P1 P

CH3

CH3CH2

N

HN

NHN

CH3

CH3CH2

NN

ribose

P

ribose

—

O2

FIGURE 20.2 A general outline for the biosynthesis of isopentenyl adenine (iP) andtrans-zeatin (tZ) from adenosine monophosphate and isopentenyl pyrophosphate.Adenosine diphosphate and adenosine triphosphate may also be used as precur-sors. Reaction 1 is the rate limiting step and is catalyzed by the enzyme adeninephosphate-isopentenyl transferase (IPT).

20.3 Cytokinins are Required for Cell Proliferation 343

N

N

N

NH

HN

A. 0-�-Glucosylzeatin (OG)Z

OO

HO

OHOH

OH

6

3

57

89

421

N

N

N

NH

HN

B. 7-�-Glucosylzeatin [7G]Z

O

OHHO

OH

OH

CH2OH

N

N

N

N

HN

C. 9-Alanylzeatin [9Ala]Z

CH2OH

CH2

CH COOHH2N

FIGURE 20.3 Examples of cytokinin conjugates. Con-jugates of zeatin are shown. (A) O-glucosylation:the side chain hydroxyl group is glucosylated.(B) N-glucosylation: the adenine ring is glucosylatedat the 3-, 7-, or 9-nitrogen. (C) Alanyl conjugates: the9-nitrogen is conjugated with the amino acid alanine.

sap of roots from a variety of sources. It is generallyconcluded that roots are a principal source of cytokininsin most, if not all, plants and that they are transportedto the aerial portion of the plant through the xylem.Indirect support for this conclusion is provided by theobservation that excised leaves from many species canbe maintained in a moist sand bed only if adventitiousroots are permitted to form at the base of the petiole.If these roots do not form or are removed as they form,the leaves will quickly senesce. The delayed senescence

when roots are allowed to form is apparently due to thepresence of cytokinins, which are synthesized in the rootand transported to the leaf through the vascular tissue.

Immature seeds and developing fruits also containhigh levels of cytokinins; the first naturally occurringcytokinins were isolated from milky endosperm of maizeand developing plum fruits. While there is some evi-dence that seeds and fruits are capable of synthesizingcytokinins, there is also evidence to the contrary. Thus itremains equally possible that developing seeds, becauseof their high metabolic activity and rapid growth, maysimply function as a sink for cytokinins transported fromthe roots. On the other hand, there is now evidence thatcytokinins are not always a long-distance messenger. Aswe will see later, meristematic cells in the shoot apicalmeristem and floral meristems in particular are underthe control of locally produced cytokinins.

20.3 CYTOKININS AREREQUIRED FOR CELLPROLIFERATION

The role of cytokinins in regulating cell division firstbecame apparent as a result of attempts to cultureisolated carrot and tobacco tissues on defined media(see Box 20.1). Cell proliferation occurred only whenauxin plus some ‘‘cell-division factor’’ was present in themedium. The ‘‘cell division factor’’ turned out, of course,to be kinetin. It was soon learned that kinetin and othercytokinins, always in the presence of auxin, stimulated celldivision in a wide variety of tissues. The significance ofthese studies was that no cambium or other meristematictissue was present in the cultures, therefore demonstrat-ing that cytokinins have the capacity to initiate divisionin quiescent, or nondividing, cells.

20.3.1 CYTOKININS REGULATEPROGRESSION THROUGHTHE CELL CYCLE

Although much remains to be learned about howcytokinins regulate cell division, studies of tobaccosuspension-cultured cells and Arabidopsis indicate adirect role for cytokinins in regulating progressionthrough the cell cycle.

Freshly established tobacco cell cultures requireboth auxin and cytokinin for continued cell division.The absence of either hormone causes the cells to bearrested in either the G1 or G2 phase of the cell cycle.Following addition of the missing hormone, the onsetof cell division can be detected within 12 to 24 hours. In1996, K. Zhang and coworkers reported that culturedcells arrested in G2 by the absence of cytokinin con-tained cyclin-dependent kinases (CDK) with reducedactivity, due to a high level of phosphorylation on a

344 Chapter 20 / Hormones III: Cytokinins

tyrosine residue. When such cultures were re-suppliedwith cytokinin, the tyrosine was dephosphorylated, theenzyme reactivated, and cell division resumed.

The current evidence suggests that a specific tyro-sine residue in the CDK catalytic unit is phosphorylatedduring the S phase by another kinase (Figure 20.4).Although the phosphorylated CDK catalytic unit is ableto combine with cyclin (in this case, a B-type cyclinor CycB), the phosphorylated complex remains inactiveuntil the inhibitory phosphate group is removed by acytokinin-dependent phosphatase. Thus the principalrole of cytokinin in cultured tobacco cells appears to bethat of generating an active CDK complex that initiatescell division by catalyzing the transition from the G2phase to mitosis, or M phase.

A second type of cytokinin action has been describedfor Arabidopsis. A mutant of Arabidopsis with a highlevel of cytokinin was found to contain as well a highlevel of a D-type cyclin (CycD3). The accumulation ofCycD3 was rapidly induced when cytokinin was addedto wildtype cell cultures or applied to whole plants.Moreover, cells of cultured callus tissues from transgenicplants that were constructed to overproduce CycD3continue to divide in the absence of cytokinin.

While it is well established that the B-type cyclinsact at the transition from G2 to mitosis, the level ofD-type cyclins in plants remains fairly constant through-out the cell cycle and no specific activity for D-typecyclins has yet been described. However, based on anal-ogy with similar cyclins in animals, it has been proposedthat they control the transition from G1 to S phase. Ifthis is true, the Arabidopsis results would indicate thatcytokinin initiates cell division at the G1 to S transi-tion through the induction of CycD3. Interestingly, twoother Arabidopsis D-type cyclins (CycD2 and CycD4)are induced by sucrose. As more is learned about the

links between cytokinins or other developmental signalsand cell cycle regulatory proteins, we can look forwardto a better understanding not only of how cell divisionis initiated but also how cells exit the cell cycle in orderto begin differentiation.

20.3.2 THE RATIO OF CYTOKININ TOAUXIN CONTROLS ROOT ANDSHOOT INITIATION IN CALLUSTISSUES AND THE GROWTH OFAXILLARY BUDS

Auxin and cytokinins have antagonistic actions withrespect to root shoot formation in cultured tobaccotissues (Figure 20.5). Both auxin and cytokinins arerequired to maintain callus cultures. However, whenauxin is present alone, or if the ratio of auxin to cytokininis high, cultures will initiate root formation. Conversely,a high cytokinin-to-auxin ratio promotes shoot produc-tion and roughly equal amounts of auxin and cytokininwill cause continued proliferation of undifferentiatedcallus. This phenomenon has been put to practical appli-cation in the technique of regenerating large numbersof plants by micropropagation (Box 20.2).

Cytokinins also antagonize the auxin effect in reg-ulating the growth of axillary buds, or apical domi-nance (Chapter 18). In many species the applicationof cytokinins either to the shoot apex or directly tothe axillary bud will release the bud from inhibition.Tomato mutants expressing strong apical dominancecontain lower amounts of cytokinins than those withnormal dominance. More recent studies with transgenicplants have confirmed that plants with elevated auxinlevels show increased apical dominance and that over-production of cytokinin reduces apical dominance. Most

FIGURE 20.4 A simplified model for hor-monal control of the cell cycle in plants.Cytokinin promotes the onset of mito-sis (the G2 to M transition) by activatinga phosphatase that removes an inhibitoryphosphate group from the cyclin-dependentkinase (CDK)/cyclin B complex. Auxin andcytokinin also promote the accumulation ofG1 cyclins (shown here as cyclin D), neces-sary for the onset of the S (synthesis) phase.

M

Cell Cycle

Cytokinin-dependent

phosphataseG2

G1

SCycB

CycD

CycD

AuxinCytokinin

CDK

CDK

P

CDK

P

P CDK

CycB

CDK

CycB

20.3 Cytokinins are Required for Cell Proliferation 345

CH2HN

NHN

N N O6

BOX 20.2TISSUE CULTUREHAS MADEPOSSIBLELARGE-SCALECLONING OFPLANTS BY MICRO-PROPAGATION

With a relatively small investment in space, technicalsupport, and materials, tissue culture has made it possibleto produce literally millions of high-quality, geneticallyuniform plants. The process is known as micropropa-gation. The most common technique is to place excisedmeristematic tissue on an artificial medium containing acytokinin/auxin ratio that reduces apical dominance andencourages axillary bud development (See Figure 20.5).The new shoots can be separated and sub-cultured toproduce more axillary shoots, or placed on a mediumthat encourages rooting. Once roots appear, the plantletscan be planted out and allowed to develop into matureplants. Alternatively, excised tissues can be used to estab-lish callus cultures, which may then be induced to formroots and shoots by manipulating the cytokinin/auxinratio.

Micropropagation can also be an effective way toeliminate viruses and other pathogens and produce com-mercial quantities of pathogen-free propagules. Thefirst plants to be mass-produced by tissue culture were

virus-free orchids of the genus Cymbidium, but the tech-nique has also been found useful for potato, lilies, tulips,and other species that are normally propagated vegeta-tively. Potato, for example, is vegetatively propagatedthrough buds on the tubers, a system that readily trans-mits viruses to the next generation. Micropropagationof potato from meristem cultures has proven to be aneffective way to isolate virus-free lines.

Micropropagation is also used extensively in theproduction of forest tree species. Here the propagulesare generated primarily from cultures of axillary andadventitious buds; callusing and differentiation of newbuds is rarely used. A similar approach has been appliedsuccessfully to cultivars of apple (Malus), peach (Pyrus),and pear (Prunus). Because most temperate fruits arehighly heterozygous, they do not breed true from seedbut are propagated by vegetative cuttings. Rooting ofmicrocuttings in culture is now a routine procedurein many commercial laboratories. By the early 1980s,growers in the Netherlands were producing more than21 million plants by micropropagation.

In spite of the fact that plantlets derived from tissueculture are cloned from presumably identical somatic(nonsexual) cells, the regenerated plants can exhibit sig-nificant variation in their morphology and physiology.This is known as somaclonal variation. The causeof somaclonal variation is not clear, but it appears toinvolve spontaneous genetic mutation as a result of theculture conditions. The value of somaclonal variation isthat occasionally the variants exhibit some agronomi-cally useful trait, such as disease resistance or variationsin floral color patterns.

interesting, however, is the observation that an exoge-nous application of cytokinin to the suppressed apicalbuds of auxin-overproducing plants will release the budsfrom dominance. Thus, it appears that it is the ratio ofauxin to cytokinin, not the absolute level of auxin, whichsuppresses axillary bud growth.

The cytokinin–auxin antagonism is believed toaccount for the phenomenon of ‘‘witch’s broom,’’ anexample of extreme axillary bud release (Figure 20.6).The witch’s broom syndrome appears on a wide vari-ety of plants and most often results from parasitismby fungi, bacteria, or mistletoes (Arceuthobium sp.), adwarf flowering shrub that parasitizes predominantlyspruce (Picea), larch (Larix), and pine (Pinus). Althoughthe exact nature of the hormonal imbalance is notknown, it is believed that the parasitism stimulatesan overproduction of cytokinin. The resulting releaseof apical dominance produces a dense mass of shortbranches.

20.3.3 CROWN GALL TUMORS AREGENETICALLY ENGINEEREDTO OVERPRODUCE CYTOKININAND AUXIN

Agrobacterium tumifaciens is a bacterium that causes neo-plastic or tumorous growth, known as crown gall, onstems (Figure 20.7). Crown gall tissues can be excisedand maintained on a simple medium containing mineralsalts, some vitamins, and sucrose as a carbon source. Nohormones need be added to the medium because theinfected tissues have been naturally engineered with thecapacity to synthesize both cytokinins and auxin.

A. tumefaciens contains a typical bacterial plasmid—a small circular strand of DNA separate from the restof the bacterial genome. The A. tumefaciens plasmid iscalled a tumor-inducing, or Ti, plasmid because it car-ries the genes responsible for the neoplastic growth thatoccurs when the bacterium infects a wounded plant. A

346 Chapter 20 / Hormones III: Cytokinins

FIGURE 20.5 Shoot regeneration in callus culture. Apiece of pith tissue from the center of a tobacco stemwas explanted onto a medium containing mineral salts,vitamins, sucrose, auxin (left), or auxin plus cytokinin(right). The tissue grown on auxin alone proliferated asan undifferentiated callus. The tissue grown on auxinplus cytokinin regenerated shoots and roots. Theseplantlets can be eventually planted into soil and will pro-duce a fully competent, mature tobacco plant.

portion of the plasmid DNA, known as T-DNA (fortransferred DNA), contains the genes for three classesof proteins. Two of these are the enzymes necessaryfor the synthesis of cytokinins and auxins. The third isthe enzyme that causes the plant to produce opines,unusual amino acids that serve as nutrients for the bac-terium. When A. tumefaciens invades a host plant, theT-DNA portion of the plasmid is inserted into the hostcell nuclear DNA. These genes are then replicated alongwith the host cell genome. A. tumefaciens is in fact a natu-ral genetic engineer. It transforms the host cell, which isthen programmed to overproduce cytokinins, auxin, andopines. The cytokinins and auxin together encourage

FIGURE 20.6 Witch’s broom on white pine (Pinus strobus).Witch’s broom is the result of a fungal infection thatstimulates an overproduction of cytokinin. The resultis an uncontrolled release of axillary bud development.

FIGURE 20.7 Crown gall is a neoplastic, or tumorous,growth shown here on the stem of a Bryophyllum plant.Crown galls are the result of an infection with the bac-terium Agrobacterium tumefaciens. The bacterium trans-forms the host plant cells with bacterial genes that causean overproduction of auxin and cytokinin.

cell proliferation and neoplastic growth, while opinesfeed the invading bacterium.

20.3.4 CYTOKININS DELAYSENESCENCE

At present, there are three lines of evidence indicating arole for cytokinins as inhibitors of senescence. First is theobservation that exogenous application of cytokinin todetached leaves will delay the onset of senescence, main-tain protein levels, and prevent chlorophyll breakdown.Application of cytokinins will also delay the naturalsenescence of leaves on intact plants.

The second line of evidence consists of correlationsbetween endogenous cytokinin content and senescence.For example, detached leaves that have been treatedwith auxin to induce root formation at the base ofthe petiole will remain healthy for weeks. In thiscase, the growing root is a site of cytokinin synthesisand the hormone is transported through the xylem tothe leaf blade. If the roots are continually removed asthey form, senescence of the leaf will be accelerated. Ithas also been observed that when a mature plant beginsits natural senescence, there is a sharp decrease in thelevel of cytokinins exported from the root.

A third and particularly convincing line of evidencecomes from recent studies employing recombinantDNA techniques. Tobacco plants (Nicotiana tobacum)have been transformed with the Agrobacterium gene forcytokinin biosynthesis, designated TMR (Figure 20.8).The Agrobacterium TMR gene encodes for the enzymeisopentenyl transferase, which catalyzes the rate-limiting step in cytokinin biosynthesis. In this case, the

20.3 Cytokinins are Required for Cell Proliferation 347

A. B. C. D.

FIGURE 20.8 Cytokinin control of senescence and bud growth in tobacco. Tobaccocallus cells, genetically transformed such that cytokinin production could be stim-ulated by heat shock, were allowed to regenerate plantlets. (A) Transformedheat-shocked plantlets. (B) Untransformed heat-shocked plantlets. (C) Transformedcontrols (no heat shock). (D) Untransformed controls. Note especially the prolif-eration of lateral buds and absence of senescence in the transformed, heat-shockedplantlets. The large, white areas in B and D are senesced leaves. The transformedcontrols (C) do not, as expected, exhibit the cytokinin effect on lateral bud growth butdo not exhibit senescence. This probably indicates the transformed gene is ‘‘leaky’’and a small amount of cytokinin is produced in the absence of heat shock. (FromSmart, C. et al. 1991. The Plant Cell 3:647. Copyright American Society of PlantPhysiologists. Photo courtesy of C. Smart.)

TMR gene was linked to a heat shock promoter. Apromoter is a sequence of DNA that signals wherethe transcription of messenger RNA (mRNA) shouldbegin. The heat shock promoter is normally involvedin the heat shock response of plants, which is inducedby a brief period of high temperature. Nomally, theheat shock response involves the synthesis of a newset of proteins called heat shock proteins. The heatshock promoter is thus active only when subjected to ahigh temperature treatment. By linking the TMR geneto the heat shock promoter, cytokinin biosynthesiscan be turned on in the transformed plants simplyby subjecting the plants to a brief period of hightemperature. A heat shock of 42◦C for 2 hours causeda 17-fold increase in zeatin levels in transformed plantscompared with untransformed control plants. Whensubjected to heat shock on a weekly basis over a 12-weekperiod, transformed plants exhibited a marked releaseof lateral buds from apical dominance as well as delayedsenescence. Transformed but non-heat-shocked plantsalso remained green longer than normal plants butdid not exhibit release from apical dominance. This isprobably due to ‘‘leakiness’’ on the part of the promoter,allowing production of a very small but effective amountof cytokinin even at normal temperature.

The mechanism by which cytokinins are able todelay senescence is not clear, but there is some evidencethat cytokinins exert a role in mobilizing nutrients. The

classic experiment of K. Mothes and coworkers, sum-marized in Figure 20.9, illustrates this point. In thisexperiment, a nutrient labeled with radioactive carbon(e.g., 14C-glycine) is applied to a leaf after a portionof the leaf has been treated with cytokinin. Invariablythe radioactivity is transported to and accumulates inthe region of cytokinin treatment. A variety of similarexperiments have led to the hypothesis that cytokininsdirect nutrient mobilization and retention by stimu-lating metabolism in the area of cytokinin application.This creates a new sink—an area that preferentiallyattracts metabolites from the region of application (thesource). It is unlikely that cytokinins act directly throughstimulating protein synthesis since the mobilizationof nonmetabolites such as α-aminoisobutyric acid isdirected by cytokinins equally well.

20.3.5 CYTOKININS HAVE ANIMPORTANT ROLE INMAINTAINING THE SHOOTMERISTEM

The fundamental characteristic of a plant meristem isthe capacity to maintain the spatial distinction betweendividing cells and differentiating cells. The entire courseof plant development depends on maintaining that smallpopulation of perpetually dividing cells. Ever sinceSkoog and Miller demonstrated that cytokinins inducedcell division and shoot regeneration fifty years ago,

348 Chapter 20 / Hormones III: Cytokinins

Kinetin treatedControl

Kinetin Kinetin

FIGURE 20.9 Diagram of an experiment demonstratingthe role of cytokinin in nutrient mobilization. Radioac-tive 14C-aminobutyric acid was applied to the area indi-cated by the dark spot. Left: Control, no cytokinin treat-ment. Radioactivity spreads into the vascular tissue forexport through the petiole. Center: Radioactivity accu-mulates in the left half of the leaf, which has been treatedwith kinetin. Right: Radioactivity is retained near thepoint of application when the right half of the leaf istreated with kinetin. (Based on experiments of Mothes,K. 1963. Regulateurs Naturels de la Croissance Vegetale,123. Centre National de la Recherche Scientifique.)

cytokinins have been routinely used to induce shootformation in tissue culture for plant propagation and forthe production of transgenic plants. It has always beenassumed that cytokinins had a significant role in main-taining the meristem in planta, but direct evidence forsuch a role has been difficult to obtain. The traditionalmethod involving exogenous application of cytokininsis something of a shotgun approach that provides rela-tively little solid information. When you simply spray aplant with hormone solution, for example, it is virtuallyimpossible to know how much hormone actually getsinto the plant or where it goes. Moreover, excessivehormone levels may cause artifactual, nonphysiologiceffects.

Several new lines of evidence, however, now pointto a positive role for cytokinins in the shoot apicalmeristem. Most of these experiments involve reducingthe in planta concentration of cytokinins, either byoverexpressing appropriate genes in transgenic plantsor through loss-of-function mutants. For example,cytokinin levels can be reduced in planta by overex-pressing the genes for cytokinin oxidase/dehydrogenase(CKX), which degrades active cytokinins. Arabidopsishas seven CKX genes and, depending on which of thesegenes is overexpressed, the cytokinin content can bereduced to 30 to 45 percent of wildtype plants. Theresult in all cases was retarded shoot development;dwarfed, late flowering plants; and reduced size of the

shoot meristem. The formation of leaf primordia wasslower in cytokinin-deficient plants and the numberof leaf cells was significantly reduced. Where CKXwas most strongly expressed, growth of the shootwas arrested completely shortly after germination.Similar results were obtained in experiments where thecytokinin content was reduced through loss-of-functionmutants of the gene for IPT or of genes for knowncytokinin receptors (Section 20.4).

Further evidence for the maintenance of the shootapical meristem by cytokinins is offered by the discoveryof a cytokinin-deficient rice mutant that was given theintriguing name of lonely guy (log). Rice flowers areborne in a typical, highly branched grass inflorescencecalled a panicle (Figure 20.10). The flowers, or spikelets,normally contain a single pistil surrounded by severalstamens. In log mutant plants, the size of the paniclewas severely reduced. There were fewer branches andthe branches bore abnormal flowers. Flowers were oftenreduced to no pistil and but a single stamen (hence thename lonely guy). Microscopic studies revealed that afterthe transition from vegetative to reproductive stage thenormally dome-shaped floral meristem flattens and thedifferentiation of floral organs is prematurely shut down.

Clearly the log mutant is characterized by a deficientmeristem, but why? When the LOG gene was isolatedand cloned, it was found that expression of the LOGgene is localized in regions of active cell division in themeristem such as the apex of the meristem and branchprimordia. It was also found that LOG encodes a phos-phoribohydrolase enzyme. This means that the enzymeactivates cytokinins by removing the ribose phosphategroup from an inactive cytokinin nucleotide to leavethe active free base. The absence of this enzyme in thelog mutant thus reduces the level of active cytokinin inthe critical region of cell division and the result is animproperly maintained meristem.

20.3.6 CYTOKININ LEVELS IN THESHOOT APICAL MERISTEM AREREGULATED BY MASTERCONTROL GENES

If cytokinins maintain the meristem by controllingcell division, then what triggers differentiation? Plants,fungi, and animals all contain genes that are involvedin patterning and organ initiation. Most of these genesshare a sequence of about 180 base pairs called thehomeobox or homeodomain. The resulting proteinsare therefore referred to as homeobox or homeodomainproteins. In plants, the homeobox proteins are identifiedas KNOX proteins, based on the KNOTTED1 proteinin maize—the first homeobox protein to be identified inplants. Homeobox genes are often referred to as mastercontrol genes because of their fundamental role in devel-opment and their capacity to specify the fate of tissues

20.3 Cytokinins are Required for Cell Proliferation 349

Log1 mutantwildtype

B.

A. C.

D.

stamenstamens

paleapaleapistil

FIGURE 20.10 Diagrammatic representations of the phe-notypes of wildtype and the cytokinin-deficient log-1mutants in rice (Oryza sativa). The wildtype rice inflo-rescence (A) is a typical grass panicle with numer-ous branches covered with lateral flowers, or spikelets(shown here in green). Each branch ends with a termi-nal spikelet (shown in red). The number of branchesand spikelets is determined by the timing of when theshoot apical meristem of the main stem or branches isconverted to a terminal spikelet meristem. The wild-type spikelet contains a pistil and numerous stamens (B).With less cytokinin in the mutant, the branch meristemsare not properly maintained. The result is fewer, shorterbranches with fewer spikelets (C). Floral meristems arealso affected and the spikelet may be reduced to a sin-gle stamen and no pistil (D). (Based on Kurakawa, T. etal. 2007. Nature 445:652; and Kyozuka, J. 2007. CurrentOpinion in Plant Biology 10:442.)

and organs. In the case of plants, one fundamental roleof KNOX genes is to specify meristems by regulatingthe biosynthesis of cytokinins and gibberellins.

The impact of KNOX genes on development is illus-trated by two types of mutations. Expression of wildtypeKNOX genes is localized to the nuclei of shoot apicalmeristems. They are not normally expressed in leaf pri-mordia or developed leaves of wildtype plants. Thereare several dominant mutations, however, that alter thispattern. These dominant mutations are gain-of-functionmutations, meaning that the genes are overexpressedin the mutant seedlings. The result is that some leaftissues fail to differentiate and continue to divide, form-ing sporadic outgrowths, or ‘‘knots,’’ on the leaf blade.In extreme cases, ectopic shoots can actually be seen

developing on the leaf surface, indicating the presence ofactive meristem-like cells. In other words, when KNOXproteins accumulate due to overexpression of the KNOXgenes, cells in the leaf which should have switched todifferentiation fail to do so and continue dividing.

The Arabidopsis gene SHOOTMEREISTEMLESS(STM) also encodes a KNOX protein and the effect ofthe recessive loss-of-function mutant, stm, is just theopposite of the gain of function dominant mutatntsdescribed above. Plants carrying the mutant stm fail todevelop any shoot apical meristem during embryogen-esis. The pattern of STM expression is very similar tothe pattern of KNOTTED1expression in maize. Usingtechniques to visualize mRNA in situ, the pattern ofSTM mRNA was followed during embryogenesis. STMmRNA initially appears in one or two cells in theearlier globular stage embryos (Figure 20.11). As theembryo enters the heart-shaped and torpedo stages,STM expression is restricted to the notch between theembryonic cotyledons— i.e., expression of STM is lim-ited to the cells that will eventually organize as theSAM. As the plant grows into the seedling stage andadult plant, STM expression persists into the vegetative,axillary, inflorescence, and floral meristems. But it is notexpressed in leaves or leaf primordia.

It thus appears that the Knox proteins are transcrip-tion factors based almost exclusively in the meristem andthat regulate the switch between indeterminant (i.e.,meristematic) growth and differentiation. If so, whatgenes are targeted by these transcription factors? Theprimary candidates appear to be genes that control thesynthesis of gibberellins and cytokinins. We’ve alreadyseen that LOG maintains the meristem by catalyzingthe formation of active cytokinins. In early studies ofKNOX proteins, it was noted that overexpression ofthese transcription factors was accompanied by a sig-nificant increase in the cytokinin content, especiallytrans-zeatin (tZ) and isopentenyl adenine (iP). Recentstudies in rice (Oryza sativa) have confirmed that thehigher cytokinin levels found in transgenic plants over-producing KNOX proteins is a direct result of increasedtranscription of the rate-limiting enzyme isopentenyltransferase (IPT).

At the same time, it has been reported that KNOXproteins suppress the transcription of GA20-oxidasegenes in at least four different species: Arabidopsis,tobacco (Nicotiana tabaccum), rice (Oryza sativa), andpotato (Solanum tuberosum). GA20-oxidase catalyzes theconversion of inactive GA20 to the active gibberellin,GA1. Thus it appears that cytokinins are primarily res-ponsible for initiating and maintaining populations ofdividing cells. Gibberellins, on the other hand, are moreinvolved in the subsequent differentiation of cells. Thebalance between division and differentiation is main-tained by KNOX proteins; transcription factors whosejob it is to maintain a high CK/GA ratio in the shoot

350 Chapter 20 / Hormones III: Cytokinins

FIGURE 20.11 Schematic repre-sentation of STM expression dur-ing embryogenesis in Arabidop-sis. STM mRNA, shown in blue,is first detected in a single cell inthe early globular stage embryo(32–64 cell stage) (Left). In the later,torpedo-stage embryo, STM mRNAis confined to the nascent meris-tem (Right). C = cotyledons. (Basedon Long, J. A. et al. 1996. Nature379:66–69.) Shoot/Root axis

apical meristem. This is accomplished by simultane-ously stimulating cytokinin biosynthesis and suppressinggibberellin biosynthesis. The resulting high-CK/lowGA condition is required for both the formation andmaintenance of the meristem. Outside the meristem,KNOX genes are apparently turned off, cytokinin con-centrations decline, cell division effectively ceases, andgibberellins take over to encourage differentiation.

20.4 CYTOKININ RECEPTORAND SIGNALING

In spite of the fundamental role played by cytokinins incell division, the multiple other effects that cytokininshave on plant development have made it difficult toidentify cytokinin receptors and signal chains. It hasonly been within the last decade, more than fifty yearsafter Skoog and Miller purified the first cytokinin, thatthe first genes involved in cytokinin signaling have beenidentified.

The cytokinin receptor was finally discovered byT. Kakimoto and his colleagues who developed an Ara-bidopsis hypocotyl test to screen for mutants. Hypocotylsections, or explants, respond to added cytokinins bytypical cytokinin responses: rapid cell proliferation,greening, and shoot formation. The cytokinin response1(cre1) mutant shows none of these responses, evenwith a tenfold increase in cytokinin concentration. Thiswould be expected if the cytokinin receptor were eithermissing or nonfunctional in the mutant. Subsequentexperiments confirmed that the wildtype protein CRE1was in fact a cytokinin receptor. The same gene has alsobeen identified as WOODENLEG (WOL), so namedbecause its mutation retarded root growth, and Ara-bidopsis HISTIDINE KINASE 4 (AHK4).

20.4.1 THE CYTOKININ RECEPTOR ISA MEMBRANE-BASED HISTIDINEKINASE

CRE1 is the first component of a two-component reg-ulatory system—a type of regulatory system previouslyknown to operate in bacteria and other prokaryotes. Thename comes from the bacterial configuration where thereceptor (or sensor)—the first component—activatesa response regulator (RR)—the second component.Response regulators in turn either regulate the tran-scription of target genes or modulate other metabolicreactions. In addition to serving as hormone recep-tors, two-component regulatory systems also functionin osmosensing (Chapter 1), light sensing, and otherforms of sensory perception.

CRE1 is an intracellular histidine kinase (HK)with three domains (Figure 20.12). The sensor domain,at the N-terminal end of the protein, includes two small,hydrophobic membrane-spanning regions that anchorthe receptor in the plasma membrane. Between themis a hydrophilic loop that extends into the extracellu-lar space. This loop includes the cytokinin-binding sitesince a mutation at this site interferes with cytokininbinding and renders the receptor inoperative. The his-tidine kinase domain is located on the cytoplasmic sideof the membrane. The term kinase identifies HK as anenzyme involved in a phosphorylation reaction and thereference to histidine kinase means that the phospho-ryl group is added to a histidine residue. The phosphateattaches to a specific histidine residue (His459) in the his-tidine kinase domain. The histidine kinase also includestwo receiver domains, Da and Db.

Once it became evident that the acceptor was ahistidine kinase, the fully sequenced Arabidopsis genomecould be searched for other potential componentsof the signaling system. Arabidopsis is now known to

20.4 Cytokinin Receptor and Signaling 351

PP

RECEIVERDOMAINS

HISTIDINEKINASE DOMAIN

(HK)

HIS HIS HIS

MEMBRANE

INPUT DOMAIN

Da

D6Asp Asp Asp

CK

FIGURE 20.12 Predicted structure of the cytokinin receptor CRE1. The monomer(left) has three domains and two small membrane-spanning hydrophobic regions.Binding with cytokinin (right) induces dimerization and autophosphorylation. Thelocations of the histidine and aspartic acid phosphate-binding residues are indicated.

have the genes for eight different histidine kinases, sixhistidine-phosphotransfer proteins, and 23 responseregulators. Only three of the HK genes (CRE1,AHK2, AHK3) encode cytokinin receptors. At leasttwo are ethylene receptors and one is believed to bean osmosensor. The reason for such a large numberof response regulators appears to be that many areexpressed in a tissue-specific manner, which allows fora more finely tuned, tissue-specific signaling output.

20.4.2 THE CYTOKININ SIGNALINGCHAIN INVOLVES A MULTISTEPTRANSFER OF PHOSPHORYLGROUPS TO RESPONSEREGULATORS

A general scheme for cytokinin signaling is shown inFigure 20.13. Binding of a cytokinin molecule to thesensor domain induces dimerization and subsequent

352 Chapter 20 / Hormones III: Cytokinins

HIS

PlasmaMembrane

NuclearMembrane

Modulation ofOthereffectors

A-type-RRB-type-RR

Transcription of primaryresponse genes

PP

P

P

P

P

P

P D

HPT

HPT

HPT

D

D

D

D

D

H H

HPT

FIGURE 20.13 A model for cytokinin signal transduction via a multistep phosphorelay system. Cytokinin sensing occurswhen cytokinin binds with the input domain in the extracellular space. Cytokinin binding induces dimerization andautophosphorylation of the acceptor histidine kinase. The phosphorelay system begins with the transfer of the phos-phoryl group first to an aspartic acid residue (D) in the receiver domain and then to a histidine residue in a separatehistidine phosphotransfer protein (HPT). The phosphorylated HPT migrates into the nucleus where the phosphorylgroup is transferred to either a B-type or A-type response regulator (RR). The activated B-type response regulatorthen activates transcription of cytokinin primary response genes, including the A-type response regulator. The A-typeresponse regulators may down-regulate cytokinin responses by suppressing the activation of B-type response regulators.Alternatively, A-type response regulators may positively or negatively modulate other cytokinin.

Chapter Review 353

phosphorylation of a histidine residue in each of thetwo receptor molecules. Whereas most kinase enzymescatalyze the addition of a phosphoryl group to a sec-ond molecule, the histidine kinase receptor autophos-phorylates, which means that it phosphorylates itself.The phosphoryl group is then spontaneously trans-ferred to an aspartic acid residue on the Db receiverdomain.

The classic prokaryote two-component sys-tem is comprised of only a receptor kinase anda response regulator and the response regu-lator is activated by receiving a phosphoryl groupdirectly from the histidine kinase. In plants, thephosphoryl group is passed bucket-brigade fashionthrough one or more histidine-phosphotransferproteins (HPTs). The phosphoryl group is transferredfrom the receiver domain of the histidine kinaseto a histidine residue on the HP protein. Thephosphorylated HP protein then migrates into thenucleus where the phosphoryl group is transferred toan Asp residue in a response regulator. Note that thetransfer is alternately from histidine to aspartic acidto histidine to aspartic acid. The system that transfersphosphoryl groups between the various HKs, HPTs,and RRs is referred to as a phosphorelay network.

There are two classes of response regulators—theA-type and the B-type. The role of response reg-ulators is still being worked out, but in general, itappears that B-type response regulators are transcrip-tion factors. When activated by phosphorylation, B-typeresponse regulators induce the expression of genes thatare responsible for some cytokinin-regulated responses.Among the target genes for B-type response regu-lators are the genes for A-type response regulators.A-type response regulators, however, are not transcrip-tion factors and do not regulate gene expression. Theyapparently modulate cytokinin responses by influencingother aspects of metabolism.

The cytokinin system also has a built-in capacityfor shutting down the phosphorelay network whenno cytokinin is present. This conclusion is based onthe finding that CRE1, aside from its kinase function,also exhibits phosphatase activity. The activity of aphosphatase enzyme is the opposite of a kinase—aphosphatase removes phosphoryl groups. Thus, inthe absence of cytokinin, CRE1 reverses the process,unloads phosphoryl groups from HPTs, and quicklyinactivates the cytokinin response pathway

SUMMARY

Cytokinins are N6 adenine derivatives with either anisoprenoid-related side chain or an aromatic side chain.

The most common naturally occurring cytokinins areisopentenyl adenine and trans-zeatin.

Synthesis of cytokinins begins with the addition ofdimethylallyl pyrophosphate to an adenine nucleotide(AMP, ADP, or ATP). The nucleotide-phosphategroup is cleaved off to generate the active form.Although it has long been known that cytokinins aresynthesized in the roots and translocated to the shootthrough the xylem sap, it is now clear the some tissues,meristems in particular, are under the control oflocally produced cytokinins. Cytokinins are deactivatedby conjugation with sugars and amino acids, or byoxidative degradation.

Cytokinins, commonly in concert with auxin, areinvolved in numerous developmental responses. Theseinclude regulation of cell division, shoot and root initia-tion, release of axillary bud growth, delay of senescence,and maintenance of an actively dividing shoot apicalmeristem.

Cytokinin receptors are of a class known ashistidine kinases. The signal chain involves a multisteptransfer of phosphoryl group transfer referred toas a phsophorelay network. The final targets areA- and B-type response regulators. When activatedby phosphorylation, the responses regulators eitheractivate transcription of cytokinin primary responsegenes or modulate other aspects of cytokinin relatedmetabolism.

CHAPTER REVIEW

1. Auxins are identified by their control of cellenlargement in excised tissues and gibberellinsare identified largely on the basis of chemicalstructure. How are cytokinins identified?

2. Describe how cytokinins regulate the cell cycle.3. The bacterium Agrobacterium is commonly

used to produce transgenic plants. Whatcharacteristic(s) make it useful in this regard?

4. In controlling various developmental responses,cytokinins frequently require the presenceof auxin. Can you think of a reason whycytokinins would require the presence of auxin?

5. What is the evidence that cytokinins are requiredfor the maintenance of the shoot apical meristem?

6. What role do KNOX proteins have in maintainingthe apical meristem?

7. What is the apparent role of gibberellins in theshoot apical meristem?

8. What is a two-component regulatory sys-tem? How does a two-component regula-tory system relate to cytokinin activity?

354 Chapter 20 / Hormones III: Cytokinins

FURTHER READING

Davies, P. J. 2004. Plant Hormones: Biosynthesis, Signal Trans-duction, Action. Dordrecht: Kluwer Academic Publishers.

Ferreira, F., J. J. Kleiber. 2005. Cytokinin signaling. CurrentOpinion in Biology 8:518–525.

Heyl, A., T. Schmulling. 2003. Cytokinin signal percep-tion and transduction. Current Opinion in Plant Biology6:480–488.

Kurakawa, T. et al. 2007. Direct control of shoot meris-tem activity by a cytokinin-activating enzyme. Nature445:652–655.

Kyozuka, J. 2007. Control of shoot and root meristemfunction by cytokinin. Current Opinion in Plant Biology10:442–446.

Mok, D. W., M. C. Mok. 2001. Cytokinin metabolism andaction. Annual Review of Plant Physiology and Plant Molec-ular Biology 52:89–118. (This series of reviews is acces-sible in most data bases under the current title: AnnualReview of Plant Biology.)

Muller, B., J. Sheen. 2007. Advances in cytokinin signaling.Science 318:68–69.

Sakakibara, H. 2006. Cytokinins: Activity, biosynthesis, andactivity. Annual Review of Plant Biology 57:431–449.

Shani, E., O. Yanai, N. Ori. 2006. The role of hormones inshoot apical meristem function. Current Opinion in PlantBiology 9:484–489.

ABA

A- PA

ABA

ER

K+Ca2+

K+

in

Ca2+channel

Kinase

TPP

H+

ATP

Ca2+

Guard cell [Ca2+]in

PLDR

G

K

out

21Hormones IV: Abscisic Acid, Ethylene,

and Brassinosteroids

Abscisic acid (ABA), ethylene, and brassinosteroids(BR) are three hormone classes that are noted for theirinvolvement in a wide variety of plant responses as wellas extensive interactions with auxins and gibberellinsand, to a lesser extent, with cytokinins. Most of thecurrent work on each of these three hormones focuseson working out their physiology, metabolism, and modeof action by screening for mutations.

In this chapter, we will take each of these threehormones in turn and look at

• their biosynthesis and metabolism,• the range of physiological effects, and• our current understanding of signal perception and

transduction.

21.1 ABSCISIC ACID

Unlike auxins, gibberellins, and cytokinins, the hormoneabscisic acid (ABA) is represented by a single 15-carbonsesquiterpene (Figure 21.1). ABA also appears to havea more limited range of specific effects than auxins,gibberellins, and cytokinins. The name is based on theonce held belief that it was involved in the abscission of

leaves and other organs. It now appears to have nothingto do with abscision, but the name has stuck (Box 21.1).

The primary functions of ABA are (1) prohibitingprecocious germination and promoting dormancy inseeds and (2) inducing stomatal closure and the produc-tion of molecules that protect cells against desiccationin times of water stress. ABA has also been implicatedin other developmental responses, including the induc-tion of storage protein synthesis in seeds, heterophylly(leaves of different shape on the same plant), initiationof secondary roots, flowering, and senescence.

21.1.1 ABSCISIC ACID IS SYNTHESIZEDFROM A CAROTENOIDPRECURSOR

Once the structure of ABA had been determined,two possible pathways for the synthesis of ABA wereproposed. In the ‘‘direct pathway,’’ ABA would be syn-thesized from a 15-carbon terpenoid precursor suchas farnesyl diphosphate (see Figure 19.4). By the late1970s it had been clearly established that this pathwaywas operative in certain fungal plant pathogens thatactively synthesized ABA, but not in plants themselves.According to the second, or ‘‘indirect pathway,’’ ABA

355

356 Chapter 21 / Hormones IV: Abscisic Acid, Ethylene, and Brassinosteroids

BOX 21.1THE DISCOVERYOF ABSCISIC ACID

As more investigators became interested in planthormone research, it soon became evident thatether extracts of plant material—used to extractauxins—frequently contained substances that inter-fered with the auxin response in the Avena coleoptilecurvature test. Initially, the principal interest ofinvestigators was to rid extracts of these interferingsubstances. As time went on, however, interest turnedtoward the possibility that these inhibitors mightthemselves be growth regulators in their own right.The advent of paper chromatography as an analyticaltool made it possible to achieve better separation ofthe various substances in crude extracts. In 1953,Bennet-Clark and Kefford reported that plant extractscontained, in addition to IAA, a substance that inhibitedgrowth of coleoptile sections, which they calledinhibitor β. The observation that large amounts ofinhibitor β could be isolated from axillary buds andthe outer layer of dormant potato tuber led Kefford tosuggest that it was involved in apical dominance andmaintaining dormancy in potatoes. Meanwhile, other

investigators reported the occurrence of inhibitorsin buds and leaves that appeared to correlate withthe onset of dormancy in woody plants. In 1964,P. F. Waring proposed the term ‘‘dormin’’ for theseendogenous, dormancy-inducing substances.

In another line of study, substances that acceler-ated abscission were isolated from senescing leaves ofbean and from cotton and lupin fruits. These substanceswould accelerate abscission when applied to excisedabscission zones and were called ‘‘abscission II.’’ Theseseveral lines of study came to a head in the mid-1960swhen three laboratories independently reported thepurification and chemical characterization of abscisinII, inhibitor β, and dormin. All three substances provedto be chemically identical.

It is not unusual in such cases that there was somedisagreement over what this substance should be called.Although abscisin II had priority (it was the first to becrystallized and chemically characterized), some felt theterm awkward and argued it did not adequately describeits range of effects. Finally, a panel of scientists activein research on abscisin II and dormin was charged withproposing an acceptable name. The name abscisic acidand abbreviation ABA were recommended by this panelto the 1967 International Conference on Plant GrowthSubstances, which met in Ottawa. The recommendationwas accepted by the Conference and the term abscisicacid is now in universal use.

FIGURE 21.1 Abscisic acid is a class of hormones repre-sented by a single compound.

was produced from the cleavage of a carotenoid suchas β-carotene. Originally proposed in the late 1960s,the indirect pathway was based on structural similar-ities between carotenoid pigments and ABA and hassince received support from a variety of biochemicalstudies, 18O2-labeling experiments, and, most recently,the characterization of ABA biosynthetic mutants. Thecleavage of carotenoids, especially β-carotene, to pro-duce useful biochemicals is not without precedent. Thecyanobacterium Microcystis, for example, produces a C10metabolite by cleavage of β-carotene. Mammals pro-duce vitamin A by cleavage of β-carotene and cleavage ofβ-carotene to produce 2 molecules of the photoreceptorretinal (C20) has been reported.

There is now a growing body of evidence sup-porting the indirect synthesis of ABA from β-carotenevia the 40-carbon terpene violaxanthin (Figure 21.2).First, a series of viviparous mutants in maize (describedfurther below) were found to have reduced levels ofboth carotenoids and ABA. These mutants, shown tobe affected in the early steps of carotenoid biosynthesis,establish a strong correlation between carotenoid andABA biosynthesis. Second, the carbon skeleton of ABAand the position of the oxygen-containing substituentsare very similar to that of violaxanthin. J. A. D. Zeevaartand his colleagues compared the incorporation of 18O2,a stable isotope of oxygen, into ABA in water-stressedleaves and turgid leaves of several species. The pat-tern of 18O2-enrichment in the carboxyl group of ABAwas consistent with the cleavage of a xanthophyll andits rapid conversion to ABA in water-stressed leaves.Third, it is known that violaxanthin can be degraded inthe light in vitro to a 15-carbon derivative, xanthoxin, anatural constituent of plants. If radio-labeled xanthoxinis fed to bean or tomato plants, some of the radioactivityappears in ABA. In ABA-deficient tomato mutants, how-ever, conversion of radio-labeled xanthoxin into ABA isreduced relative to wildtype plants. Finally, at least

21.1 Abscisic Acid 357

G3P+Pyruvate

MEPPathway

IPP

Vioaxanthin

Chloroplast

(C40)

(C5)

(C40)

(C15)

(C15)

(C15)

(C15)

Cytoplasm

Xanthoxin

Abscisic Aldehyde

Abscisic Acid

Xanthoxin+C25 byproduct

�-carotene

(NCED)

FIGURE 21.2 A flow sheet for the biosynthesis of abscisicacid. ABA biosynthesis begins in the chloroplast withthe synthesis of isopentenylpyrophosphate (IPP) fromglyceraldehydes-3-phosphate (G3P) and pyruviate viathe methylerythritol-4-phosphate (MEP) pathway. IPPin the chloroplast gives rise to a variety of C10, C20, andC40 terpenoids, including β-carotene. β-Carotene is con-verted to violaxanthin, which is cleaved by the enzymenine-cis-epoxycarotenoid dioxygenase (NCED) to yieldxanthoxin, a C15 precursor to ABA, and a 25-carbon‘‘byproduct.’’

two groups have reported a stoichiometric relationshipbetween losses of violaxanthin and increases in ABA instressed etiolated bean leaves.

Although ABA is synthesized in the cytosol, itsbiosynthetic pathway begins in the chloroplast (andpossibly other plastids in nongreen cells), which iswhere carotenoid pigments are produced (Figure 21.2.See also Figure 19.4). The critical enzyme is nine-cis-expoycarotenoid dioxygenase (NCED). This enzymecleaves the 40-carbon carotenoid violaxanthin to pro-duce a 15-carbon product, xanthoxin, and a 25-carbon‘‘by-product.’’ Xanthoxin is then converted to abscisicaldehyde by an alcohol dehydrogenase. Abscisic al-dehyde is in turn oxidized to abscisic acid by abscisicaldehyde oxidase. The enzyme NCED and, conse-quently xanthoxin production, is known to be targetedin the chloroplast while the alcohol dehydrogenase andabscisic aldehyde oxidase are located in the cytosol.This means that xanthoxin must migrate from thechloroplast into the cytosol, although the mechanismof migration is not yet known.

21.1.2 ABSCISIC ACID IS DEGRADEDTO PHASEIC ACID BYOXIDATION

Abscisic acid is rapidly metabolized when it is appliedexogenously to plant tissues. In wilted bean leaves,for example, the half-time for turnover (the time forone-half of the labeled ABA to be destroyed) was esti-mated to be about three hours. A glucose ester ofABA has been found in low concentration in a variety ofplants, but the principal metabolic route seems to be oxi-dation to phaseic acid (PA) and subsequent reduction ofthe ketone group on the ring to form dihydrophaseic acid(DPA) (Figure 21.3). At least some tissues appear to carrythe metabolism further to form the 4′-glucoside of DPA.DPA and its glucoside are both metabolically inactive.

21.1.3 ABSCISIC ACID IS SYNTHESIZEDIN MESOPHYLL CELLS, GUARDCELLS, AND VASCULAR TISSUE

There are a lot of open questions about the sites ofABA synthesis in the plant. Earlier physiological studiesindicated that abscisic acid was found predominantly inmature, green leaves, especially in water-stressed plants.This would fit with the more recent biochemical andgenomic studies described above showing that ABA pre-cursors originate in chloroplasts but ABA itself is formedin the cytoplasm. There is also evidence that ABA maybe stored in the chloroplasts (Chapter 13). At low pH,ABA exists in the protonated form ABAH, which freelypermeates most cell membranes. The dissociated formABA− is impermeant because it is a charged moleculethat does not readily cross membranes. In activelyphotosynthesizing mesophyll cells the cytosol will bemoderately acidic (pH 6.0 to 6.5) while the chloroplaststroma is alkaline (pH 7.5 to 8.0). Thus, ABAH diffusesreadily from the cytosol into the chloroplast stroma,where it dissociates and beomes trapped. This storedABA can later be released when photosynthesis shutsdown and the stroma pH declines.

Abscisic aldehyde oxidase (AAO) expression isinduced in guard cells under conditions of water stressand NCED expression has been detected in guard cellsof senescing leaves and cotyledons. Thus it appearsthat ABA is also synthesized directly in the guard cells.Furthermore, expression of ABA biosynthetic genes(NCED and others) has been localized in phloemcompanion cells and xylem parenchyma cells of fullyturgid plants. This indicates that vascular tissues arealso a site of ABA synthesis in unstressed plants.

Abscisic acid is highly mobile and moves quicklyout of the leaves to other parts of the plant, especiallysink tissues. For example, radioactively labeled abscisicacid applied to soybean leaves can be detected in theroots within 15 minutes. Developing seeds also import

358 Chapter 21 / Hormones IV: Abscisic Acid, Ethylene, and Brassinosteroids

FIGURE 21.3 Oxidative degradation of abscisic acid to phaseic acid and dihydrophaseicacid.

large amounts of abscisic acid from the leaves. There isalso some evidence that under conditions of water stress,ABA either stored or synthesized in the roots is rapidlyexported to the leaves (See Section 21.1.5).

21.1.4 ABSCISIC ACID REGULATESEMBRYO MATURATION ANDSEED GERMINATION

The development of embryos and subsequent germi-nation of the seed is characterized by often dramaticchanges in hormone levels (refer to Figure 16.10). Inmost seeds, cytokinin levels are highest during the veryearly stages of embryo development when the rate of celldivision is also highest. As the cytokinin level declinesand the seed enters a period of rapid cell enlargement,both GA and IAA levels increase. In the early stages ofembryogenesis, there is little or no detectable ABA. Itis only during the latter stages of embryo development,as GA and IAA levels begin to decline, that ABA levelsbegin to rise. ABA levels generally peak during the mat-uration stage, when seed volume and dry weight alsoreach a maximum, and then return to lower levels inthe dry seed. Maturation of the embryo is characterizedby cessation of embryo growth, accumulation of nutri-ent reserves in the endosperm, and the development oftolerance to desiccation.

The timing of ABA accumulation to coincide withembryo maturation reflects the critical role that ABAplays in the maturation process. One of the functions ofa seed, of course, is to disperse the population and ensuresurvival of the species through unfavorable conditions.A seed would be of little value if the embryo did notenter dormancy but continued to grow and establish anew plant before dispersal could occur. One functionof ABA is to prevent such precocious germination, orvivipary, while the seed is still on the mother plant.

The relationship between ABA and precocious ger-mination is clear. Vivipary can be chemically inducedin maize by treatment of the developing ear at theappropriate time with fluridone, a chemical inhibitorof carotenoid biosynthesis. Since carotenoids and ABAshare early biosynthetic steps, fluridone inhibits biosyn-thesis of ABA as well. Fluridone-induced vivipary can beat least partially alleviated by application of exogenous

ABA. Soybean embryos can be encouraged to germi-nate precociously by treatments such as washing or slowdrying, both of which lower the endogenous ABA level.Precocious germination will occur when the ABA con-centration is reduced to 3 to 4 μg per g fresh weight ofseed, a level that is not normally reached until the latestages of seed maturation.

The strongest indication of a role for ABA in pre-venting precocious germination, however, comes fromthe study of viviparous mutants. At least four viviparousmutants in maize (vp2, vp5, vp7, vp9) are known to beABA-biosynthetic mutants with reduced levels of ABA inthe seeds. One maize mutant, vp1, appears to have nor-mal ABA levels but is missing what is believed to be anABA-specific transcription factor. All of these mutantsgerminate prematurely on the cob before the seeds haveentered dormancy. Viviparous mutants are also knownfor Arabidopsis. ABA also stimulates protein accumula-tion in the latter stages of soybean embryo developmentand is known to prevent GA-induced α-amylase biosyn-thesis in cereal grains. All of these results establish astrong connection between ABA and seed maturationand/or prevention of precocious germination.

ABA also initiates desiccation of the seed, althoughthe mechanisms are unknown. This may involve ABAregulation of genes which encode proteins that areinvolved in desiccation tolerance.

21.1.5 ABSCISIC ACID MEDIATESRESPONSE TO WATER STRESS

Plants generally respond to acute water deficits by clos-ing their stomata in order to match transpirational waterloss from the leaf surface with the rate at which watercan be resupplied by the roots. Since the discovery ofABA in the late 1960s, it has been known to have aprominent role in stomatal closure during water stress.In fact, ABA has long been recognized as antitranspirantbecause of its capacity to induce stomatal closure andthus reduce water loss through transpiration.

ABA accumulates in water-stressed (that is, wilted)leaves and exogenous application of ABA is a powerfulinhibitor of stomatal opening. Furthermore, two toma-to mutants, known as flacca and sitiens, fail to accumulatenormal levels of ABA and both wilt very readily. Theprecise role of ABA in stomatal closure in water-stressed

21.1 Abscisic Acid 359

H2O

Substomatal airspace

Apoplast Symplast

Cuticle

FIGURE 21.4 ABA movements in the apoplast. ABA syn-thesized in the roots is carried to the leaf mesophyll cells(heavy arrows) in the transpiration stream (light arrows).ABA equilibrates with the chloroplasts of the photosyn-thetic mesophyll cells or is carried to the stomatal guardcells in the apoplast.

whole plants has, however, been difficult to decipherwith certainty. This is because ABA is ubiquitous, oftenoccurring in high concentrations in nonstressed tissue.Also, some early studies indicated that stomata wouldbegin to close before increases in ABA content could bedetected.

According to current thinking, the initial detectionof water stress in leaves is related to its effects onphotosynthesis. Inhibition of electron transport andphotophosphorylation in the chloroplasts would disruptproton accumulation in the thylakoid lumen and lowerthe stroma pH. At the same time, there is an increase inthe pH of the apoplast surrounding the mesophyll cells.The resulting pH gradient stimulates a release of ABAfrom the mesophyll cells into the apoplast, where it canbe carried in the transpiration stream to the guard cells(Figure 21.4).

As noted above, wilted leaves accumulate largequantities of ABA. In most cases, however, stomatalclosure begins before there is any significant increase inthe ABA concentration. This could be explained by therelease of stored ABA into the apoplast, which occursearly enough and in sufficient quantity—the apoplastconcentration will at least double—to account for initialclosure. Increased ABA synthesis follows and serves toprolong the closing effect.

Stomatal closure does not always rely on the per-ception of water deficits and signals arising within theleaves. In some cases it appears that the stomata close inresponse to soil desiccation well before there is any mea-surable reduction of turgor in the leaf mesophyll cells.Several studies have indicated a feed-forward controlsystem that originates in the roots and transmits infor-mation to the stomata. In these experiments, plants are

grown such that the roots are equally divided betweentwo containers of soil (Figure 21.5A). Water deficitscan then be introduced by withholding water fromone container while the other is watered regularly.Control plants receive regular watering of both con-tainers. Stomatal opening along with factors such asABA levels, water potential, and turgor are comparedbetween half-watered plants and fully watered controls.Typically, stomatal conductance, a measure of stomatalopening, declines within a few days of withholding waterfrom the roots (Figure 21.5B), yet there is no measurablechange in water potential or loss of turgor in the leaves.In experiments with day flower (Commelina communis),there was a significant increase in ABA content of theroots in the dry container and in the leaf epidermis(Figure 21.6). Furthermore, ABA is readily translocatedfrom roots to the leaves in the transpiration stream, evenwhen roots are exposed to dry air. These results suggestthat ABA is involved in some kind of early warningsystem that communicates information about soil waterpotential to the leaves.

21.1.6 OTHER ABSCISIC ACIDRESPONSES

There is recent evidence that ABA may also have arole in lateral or secondary root development. Theinitiation and development of lateral roots is known tobe primarily under the control of auxin, but lateral rootdevelopment can be inhibited by ABA if the hormone isapplied during early stages of lateral root development,before the lateral root meristem becomes organized.

Earlier studies also indicated an impact of exoge-nous ABA on flower formation under certain conditions,but the data was equivocal. In particular, no causal rela-tionship could be established between endogenous ABAlevels and flowering behavior. However, the prospect ofa role for ABA in flowering has been revived recentlywith the discovery that, under conditions that wouldnormally delay flowering, ABA-deficient mutants ofArabidopsis produce flowers somewhat earlier than wild-type plants. This observation suggests that endogenousABA may normally inhibit or delay flowering in Ara-bidopsis. Further support comes from the discovery thata gene (FCA) previously known to be involved in con-trolling the time of flowering also has the properties ofan abscisic acid receptor. We will take a closer look atthis receptor in the next section and the role of the FCAgene in flowering in Chapter 25.

21.1.7 ABA PERCEPTION AND SIGNALTRANSDUCTION

ABA perception and signaling appears to be particular-ly complex and, although its metabolism and physiology

360 Chapter 21 / Hormones IV: Abscisic Acid, Ethylene, and Brassinosteroids

Time (days)

B.

A.

0.0

0.1

0.2

6543210

0.3

7

FIGURE 21.5 (A) An experimental setup fortesting the effects of desiccated roots onABA synthesis and stomatal closure. Rootsof a single plant are divided equally betweentwo containers. Water supplied to one con-tainer maintains the leaves in a fully turgidstate while water is withheld from the sec-ond container. Withholding water from theroots leads to stomatal closure, even thoughthe leaves are not stressed. (B) Stomatal clo-sure in a split-root experiment. Maize (Zeamays) plants were grown as shown in (A). Con-trol plants (open circles) had both halves of theroot system well-watered. Water was withheldfrom half the roots of the experimental plants(closed circles) on day zero. Stomatal open-ing, measured as leaf conductance, declined inthe plants with water-stressed roots. (B fromBlackman, P. G., W. J. Davies. 1985. Journalof Experimental Botany 36:39–48. Reprintedby permission of The Company of Biologists,Ltd.)

have been studied for decades, the mechanism ofABA perception and its subsequent signal chain haveremained elusive. As noted earlier, ABA is a weak acid.As such it is likely to exist in both the protonatedand unprotonated forms in the relatively acidicenvironment of the apoplast. In the protonated state itmay diffuse across the plasma membrane and react withan intracellular receptor or, in the unprotonated form,it may remain outside the cell to be sensed by a site onthe plasma membrane. Indeed, experiments employingimpermeable ABA derivatives and/or microinjection ofABA into cells have indicated multiple ABA receptorsat multiple locations.

Over the last 20 years, methods that have normallybeen employed to identify hormone receptors have pro-ven relatively unsuccessful in the search for ABAreceptors. A more recent approach has made use ofantigen-antibody reactions with what are called anti-idiotypic antibodies. In this method, antibodies raisedagainst ABA are themselves used as antigens to raisea second group of antibodies—the anti-idiotypicantibodies—that have binding characteristics similarto ABA. Thus, any protein that binds with theanti-idiotypic antibodies could be a putative ABAreceptor. The anti-idiotypic antibodies were then usedto screen the proteins encoded by a complimentary

Percentage of original fresh weight

20

AB

A (

ng/1

00

mg

d.w

t)

100 90 80 70 65 60

40

60

80

100

120

60

80

100

RW

C (

%)

FIGURE 21.6 Effect of air drying on the ABA contentof Commelina communis root tips. Root tips were airdried to the relative water contents shown in the uppercurve. Lower curve shows the dramatic increase in ABAcontent as the fresh weight decreases. (From Zhang,J., W. J. Davies. 1987. Journal of Experimental Botany38:2015–2023. Reprinted by permission of The Com-pany of Biologists, Ltd.)

21.1 Abscisic Acid 361

DNA (cDNA) library for barley aleurone cells. Thisapproach led to the identification of ABAP1, a pro-tein that is located in the plasma membrane of barleyaleurone cells and that specifically and reversibly bindsABA in vitro.

Since the discovery of ABAP1, at least three otherputative ABA receptors have been identified. One isa chloroplast protein Magnesium Protoporphyrin-IXChetalase H subunit (CHLH, also known as ABAR).The second is a soluble, flowering-time control proteinFCA isolated from Arabidopsis. Based on similarity ofamino acid sequence, FCA is homologous to the bar-ley protein ABAP1. FCA interacts with another protein(FY) to regulate the processing of functional mRNA (seeChapter 25 for the role of FCA in flowering). The thirdputative receptor is a membrane-localized G-proteincoupled receptor (GPCR) identified as GCR2. Thesimple fact that these proteins bind ABA in vitro,however, does not prove they are true receptors. Itstill needs to be demonstrated that loss-of-functionor gain-of-function mutants alter ABA functions in apredictable manner.

The signal chain for ABA effects, both upstream anddownstream from the hormone, is a subject of intensivestudy. The apparently complex interactions betweenabiotic signals, receptors, second messengers, andABA-induced gene transcription—let alone crosstalkwith other signals—make it difficult to assemble adefinitive scheme. Still, a number of componentsare beginning to fall into place. Most of the recentprogress has been made through newly discovered

ABA

A- PA

ABA

ER

K+Ca2+

K+

in

Ca2+channel

Kinase

TPP

H+

ATP

Ca2+

Guard cell [Ca2+]in

PLDR

G

K

out

FIGURE 21.7 A simplified schematic illustrating the coordination of ion pumpsby ABA and Ca2+ during closure of stomatal guard cells. ABA is perceived by anunknown receptor (ABA R) that transmits the ABA signal to inward-rectifying cal-cium channels via a membrane-associated protein kinase. The kinase is antagonizedby a protein phosphatase (PP). ABA also stimulates the release of Ca2+ from internalstores such as the endoplasmic reticulum (ER), possibly mediated by phospholipidsignaling and/or G protein. The increased cytosolic Ca2+ concentration inhibits K+

inchannels and opens both K+

out and anion channels (A−). The result is a net loss ofions from the guard cell, followed by a loss of water and turgor, and closure of thestomatal pore.

ABA-insensitive gene mutations and can be summarizedin the following points.

1. There appears to be rapid turnover of ABA in bothstressed and unstressed plants, but the events thatsense abiotic stress and initiate ABA accumulationremain unknown.

2. Ca2+ appears to be an important part of the ABAsignal chain, especially in stomatal guard cells.Ca2+ mediates ABA-induced turgor adjustmentsby activating plasma membrane anion channels(Figure 21.7).

3. The promoter region of some genes contains asequence called the ABA response element (ABRE).Transcription factors known as ABA response ele-ment binding factors (ABFs) bind to this promoterregion to regulate the activity of many ABA-inducedgenes. These genes include putative protective pro-teins such as enzymes required for the synthesisof osmolytes or compatible solutes that help theplant adapt to water stress (Chapter 13), and tran-scription factors that in turn regulate other changesin gene expression

4. A number of ABA-insensitive (abi) mutants havebeen identified. At least three insensitive mutants,abi3, abi 4, and abi 5, impair only seed germinationand early seedling development. All three wildtypegenes (ABI3, 4, 5) encode transcription factors thatare expressed mainly in seeds, suggesting that therole of ABA in seeds requires gene transcription.

362 Chapter 21 / Hormones IV: Abscisic Acid, Ethylene, and Brassinosteroids

5. A number of ABA-activated protein kinases thatpositively regulate ABA responses have been identi-fied. In addition, ABI1 and ABI2 are protein phos-phatases that negatively regulate ABA responses. So,protein phosphorylation events are clearly impor-tant in ABA signaling.

It will no doubt take some time to sort out all ofthese components and those yet to be discovered andconstruct a clear model of the signaling chains for variousABA-mediated responses.

21.2 ETHYLENE

Ethylene is another class of hormones with a singlerepresentative. Ethylene is a simple gaseous hydrocar-bon with the chemical structure H2C CH2. Ethyleneis apparently not required for normal vegetative growth,although it can have a significant impact on the devel-opment of roots and shoots. Ethylene appears to besynthesized primarily in response to stress and maybe produced in large amounts by tissues undergoingsenescence or ripening. Ethylene is commonly used toenhance ripening in bananas and other fruits that arepicked green for shipment (Box 21.2). Ethylene is fre-quently produced when high concentrations of auxinsare supplied to plant tissues and many of the inhibitoryresponses to exogenously applied auxin appear to be dueto auxin-stimulated ethylene release rather than auxinitself.

Ethylene occurs in all plant organs—roots, stems,leaves, bulbs, tubers, fruits, seeds, and so on—althoughthe rate of production may vary depending on the stageof development. Ethylene production will also vary fromtissue to tissue within the organ, but is frequently locatedin peripheral tissues. In peach and avocado seeds, forexample, ethylene production appears to be localizedprimarily in the seed coats, while in tomato fruit andmung bean hypocotyls it originates from the epidermalregions. The off-gassing of ethylene by natural vegeta-tion is also a significant source of atmospheric volatileorganic carbon (VOC).

21.2.1 ETHYLENE IS SYNTHESIZEDFROM THE AMINO ACIDMETHIONINE

Despite the early discovery of ethylene, its knownimportance in plant development, and its relativelyuncomplicated chemistry, the pathway for ethylenebiosynthesis initially proved difficult to unravel. Thisis partly because there were a large number of potentialprecursors (sugars, organic acids, or peptides) known tobe present in plant tissues. In addition, until recently,the enzymes involved have proven too labile to isolateand study in vitro. Consequently, most of the work has

been carried out in vivo, with all the pitfalls inherent insuch experiments. Moreover, ethylene is a volatile gasand available analytical methods for its measurementwere simply too insensitive. It wasn’t until the early1960s that developments in gas chromatography madeit possible to analyze ethylene at physiologically activeconcentrations. With the advent of gas chromatography,the study of ethylene began to advance rapidly.

M. Lieberman and L. W. Mapson first demon-strated in 1964 that methionine was rapidly convertedto ethylene in a cell-free, nonenzymatic model sys-tem. In subsequent studies, Lieberman and coworkersconfirmed that plant tissues such as apple fruit con-verted [14C]-methionine to [14C]-ethylene and that theethylene was derived from the third and fourth car-bons of methionine. Little progress was made until1977 when D. Adams and F. Yang demonstrated thatS-adenosylmethionine (SAM) was an intermediate inthe conversion of methionine to ethylene by apple tis-sue. In 1979, Adams and Yang further demonstrated theaccumulation of 1-aminocyclopropane-1-carboxylicacid (ACC) in apple tissue fed [13C]-methionine underanaerobic conditions—conditions that inhibit the pro-duction of ethylene. However, upon reintroduction ofoxygen, the labeled ACC was rapidly converted to ethy-lene. ACC is a nonprotein amino acid that had beenisolated from ripe apples in 1957, but its relationshipto ethylene was not obvious at that time. These resultsestablished that ACC is an intermediate in the biosyn-thesis of ethylene.

The three-step pathway for ethylene biosynthesis inhigher plants is shown in Figure 21.8. In the first step, anadenosine group (i.e., adenine plus ribose) is donated tomethionine by a molecule of ATP, thus forming SAM.An ATP requirement is consistent with earlier evidencethat ethylene production is blocked by inhibitors ofoxidative phosphorylation, such as 2,4-dinitrophenol.Conversion of methionine to SAM is catalyzed bythe enzyme methionine adenosyltransferase or SAMsynthetase.

The cleavage of SAM to yield 5′-methylthio-adenosine (MTA) and ACC, mediated by the enzymeACC synthase, is the rate-limiting step. ACC synthasewas the first enzyme in the pathway to be studied indetail. The enzyme has been partially purified fromtomato and apple fruit but, because of its instability andlow abundance, progress toward its purification andcharacterization has been slow. More recently, genesfor ACC synthase have been isolated from zucchini(Cucurbita) fruit and tomato pericarp tissue. The clonedgenes direct the synthesis of active ACC synthase in thebacterium E. coli and yeast, making it possible to producethe enzyme in sufficient quantity for further study.The enzyme that catalyzes the oxidation of ACC toethylene, previously referred to as the ethylene-formingenzyme but now known as ACC oxidase, proved

21.2 Ethylene 363

H2C CH2

BOX 21.2THE DISCOVERYOF ETHYLENE

The effect of ethylene on plants was originally describedby Dimitry Nikolayevich Neljubow, a graduate studentin Russia in 1886, who found that abnormal growth ofdark-grown pea seedlings could be traced to ethyleneemanating from illuminating gas. Interest in ethylene asa plant growth factor, however, did not gain real momen-tum until it was found to have commercial implications.

Those whose business involves the shipping andstoring of fruit have long been aware that ripe and

rotting fruit could accelerate the ripening of other fruitstored nearby. For example, bananas picked in Cubaand shipped by boat often arrived in New York inan overripe and unmarketable condition. One of theearliest reports that these effects were due to a volatilesubstance given off by plant tissue was published in 1910by H. H. Cousins in an annual report of the JamaicanDepartment of Agriculture. He discovered that ripeoranges released a volatile product that would accelerateripening of bananas stored with them. A number ofsimilar reports appeared in the early 1930s, showingthat volatile emanations from apples caused epinastyin tomato seedlings and respiratory changes associatedwith the ripening process. In 1934, R. Gane providedindisputable evidence that the volatile substance wasethylene.

especially difficult to isolate in the active form. ACCoxidase was finally identified when a gene cloned fromripening tomato fruit (pTOM13) was linked to ethyleneproduction.

Another important aspect of ethylene biosynthe-sis is the limited amount of free methionine availablein plants. In order to sustain normal rates of ethy-lene production, the sulfur released during ethyleneformation must be recycled back to methionine. Thisis accomplished by what is commonly referred to asthe methionine cycle (Figure 21.8). This cycle is alsoknown as the Yang cycle, after S. F. Yang, who carriedout much of the pioneering work on ethylene biosyn-thesis. Double-labeling experiments have shown that

the CH3S-group is salvaged and recycled as a unit. Theremaining four carbon atoms of methionine are suppliedby the ribose moiety of the ATP used originally to formSAM. The amino group is provided by a transaminationreaction.

Ethylene production is promoted by a numberof factors including IAA, wounding, and water stress,principally by the induction of the synthesis of ACCsynthase. Induction of this enzyme in plant tissues isblocked by inhibitors of both protein and RNA syn-thesis, suggesting that induction probably occurs at thetranscriptional level. In E. coli carrying the cloned ACCsynthase gene, the physical abundance of ACC syn-thase messenger RNA also increases in response to IAA

C

Methionine

I II III

EthyleneS-Adenosylmethionine(SAM)

I-Amino-cyclopropane-I-carboxylic acid

(ACC)

CH3

S

CH2

CH2

CO2 O2

CH3 − S − Ribose(MTR)

CH3 − S − Adenosine(MTA)

N − Malonyl − ACC(MACC)

COO− COO−

CH3

+ S − Adenosine

CH2

CH2

CH − NH3+CH − NH3

+

COO−H2C

H2C NH3+

COO−H2C

H2C NH − CO − CH2 − COO−

H2C CH2

Adenine

ATP

EFE

PPi + Pi

C

Yang cycle

FIGURE 21.8 A scheme for ethylene biosynthesis in higher plants. The enzymes areI: SAM synthetase; II: ACC synthase; and III: ACC oxidase. The ethylene group ishighlighted in yellow. The Yang cycle for sulfur recovery is highlighted in orange.

364 Chapter 21 / Hormones IV: Abscisic Acid, Ethylene, and Brassinosteroids

and wounding. Control of ethylene production thusappears to be exercised primarily through transcrip-tional regulation of the ACC synthase gene. Ethyleneproduction is also stimulated by ethylene itself, a form ofautocatalysis. This is commonly seen in ripening fruits(see Chapter 25) where ethylene apparently stimulatesan increase in both ACC synthesis and its subsequentconversion to ethylene.

21.2.2 EXCESS ETHYLENE IS SUBJECTTO OXIDATION

Unlike other hormones, ethylene is a volatile gas thatis readily given off to the atmosphere. Ethylene can,however, be metabolized by oxidation to carbon dioxideor by conversion to either ethylene oxide or ethyleneglycol. It has not yet been established whether ethylenemetabolism has any active role in the physiologicalaction of the hormone. In fact, kinetic studies haveindicated that ethylene metabolism is a straightforwardchemical reaction not subject to normal physiologicalcontrols. It may thus be only a nonessential consequenceof high ethylene levels in the tissue. It is therefore likelythat most tissues lose excess ethylene by simple diffusioninto the surrounding atmosphere.

21.2.3 THE STUDY OF ETHYLENEPRESENTS A UNIQUE SETOF PROBLEMS

Because ethylene is a simple gaseous hydrocarbon thatreadily diffuses from its site of synthesis, study of itsrole in development presents a unique set of prob-lems. Although known primarily for its effects on fruitripening and its synthesis by many tissues in responseto stress, ethylene is known to affect virtually everyaspect of plant growth and development. As a byproductof hydrocarbon combustion, ethylene is also a com-mon environmental pollutant that can play havoc withgreenhouse cultures or laboratory experiments. In addi-tion, ethylene biosynthesis is also stimulated by highlevels of auxin and other hormones. Still, our under-standing of ethylene physiology has made tremendousstrides over the past several decades, owing largelyto development of the gas chromatograph, the avail-ability of ethylene-releasing agents, and the study ofethylene-insensitive mutants. The gas chromatographhas made possible quantitative analysis of ethylene atextremely low concentrations that could not other-wise be measured. Ethephon (2-chloroethylphosphonicacid) is a compound that, at physiological pH, readilydecomposes to produce ethylene. Use of ethephon isadvantageous in the laboratory as its application andconcentration is often more easily controlled comparedwith gaseous ethylene.

21.2.4 ETHYLENE AFFECTS MANYASPECTS OF VEGETATIVEDEVELOPMENT

Ethylene is known primarily for its promotion of fruitripening and senescence. Ethylene control of fruit devel-opment has been studied extensively in tomato, whichis a climacteric fruit. During the development of cli-macteric fruits there is a characteristic developmentallyregulated burst in respiration called the climacteric rise.The climacteric rise is normally accompanied by ethy-lene production and is followed by the expression ofa set of genes that enhance ripening-related activitiessuch as development of fruit color, flavor, and texture.The tomato never ripe mutant is insensitive to ethylenebecause it has a defective ethylene receptor protein.Consequently, the ‘‘ripening genes’’ are not expressedand, although the fruit reaches full size, it never developsthe red color, flavor, and texture characteristic of a ripetomato.

Ethylene has been shown to stimulate elongationof stems, petioles, roots, and floral structures of aquaticand semiaquatic plants. The effect is particularly notedin aquatic plants because submergence reduces gas dis-persion and thus maintains higher internal ethylenelevels. In rice, ethylene is ineffective in the presence ofsaturating levels of gibberellins, which also promotesstem elongation. Moreover, ethylene promotes gib-berellin synthesis in rice and the elongation effect canbe blocked with antigibberellins, which suggests thatgibberellin mediates this ethylene effect. By contrast,root and shoot elongation in nonaquatic plants such aspeas (Pisum sativum) is inhibited by ethylene.

Ethylene stimulates many inhibitory and abnormalgrowth responses such as the swelling of stem tissuesand the downward curvature of leaves, or epinasty. Leafepinasty occurs because of excessive cell elongation onthe adaxial (i.e., upper) side of the petiole. Epinasty isa common response to water logging of flood-sensitiveplants such as tomato (Lycopersicum) and is actually aresponse to anoxia in the region of the roots. Themore vertical orientation of epinastic leaves reduces theabsorption of solar energy and, consequently, transpi-rational water loss. This helps to bring water loss moreinto line with reduced capacity for water uptake in plantssuffering root anoxia.

A role for ethylene has also been noted for pro-motion of seed germination, inhibition of bud break,reduced apical dominance, fruit ripening, cell death,and pathogen responses. Ethylene can be a problem incommercial greenhouse that are heated with gas-firedheating systems.

21.2.5 ETHYLENE RECEPTORSAND SIGNALING

One of the best-known effects of ethylene is referred toas the ‘‘triple response’’ of etiolated (dark-grown) dicot

21.2 Ethylene 365

seedlings (Figure 21.9). This response is characterizedby inhibition of hypocotyl and root cell elongation, apronounced radial swelling of the hypocotyl, and exag-gerated curvature of the plumular hook. The responseis rapid (3 days post-germination) and allows largepopulations of seedlings to be screened for ethylenemutations. An absence of the triple response in the pres-ence of exogenous ethylene has been used successfully toidentify ethylene-resistant mutants and other ethyleneresponse defects, especially in experiments conductedwith Arabidopsis. These mutants generally fall into one ofthree distinct categories: (1) constitutive triple-responsemutants that exhibit the triple response in the absenceof ethylene, (2) ethylene-insensitive mutants, and (3)mutants in which ethylene-insensitivity is limited tospecific tissues, such as the plumular hook or root elon-gation. As a result of these experiments, several ethylenereceptors and downstream elements in the ethylenesignal chain have been identified.

Ethylene is perceived by a family of five membrane-associated, two-component histidine kinase receptors.Unlike most other two-component receptors, whichare localized in the plasma membrane, it has beenshown convincingly that the Arabidopsis receptor ETR1is associated with the membrane of the endoplasmicreticulum (ER). The specific advantage(s) to localiza-tion of the receptor in the ER rather than the plasmamembrane is not clear, but since ethylene diffuses read-ily in both aqueous and lipid environments, ethylenewould have ready access to any subcellular location.

Plumular hook

Hypocotyl

Cotyleadons

Root

Air Ethylene

FIGURE 21.9 Diagramatic illustration of the ethylenetriple response in dark-grown dicot seedlings. Note theinhibition of hypocotyl and root cell elongation, a pro-nounced radial swelling of the hypocotyl, and exagger-ated curvature of the plumular hook in the seedlingsexposed to ethylene. (Based on Guzman, Ecker. 1990.Plant Cell 2:513–523.)

The sensor domain of the ethylene receptor has threemembrane-spanning regions and is assumed to functionas a dimer. The sensor domain also contains a coppercofactor that is necessary for ethylene binding. A uniquecharacteristic of the ethylene response system is that thereceptors are believed to be constitutively active. Thismeans that the receptors and subsequent signal chainare functional in the absence of ethylene and that ethylene,in effect, turns the system off.

Although there are some variations, ethylenesignaling appears to follow the general model shownin Figure 21.10. In the absence of ethylene, the signalchain begins with a protein called Constitutive Triple

+ETHYLENEAIR

Cu Cu

ETR1

CTR1

ProteionKinaseCascade

ATP

NATP

ADP

NADP

PTF

P

Promoter Gene

TF

Nucleus

CTR1

ETR1

FIGURE 21.10 A model for gene regulation by the ethy-lene perception and response pathway. Ethylene issensed by a family of two-component histidine kinasereceptors (ETR) that are located in the membrane ofthe endoplasmic reticulum. In the absence of ethy-lene the receptors are functionally active and activatea serine-threonine kinase (CTR1). CTR1 is the firstcomponent in a protein kinase cascade that ultimatelytargets one or more transcription factors. Phosphoryla-tion activates the transcription factors that are then ableto bind to the promoter regions of ethylene-sensitivegenes and enable transcription of the genes. Ethylenebinding inhibits receptor function and blocks the acti-vation of CTR1, thus shutting down the protein kinasecascade, preventing phosphorylation of the transcriptionfactor, and turning off the gene.

366 Chapter 21 / Hormones IV: Abscisic Acid, Ethylene, and Brassinosteroids

Response 1 (CTR1). CTR1 physically interacts withthe histidine kinase domain of the receptor ETR1.This interaction leads to the phosphorylation of CTR1and initiates the signal transduction stream. CTR1 is aserine/threonine protein kinase. According to thismodel, CTR1 initiates a protein kinase cascade thatultimately results in the phosphorylation of one or more

transcription factors and the constitutive expression ofcertain genes. The protein kinase cascade is very similarto a widely known group of mitogen-activated proteinkinases that serve a critical role in the transduction ofmany signals in animals, plants, and fungi (Box 21.3).CTR1 is equivalent to a MAPKKK and several potentialcandidates for the other kinases in the MAPK cascade

MAPKKK

MAPKK

MAPK

BOX 21.3MITOGEN-ACTIVATEDPROTEIN KINASE:A WIDESPREADMECHANISMFOR SIGNALTRANSDUCTION

The capacity of extracellular signals such as hormones,light, osmotic status, and stress to effect a change inthe physiology of cells often depends on regulating thetranscription of genes. The expression of a gene inturn depends on the binding of activated transcriptionfactors to the DNA in the promoter region of thegene. One of the primary means for regulating thisinteraction between a transcription factor and DNAis phosphorylation. Some transcription factors will notbind to DNA unless they are phosphorylated, whilephosphorylation inhibits the binding of others.

Many extracellular signals are linked to transcrip-tion factor phosphorylation by the mitogen-activatedprotein kinase (MAPK) system (Figure 21.11). Origi-nally named for its role in activation of genes involved incell proliferation (mitogen = to stimulate mitosis), theMAPK system is known to mediate gene transcriptionin response to a variety of signals in animals, plants, andfungi.

The core of the MAPK system is a sequence ofthree kinase enzymes. Each enzyme acts to effect thephosphorylation of the next enzyme in the sequence.Starting at the bottom end, the third enzyme in thesequence, MAP kinase (MAPK), is responsible for phos-phorylating a transcription factor. MAPK is in turnphosphorylated by an MAP kinase kinase (MAPKK orMAP2K), which is phosphorylated by an MAP kinasekinase kinase (MAPKKK, or MAP3K). MAPKKK canbe activated directly by a signal-receptor complex, aG-protein, or some other secondary signal.

There are two advantages to this system. The first isthat each kinase is able to phosphorylate multiple copies

of the next component—hence the name ‘‘cascade.’’The second advantage is that each of the componentsis represented by a small family of proteins. By utilizingdifferent members of each family in various combi-nations, the cell is able to assemble a large number ofdifferent pathways that can interpret different extracellu-lar signals and activate a variety of different transcriptionfactors.

Stimulus

Receptor

G-protein

MAPKKK

MAPKK

MAPK

Transcription factor

MKP1 Gene Gene products

MAPKmodule

FIGURE 21.11 The principal components of a generalizedMAP kinase cascade. The activation of a receptor by anextracellular stimulus initiates the kinase cascade involv-ing a sequence of serine-threonine kinases. The initialactivation may be mediated by a G-protein or other sec-ond messenger system. The first kinase in the cascade,mitogen-activated protein kinase kinase kinase (MAP-KKK) phosphorylates the second kinase, MAP kinasekinase (MAPKK), which in turn phosphorylates the thirdcomponent, MAP kinase (MAPK). The ultimate target isthe phosphorylation of a transcription factor. The acti-vated transcription factor binds to the promoter regionof the target gene and up-regulates transcription of thegene. One of the genes whose expression is stimulated isMKP1, a protein phosphatase that can remove the phos-phate group from MAPK and block gene activation.

21.3 Brassinosteroids 367

stream have recently been identified in Arabidopsis.When ethylene binds with the receptor, it preventsthe interaction of CTR1 with ETR1. This blocks theinitiation of the protein kinase cascade and subsequentgene activation. The result is that in the absence ofethylene the expression of ethylene-controlled genes isalways ‘‘on.’’ The effect of ethylene is to turn thesegenes ‘‘off ’’ by preventing the activation of the requiredtranscription factors.

21.3 BRASSINOSTEROIDS

Brassinosteroids are steroid hormones with a chemicalstructure similar to the steroid hormones in animals.Brassinosteroids elicit an impressive array of develop-mental responses, including an increased rate of stemand pollen tube elongation, increased rates of cell divi-sion (in the presence of auxin and cytokinin), seedgermination, leaf morphogenesis, apical dominance,inhibition of root elongation, vascular differentiation,accelerated senescence, and cell death. Brassinosteroidsare also implicated in mediating responses to bothabiotic and biotic stress, including salt, drought, tem-perature extremes, and pathogens.

The study of brassinosteroids as plant hormonesdates back to the early 1970s, when a group of agricul-tural researchers began screening pollen, already knownas a rich source of growth-promoting substances. Theresult was a complex mixture of lipids that stimulatedelongation of bean second internodes. Because the mostactive preparations were isolated from pollen of the rapeplant (Brassica napus), the active substances were referredto collectively as brassins.

Many of the effects of the brassins were similar tothose of GA, leading many to believe the extracts weresimply crude extracts of gibberellins, rather than a newclass of hormones as originally proposed. However, in1979, M. D. Grove and his coworkers identified theactive component as brassinolide (BL) (Figure 21.12).The very low concentration of brassinosteroids in planttissues is suggested by the fact that approximately 40kilograms of bee-collected rape pollen were required inorder to obtain only 4 milligrams of pure, crystallineBL! Fortunately, BL was chemically synthesized onlytwo years later, eliminating the need for such massiveand laborious extraction procedures.

For many years, the classification of brassinosteroidsas hormones was not widely accepted. Effects couldbe demonstrated only by exogenous application and itwas difficult to assess their function in vivo. This allchanged with the discovery of mutants of Arabidopsis,pea, and tomato that are blocked in brassinosteroidbiosynthesis. In all cases, the normal phenotype couldbe rescued by the application of brassinosteroids. These

HO

O

Brassinolide

HO

OH

OH

O

CH3

CH3

CH3

H3CH3C

H3C

FIGURE 21.12 The phytosterol brassinolide (BL) is anexample of brassinosteroid hormones.

studies clearly established that brassinosteroids havesome well-defined functions in normal plant develop-ment and qualify as a distinct class of endogenous planthormones. More than 40 analogs of BL have now beenisolated from more than 60 plant species and virtuallyall types of tissues, including pollen, seeds, leaves, stems,roots, and flowers. However, brassinolide remains themost biologically active brassinosteroid and is widelydistributed throughout the plant kingdom. Like otherhormones, brassinolide is active in micromolar concen-trations. Brassinolide at a concentration of 10−7 M isable to stimulate a fourfold increase in the length ofsoybean epicotyl sections.

Interestingly, most of the responses controlled bybrassinosteroids are also controlled by auxin. Althoughthe two hormones can act independently, it should notbe too surprising that there is considerable crosstalkbetween brassinosteroid and auxin signaling pathways.Relatively little is known about the mechanisms forBR regulation of the many responses it appears to beinvolved in. With respect to cell elongation, however,the evidence indicated that BR increases plastic exten-sibility of the cell wall by regulating genes encodingwall-modifying enzymes such as expansins and cellulosesynthase.

21.3.1 BRASSINOSTEROIDSARE POLYHYDROXYLATEDSTEROLS DERIVED FROMTHE TRITERPENE SQUALENE

Brassinosteroids are polyhydroxylated plant sterols—lipoidal substances related biosynthetically to the gib-berellins and abscisic acid (refer to Figure 19.4). Plantssynthesize a large number and variety of sterols, includ-ing sitosterol, stigmasterol, cholesterol, and campes-terol. Sterols and sterol derivatives are discussed moreextensively in Chapter 27. Sterols are triterpenoids, C30molecules that are derived from acetate through themevalonic acid pathway (see Figure 19.3A, 19.4). In

368 Chapter 21 / Hormones IV: Abscisic Acid, Ethylene, and Brassinosteroids

the synthesis of terpenes, sequential additions of the5-carbon isopentenyl pyrophosphate (IPP) produce ter-penes with 10-, 15-, or 20-carbon atoms. Triterpenesare formed when two C15 (farnesyl) units join headto head to form the C30 molecule squalene. The sub-sequent biosynthesis of plant sterols is not yet fullyunderstood, but the first step is a cyclization reaction toform cycloartenol (Figure 21.13). Using cycloartenolas a common precursor, there are probably multiplepathways leading to the several sterols found in plants.Decarboxylation and oxidation reactions are involved,as most common sterols have from 26 to 29 carbons anda single hydroxyl (—OH) group.

It is thought that most sterols, with the exceptionof stigmasterol, may serve as precursors for variousbrassinosteroids. However, the pathway for the biosyn-thesis of brassinolide is best understood. This pathwaywas established largely through studies using culturedcells of Catharanthus roseus. The precursor to brassi-nolide is campesterol, a C28 sterol. Through a seriesof largely oxidative steps, additional hydroxyl (—OH)groups are added and oxygen is introduced into theB ring. There are two parallel pathways for the con-version of campesterol to brassinolide, depending onwhether the oxidation at carbon-6 occurs early or late inthe pathway. In either case, approximately 12 steps are

HO

Squalene

Cycloartenol

A B

C D

HO

OH

OH

HOH O

O

HO

2523 24

28

2220

1817

11

19

1

45

6

7

2

3

10 8

9

12

13

14 15

16

2126

27

Campesterol

BrassinolideFIGURE 21.13 Principal steps in the biosynthesis of brassinolide from the triter-penoid squalene. The biosynthesis of sterols in plants is poorly understood. Squaleneundergoes a cyclization reaction to form cycloartenol. Cycloartenol is then subjectto various oxidations and methylations to form campesterol and other sterols. Thepathway for synthesis of brassinolide from campesterol has been established in cul-tured cells of Catharanthus roseus. Two alternate pathways are known, each involvingat least 12 steps.

21.3 Brassinosteroids 369

required to complete the conversion of campesterol tobrassinolide (Figure 21.13).

21.3.2 SEVERAL ROUTESFOR DEACTIVATIONOF BRASSINOSTEROIDSHAVE BEEN IDENTIFIED

Studies into the metabolism of brassinosteroids intomato and Ornithopus cells, cucumber, and mung beanhave revealed several deactivation mechanisms. Theα-hydroxyl groups on the A ring may be epimerizedto form a β-hydroxyl, which is then followed byesterification with fatty acids or by glucosylation.Other possibilities include cleavage of the side chain orconjugation at other hydroxyl positions. The precisemethod for deactivation clearly depends on the speciesand possibly the tissue involved.

21.3.3 BRASSINOLIDE RECEPTORSAND SIGNALING

With the discovery of brassinosteroids in plants, it is nowclear that steroid hormones are not restricted to animals.Unlike animals, however, which rely mainly on nuclearreceptors for steroid hormones, plants appear to usemembrane-based receptors to initiate a phosphorylationcascade that carries the signal into the nucleus. Althoughgenetic studies are beginning to unmask some of themany components in brassinosteroid perception andsignaling, there remains much to be learned about howthey interact with each other and with signaling forother biotic and abiotic factors. Figure 21.14 depicts ageneral model for brassinosteroid signaling.

The principal receptor for brassinosteroids requiresthe interaction of two proteins that form a plasmamembrane-associated heterodimer. The first is aserine/threonine kinase known as BRASSINOS-TEROID INSENSITIVE 1 (BRI1). (Keep in mindthat the name of the gene and protein are based onobserved mutations—hence the name ‘‘insensitive.’’)The second is the protein BRI1-ASSOCIATEDRECEPTOR KINASE (BAK1). Brassinosteroids bindto the extracellular domain of BRI1, which first inducesthe dissociation of an inhibitory protein (BKI1) thatinhibits the association of BAK1 with BRI1 and thenpromotes dimerization with BAK1 and autophos-phorylation of BRI1. The phosphorylated complexinitiates BR signaling. One target of brassinosteroidsignaling is the protein BZR1 (BRASSINAZOLERESISTANT 1). BZR1 is a transcription factorwhose location is dependent on its phosphorylationstatus. In the phosphorylated state, BZR1 is trappedin the cytoplasm while dephosphorylation allowsit to move into the nucleus. The phosphorylation

status of BZR1 is mediated by two competing factors;the BR signaling chain and a separate protein,BIN2. BIN2 mediates phosphorylation of BZR1and thus keeps the protein in the cytoplasm. Thebrassinosteroid signal, on the other hand, mediatesdephosphorylation of BZR1, which both activates thetranscription factor and encourages its migration intothe nucleus. Once in the nucleus, BZR1-P binds totarget sites in the promoter region of BR-sensitivegenes and initiates transcription. BR signaling mayalso inhibit the phosphorylating ability of BIN2, thusfurther ensuring activation and nuclear localization ofBZR1.

BRI1BAK1

BKI1

BZR1

BZR1

BIN2

?

P

P

BR Signaling

BR

Cytoplasm

Nucleus

Transcription

FIGURE 21.14 A proposed scheme for brassinosteroid(BR) signaling. In the absence of BR, the two receptorkinases BRI1 and BAK1 exist independently in the mem-brane as monomers. Dimerization, necessary for activa-tion, is inhibited by the presence of BKI1. BR inducesthe removal of BKI1, followed by the dimerization of thetwo receptors and auto phosphorylation of kinase regionof BRI1. The targets of the BR signaling chain are tran-scription factors such as BZR1. The cellular location andactivaty of BR-sensitive transcription factors is deter-mined by their phosphorylation status. The BR signalingchain dephosphorylates cytoplasmic BZR1-P, which thenmigrates into the nucleus where it enables transcriptionof BR-sensitive genes. The protein BIN2 phosphorylatesBZR1, causing it to migrate back into the cytoplasm, thusinhibiting transcription. BR signaling may also enhancetranscription by blocking BIN2.

370 Chapter 21 / Hormones IV: Abscisic Acid, Ethylene, and Brassinosteroids

SUMMARY

Abscisic acid (ABA) is a 15-carbon sesquiterpene thatis synthesized by cleavage of the 40-carbon caroteneviolaxanthin. It is synthesized primarily in green leavesand can be stored in the chloroplast, although there issome evidence that ABA may also be either synthesizedor stored in roots and exported to the leaves in times ofwater stress.

The primary functions of ABA are (1) prohibitingprecocious germination and promoting dormancy inseeds and (2) inducing stomatal closure and the produc-tion of molecules that protect cells against desiccationin times of water stress. ABA has also been implicatedin other developmental responses, including hetero-phylly (leaves of different shape on the same plant),initiation of secondary roots, flowering, and senes-cence.

ABA perception and signaling appears to be partic-ularly complex. Several putative receptors or ABA bind-ing proteins have been identified, including ABAP1 inbarley aleurone tissue, a protein involved in flowering,and a G-protein coupled receptor. A number of othercomponents of various ABA signaling chains have beenidentified, but a clear model for any one chain has yetto be constructed.

Ethylene is a simple 2-carbon gaseous hydro-carbon that appears to be synthesized primarily inresponse to stress or in senescing and ripening tis-sue. Ethylene is ubiquitous in plants, has been impli-cated in a wide range of developmental responses, andfrequently interacts with auxin and gibberellin. It’sbest-known response is the triple response of etiolateddicot seedling, characterized by inhibition of hypocotyland root cell elongation, a pronounced radial swellingof the hypocotyl, and exaggerated curvature of theplumular or epicotyl hook.

Ethylene is synthesized by a simple three-step pro-cess from the sulfur-containing amino acid methionine.In order to sustain normal rates of ethylene produc-tion, the sulphur released during ethylene formation isrecycled back to methionine via the Yang Cycle.

Ethylene is sensed by a membrane-locatedhistidine kinase receptor that initiates a mitogen-activated protein (MAP) kinase cascade that results inthe activation of transcription factors. Uniquely in thecase of ethylene, however, the receptor kinase systemis constitutively active (i.e., ‘‘on’’) and is blocked, orturned ‘‘off,’’ by ethylene.

Brassinosteroids (BR) are steroidal hormones thatelicit a wide range of effects when applied to plantsincluding an increased rate of stem and pollen tube

elongation, increased rates of cell division (in the pres-ence of auxin and cytokinin), seed germination, leafmorphogenesis, apical dominance, inhibition of rootelongation, vascular differentiation, accelerated senes-cence, and cell death. Brassinosteroid sensing involvesa serine/threonine kinase that regulates the phos-phorylation and dephosphorylation of transcriptionfactors.

CHAPTER REVIEW

1. Auxins, gibberellins, and brassinosteroids allinfluence stem elongation. In what ways do theresponses of stems to these three hormones differ?

2. What is the evidence that ABA mediates responsesto water stress?

3. What unique problems are related to thestudy of ethylene as a plant hormone?

4. Describe the hormonal changes that occur duringseed development, maturation, and germination.Offer a rationale for the observed pattern.

5. What is a ‘‘MAPK module’’? How is it involvedin signal perception and transduction?

6. How are gibberellins and brassinosteroids relatedbiosynthetically?

7. What is the evidence that ABA is synthesizedin leaf mesophyll cells? In vascular tissue?

8. What is vivipary? How do we know thatABA is involved in regulating vivipary?

9. Compare and contrast the receptor systemsfor ABA, ethylene, and brassinosteroids.

FURTHER READING

Buchanan, B. B., W. Gruissem, R. L. Jones. 2000.Biochemistry and Molecular Biology of Plants. Rockville,MD: American Society of Plant Physiologists.

Chen, Y-F., M. D. Randlett, J. L. Findell, G. E. Schaller.2002. Localization of the ethylene receptor ETR1 tothe endoplasmic reticulum. Journal of Biological Chemistry277:19861–19866.

Clouse, S. D. 2002. Brassinosteroids. In: C. R. Somerville,E. M. Meyerowitz (eds.), The Arabidopsis Book. AmericanSociety of Plant Biologists, Rockville, MD, doi: 10.1199/tab.0009, www.aspb.org/publications/arabidopsis/.

Davies, P. J. 2004. Plant Hormones: Biosynthesis, Signal Trans-duction, Action. Dordrecht: Kluwer Academic Publishers.

Further Reading 371

Gendron, J. M., Z-Y Wang. 2007. Multiple mechanismsmodulate brassinosteroid signaling. Current Opinion inPlant Biology 10:436–441.

Guo, H., J. R. Ecker. 2004. The ethylene signaling pathway:New insights. Current Opinion in Plant Biology 7:40–49.

Guzman, P., J. R. Ecker. 1990. Exploiting the triple responseof Arabidopsis to identify ethylene-related mutants. PlantCell 2:513–523.

Hardtke, C. S. 2007. Transcriptional auxin-brassinosteroidcrosstalk: Who’s talking? BioEssays 29:1115–1123.

Hirayama, T., K. Shinozaki. 2007. Perception and transduc-tion of abscisic acid signals: Keys to the function of theversatile plant hormone, ABA. Trends in Plant Science12:343–351.

Karp, G. 2008. Cell and Molecular Biology. New York: JohnWiley & Sons. (Includes a good review of MAP kinases.)

Li, J. 2005. Brassinosteroid signaling: From receptor kinases

to transcription factors. Current Opinion in Biology8:526–531.

Nambra, E., A. Marion-Poll. 2005. Abscisic acid biosyn-thesis and catabolism. Annual Review of Plant Biology56:165–185.

Razem, F. A. et al. 2004. Purification and characterization ofa barley aleurone abscisic acid-binding protein. Journalof Biological Chemistry 279:9922–9929.

Ryu, H. et al. 2007. Nucleocytoplasmic shuttling of BZR1mediated by phosphorylation is essential in Arabidopsisbrassinosteroid signaling. The Plant Cell 19:2749–2762.

Schwartz, S. H., X. Qin, J. A. D. Zeevaart. 2003. Elucida-tion of the indirect pathway of abscisic acid biosynthe-sis by mutants, genes, enzymes. Plant Physiology. 2003.131:1591–1601.

Verslues, P., J-K. Zhu. 2007. New developments in abscisicacid perception and transduction. Current Opinion inPlant Biology 10:447–452.

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Red light

Response

Far-red light

Pr Pfr

22Photomorphogenesis: Responding to Light

Plants do not enjoy the luxury of being able to changetheir environment or to seek shelter from adverse con-ditions by changing their location. Yet species survivaldictates that plants must be able to avoid adverse con-ditions. The germination of seeds and survival of theseedling that emerges are largely dictated by conditionsin their immediate environment. Many seeds will notgerminate if buried too deeply or if they lay in theshade of a forest canopy. Seedlings that emerge beneatha canopy tend to have elongated stems, as if reachingout for the light. Weeds growing in full sun at theedge of a wheat field will be shorter and more com-pact than plants of the same species competing with thecrop in the center of the field. Plants flower at differenttimes, spreading flowering through the season as thougheach species is awaiting some environmental cue. Theseand similar patterns of plant behavior have a signifi-cant survival advantage. They enable plants to make thebest use of available resources, compete effectively withother species, or anticipate unfavorable environmentalchange.

How can seeds and seedlings know where theyare? How can plants ensure they are in a position tomaximize photosynthesis? How do plants measure thepassing of the season? The answers to these and manyother questions directly related to plant survival may befound in their capacity to detect and interpret a varietyof environmental signals.

One particularly important environmental signal islight. The quantity and quality of light are constantlychanging, often in predictable ways. Plants are able tomonitor these changes and make use of this informationto direct their growth, form, and reproduction. In thischapter we

• introduce the concept of light-regulated plantdevelopment or photomorphogenesis,

• describe the phytochromes; pigments that mediateplant responses to red and far-red light,

• describe the cryptochromes; pigments that mediateplant responses to blue light,

• review the principal developmental responses regu-lated by the phytochromes and cryptochromes,

• review the basic aspects of phytochrome and cryp-tochrome signal transduction, and

• present a case study of the role of phytochrome andcryptochrome in de-etiolation in Arabidopsis.

22.1 PHOTOMORPHOGENESISIS INITIATED BYPHOTORECEPTORS

The regulation of plant development by light, orphotomorphogenesis, is a central theme in plant

373

374 Chapter 22 / Photomorphogenesis: Responding to Light

development. In order to acquire and interpret theinformation that is provided by light, plants havedeveloped sophisticated photosensory systems com-prised of light-sensitive photoreceptors and signaltransduction pathways. A photoreceptor ‘‘reads’’ theinformation contained in the light by selectivelyabsorbing different wavelengths of light. Absorptionof light normally induces a conformational change inthe pigment or an associated protein, a photochemicaloxidation-reduction reaction, or some other form ofphotochemical change. Whatever the nature of theprimary event, absorption of light by the photoreceptorsets into motion a cascade of events that ultimatelyresults in a developmental response.

There are four classes of photoreceptors in plants.The phytochromes absorb red (R) and far-red (FR)light (ca. 660 and 735 nm, respectively) and have arole in virtually every stage of development from seedgermination to flowering. Cryptochromes and pho-totropin detect both blue (400–450 nm) and UV-Alight (320–400 nm). The cryptochromes appear to playmajor roles during seedling development, flowering,and resetting the biological clock. Phototropin mediatesphototropic responses, or differential growth in a lightgradient, and will be discussed in the context of theseresponses in Chapter 23. A fourth class of photore-ceptors that mediate responses to low levels of UV-B(280–320 nm) light have not yet been characterized.

Phytochrome, cryptochrome, and phototropin areall chromoproteins (Chapter 6). Chromoproteins con-tain a light-absorbing group, or chromophore, attachedto a protein with catalytic properties, called the apopro-tein. The chromophore plus the apoprotein is referredto as the holoprotein.

22.2 PHYTOCHROMES:RESPONDING TO REDAND FAR-RED LIGHT

Phytochrome is a unique pigment that can exist in twostates—one with an absorption maximum in the red(R, or 665 nm) region of the spectrum and one withan absorption maximum in the far-red (FR, 730 nm)(Figure 22.1). The pigment is ubiquitous in plantsand its discovery solely on the basis of simple butelegant physiological experiments ranks among themajor achievements of twentieth-century plant biology(Box 22.1).

Early phytochrome studies employed almost exclu-sively a combination of physiological experiments andphysical techniques such as in vivo spectroscopy. It isinteresting to note that even these early physiologicalexperiments predicted multiple forms of phytochrome.

300 400 500 600 700 800Wavelength (nm)

Abs

orba

nce

Pfr

Pr

FIGURE 22.1 Absorption spectra of the Pr and Pfrforms of purified phytochrome. The spectrum for Pfris actually the spectrum for an equilibrium mixtureof Pfr and Pr (see text for details). Note the differen-tial absorption in the blue region of the spectrum aswell as the red, far-red region. Some blue light effectsare mediated by phytochrome, but photoconversion byred light is 50 to 100 times more effective than in theblue. Because both forms absorb equally in the greenregion (500 to 550 nm), green light does not appreciablychange the state of the pigment and can, in most cases, beused as a safe light when setting up phytochrome experi-ments.

With the advent of molecular genetics, the existenceof multiple phytochromes in higher plants was con-firmed. In most angiosperms, for example, there areat least three distinct phytochromes: phyA, phyB, andphyC, encoded by the genes PHYA, PHYB, and PHYC,respectively.1 Arabidopsis, which has been studied mostextensively, has five phytochromes (phyA–phyE). Thedifferences between the several phytochromes are in theprotein—the chromophore is common to all membersof the phytochrome family. The different phytochromesare expressed in different tissues at different times indevelopment and mediate different light responses,although it appears that phyA and phyB are the prin-cipal species that mediate red and far-red responses.Phytochromes also interact with each other as wellas with other photoreceptors and developmental stim-uli, rendering an understanding of their complex signaltransduction pathways one of the more challenging areasof current study.

1The notation convention adopted for plant photoreceptorsfollows the recommendations of Quail et al. 1994. Plant Cell6:468: e.g., phyA for the holoprotein, PHYA for theapoprotein, PHYA for the wildtype gene, and phyA for themutant gene.

22.2 Phytochromes: Responding to Red and Far-Red Light 375

BOX 22.1HISTORICALPERSPECTIVES—THE DISCOVERYOF PHYTOCHROME

It is now well established that phytochrome plays a crit-ical role in almost every stage of plant development. Itsexistence was predicated on the basis of a simple phys-iological observation: seed germination and growth ofetiolated seedlings exhibited photoreversible responsesto red and far-red light. Because of its uniquely pho-toreversible character, however, the newly proposedpigment was initially greeted with skepticism in thescientific community.

It had long been recognized that light influencedplant development under conditions that excluded sig-nificant levels of photosynthesis. Indeed, dramatic dif-ferences in the growth and form of plants in darknessand light had fascinated botanists and physiologists forcenturies. However, little real progress toward under-standing these phenomena was achieved until the early1950s. At that time, H. A. Borthwick, a botanist, andS. B. Hendricks, a physical chemist, began a study ofaction spectra for such diverse phenomena as seed ger-mination, stem elongation, and photoperiodic controlof flowering. It soon became apparent that all these phe-nomena shared similar action spectra, with peaks in thered and far-red. More interesting, however, was the dis-covery that a response potentiated by red light could benegated if the red light treatment were followed imme-diately with far-red light. Such clear photoreversibilityhad never before been described in biology. This dis-tinctive characteristic led Borthwick and Hendricks topropose the existence of a novel pigment system, latercalled phytochrome.

This new, but hypothetical, pigment would existin two forms: a red-absorbing form called Pr and afar-red-absorbing form called Pfr. The pigment wouldalso be photochromic, which means that absorption oflight would alter its absorbance properties. Absorptionof red light by Pr would convert the pigment to thefar-red-absorbing form while subsequent absorption offar-red light by Pfr would drive the pigment back tothe red-absorbing form. Solely on the basis of simplephysiological experiments, they were able to predictseveral other features of this hypothetical pigmentsystem. First, because seeds and dark-grown seedlingtissues responded initially to red light, not far-red,the pigment was probably synthesized as the Pr form.Moreover, Pr was stable and probably physiologicallyinactive. Second, because treatment with red lightinitiated germination and other developmental events,Pfr was probably the active form. On the other hand,

Pfr was apparently unstable and was either destroyed orcould revert to Pr in darkness by a nonphotochemical,temperature-dependent reaction. Third, becausethe pigment could not be seen in dark-grown,chlorophyll-free tissue, it was no doubt present at verylow concentration. Borthwick and Hendricks furthersurmised that the pigment must be acting catalyticallyand was therefore possibly a protein. It is a tributeto the scientific acumen of these investigators andtheir coworkers that every one of these predictionswas later proven true. Yet, at the time, the existenceof phytochrome was met with some skepticism withinthe scientific community, largely because there was noprecedent for such a photoreversible pigment in theplant or animal research literature.

nm

+ 01

Red irradiated

Far-red irradiated

Difference SpectrumFar-red — Red

OD 0

OD = 0.1

- 01

600 650 700 750 800 850

Δ

Δ

FIGURE 22.2 Absorbance curves for maize shoots follow-ing red or far-red irradiations. Note that these curvesrepresent the absorbance of whole tissue, not just thepigment. Note also that far-red irradiation of the tis-sue, which converts the pigment from Pfr (solid curve)to Pr (dashed curve), causes the expected increase inabsorbance in the red and a decrease in the far-redregions of the spectrum. The difference spectrumeffectively represents the absorption spectrum ofthe Pr form. These data represent the first physi-cal demonstration of the existence of phytochrome.(From Butler, W. et al. 1959. Proceedings of the NationalAcademy of Sciences USA 45:1703–1708. Reprinted bypermission.)

376 Chapter 22 / Photomorphogenesis: Responding to Light

It was clearly necessary to obtain physical evidencefor the existence of phytochrome and, ultimately, toisolate the pigment and characterize it in vitro. Thestrategy that led to a satisfactory resolution of thisproblem turned on the same unique photoreversiblecharacter that generated skepticism in the first place.Since phytochrome was the only known photochromicpigment present in plants, it should be possible to detectabsorbance changes related to photoconversion of thepigment from one form to the other. Hendricks andhis colleagues thus predicted that the conversion of Prto Pfr in red light would be accompanied by a decreasein absorbance in the red (the maximum absorbance ofPr) and a corresponding increase in absorbance in thefar-red. Subsequent irradiation with far-red light shouldcause an increase in absorbance in the red and a decreasein the far-red. These experiments would require a special

kind of spectrophotometer, one capable of measuringvery small absorbance changes in dense, light-scatteringtissue samples. Fortunately, such an instrument wasunder development in another laboratory at Beltsvilleand relatively straightforward modifications wererequired to adapt its use for phytochrome detection.

The predicted photoreversible absorbance changeswere first demonstrated in dark-grown maize shoots in1959 (Figure 22.2). This spectral analysis was the firstphysical evidence that phytochrome actually existed.A short time later, the pigment was successfully iso-lated and purified from dark-grown cereal seedlings.In the years that followed, phytochrome was foundto be ubiquitous in the plant kingdom. It is found inalgae, bryophytes (mosses and liverworts), and proba-bly all higher plants where it plays a significant role inbiochemistry, growth, and development.

22.2.1 PHOTOREVERSIBILITY IS THEHALLMARK OF PHYTOCHROMEACTION

The key to the discovery of phytochrome was the findingthat plant responses to weak red light could be nullifiedif the red treatment were followed immediately withfar-red light (Table 22.1). Such clear photoreversibil-ity had never before been described in biology. Thedata in Table 22.1 are for groups of seeds that wereallowed to imbibe water in darkness for three hoursbefore being subjected to various brief light treatments.The light treatments were either 1 minute of red light(R; λ ca. 660 nm) or 3 minutes of far-red light (FR; λ>

700 nm) at low fluence rates, or alternating R and FR insuccession.

TABLE 22.1 Photoreversible control ofgermination. Lettuce seeds were imbibed for 3hours prior to irradiations. Irradiation times were1 min of low intensity red light and 3 min of Fr.Germination was scored after 48 h in subsequentdarkness at 20◦C.Irradiations Germination (%)

R 88R, Fr 22R, Fr, R 84R, Fr, R, Fr 18R, FR, R, FR, R 72R, Fr, R, Fr, R, Fr 22

Data from a student experiment.

Following irradiation, the seeds were returned todarkness for 48 hours, and the number of germinatedseeds in each lot was then counted. Note that redlight promotes a high germination rate but that a R,FRtreatment (red light followed immediately by far-redlight) maintains germination at the dark level (22%).When the R and FR treatments are alternated, thegermination rate depends solely on whether R or FRwas presented last. It is as though germination weredependent on a switch that could be turned on by redlight and turned off by far-red light. Similar results wereobserved with responses as diverse as stem elongation indark-grown pea seedlings and photoperiodic control offlowering in cocklebur (Xanthium strumarium).

Photoreversible behavior can be explained by a sim-ple, two-state photoequilibrium model for phytochrome(Figure 22.3). The red light-absorbing form of the pig-ment is designated Pr and the far-red-absorbing form,Pfr. Phytochrome is synthesized as the Pr form, which

Red light

Response

Far-red light

Pr Pfr

FIGURE 22.3 The simple, two-state photoequilibriummodel for phytochrome as originally postulated on thebasis of seed germination and other plant responses toalternating red and far-red light. Absorption of red lightby the red-absorbing form of the pigment (Pr) convertsit to a form that absorbs far-red light (Pfr). Conversely,absorption of far-red light by Pfr returns the pigment tothe Pr form. Pfr is considered the active form and initi-ates a signal transduction chain that leads to germination.

22.2 Phytochromes: Responding to Red and Far-Red Light 377

accumulates in dark-grown tissue and is generally con-sidered to be physiologically inactive. When Pr absorbsred light, it is converted to the Pfr form, which isthe physiologically active form of the pigment for mostknown responses. Exposure of Pfr to far-red light returnsthe pigment to the Pr form. Both physiological and spec-trophotometric experiments have also indicated thatsome Pfr may revert to Pr by a temperature-dependentprocess called dark reversion.

22.2.2 CONVERSION OF PR TO PFR INETIOLATED SEEDLINGS LEADSTO A LOSS OF BOTH PFR ANDTOTAL PHYTOCHROME

Most of the early work on phytochrome was conductedwith dark-grown, or etiolated, seedlings. Dark-grownseedlings grow quickly, they accumulate relatively largeamounts of phytochrome, and the absence of chloro-phyll makes it possible to measure phytochrome directlyin tissue with spectrophotometers adapted for use withoptically dense, light-scattering materials.2 With theappropriate instrument, changes in the total amount ofphytochrome and the relative proportions of Pr and Pfrcould be monitored following controlled irradiations.These in vivo spectrophotometric studies confirmedmany of the original predictions about the dynamicproperties of phytochrome.

Typical phytochrome transformations in etiolatedseedling tissue are shown in Figure 22.4. The most dis-tinctive feature is that Pfr is relatively unstable. Notethat when the tissue is returned to darkness after Pr isconverted to Pfr with a brief pulse of low fluence redlight, the concentration of Pfr declines with a half-life of1 to 1.5 hours. This loss of Pfr is accompanied by a cor-responding decline in the total amount of phytochrome.These kinetics can be explained by the fact that bothforms of the pigment are subject to irreversible chemicaldegradation (Figure 22.5). In darkness, Pr accumulatesuntil its rate of synthesis is matched by the rate of Prdegradation, which is relatively low. The loss of phy-tochrome following a red irradiation can be explained bytwo factors. First, the rate of Pfr degradation is approxi-mately 100 times greater than the rate of Pr degradation.Immunochemical studies have demonstrated the conju-gation of Pfr with ubiquitin, indicating that Pfr is subjectto degradation by the ubiquitin/26S proteasome system.Second, it has been demonstrated that Pfr suppressesthe transcription of the phytochrome gene by feedbackinhibition. As little as 5 seconds of red light causes a

2A spectrophotometer is an instrument for measuring theabsorption of light by pigments in solution. Conventionalspectrophotometers are by design limited to optically clearsolutions, free of light-scattering particles.

Ptotal

Pfr

Pr

Time in darkness (hrs.)

100

0 1 2 3

Rel

ativ

e co

ncen

trat

ion

FIGURE 22.4 Typical phytochrome transformations inetiolated seedling tissue. Dark-grown oat coleoptile tis-sue was given a short exposure to low fluence red lightat time 0. Absorbance changes were then monitored fortotal pigment and Pfr in the ensuing dark period. Pr wascalculated as the difference between total phytochromeand Pfr.

rapid decline in translatable phytochrome mRNA inetiolated seedlings.

Although it was not known at the time of the originalstudies, it is now clear that only phyA accumulates tohigh levels in etiolated tissue and is rapidly degraded inthe light. All other phytochromes (phyB–E) are stablewhen irradiated and are present in constant, althoughmuch lower, amounts regardless of the light conditions.

Pr

Pr ′ Pfr ′

PHY gene mRNA Response

Feedback inhibition

far-red light

Thermal reversion

red lightPfr

FIGURE 22.5 The phytochrome system, based on thebehavior of phytochrome in dark-grown seedling tis-sue. Phytochrome is synthesized as the physiologicallyinactive red-absorbing form (Pr), which accumulatesin darkness. Red light (660 nm) drives a phototrans-formation to the far-red-absorbing form (Pfr). Absorp-tion of far-red light (735 nm) returns the pigment to thePr form. Pfr, the active form, induces a response. Pr′ andPfr′ represent inactive degradation products of Pr andPfr, respectively. Pfr is known to revert to Pfr in darknessby a temperature dependent process. Pfr also suppressesphytochrome mRNA transcription by feedback inhibi-tion.

378 Chapter 22 / Photomorphogenesis: Responding to Light

22.2.3 LIGHT ESTABLISHES A STATE OFDYNAMIC PHOTOEQUILIBRIUMBETWEEN PR AND PFR

The absorption spectra for Pr and Pfr (Figure 22.1)show that both forms have broad, overlapping absorp-tion spectra. Note that Pfr absorbs some light at 660 nm(although much less efficiently than Pr) and Pr absorbsslightly in the far-red. Thus, even with ‘‘pure’’ red lightat 660 nm, it is not possible to convert 100 percent ofthe pigment to Pfr. As soon as Pfr appears, a portionof it will absorb red light and immediately phototrans-formed back to the Pr form. In a similar manner, Pr alsoabsorbs a small amount of far-red light (735 nm), so thateven in pure far-red light some Pr will be converted toPfr. In other words, regardless of what light source isused, a dynamic photoequilibrium (�) is established asphytochrome cycles between Pr and Pfr. This photoe-quilibrium is conveniently defined as � = Pfr/PTOT, inwhich PTOT is the total phytochrome or the sum of Prand Pfr. The photoequilibrium established by red light(660 nm) in etiolated tissues, for example, is 0.8 whilethe value for far-red light at 720 nm is 0.03. In otherwords, pure red light will maintain about 80 percent Pfrand 20 percent Pr while far-red light will establish about3 percent Pfr. For this reason, the absorption spectrumof Pfr shown in Figure 22.1 is actually the spectrumof an equilibrium mixture of Pr and Pfr following asaturating red light treatment.

Except in the laboratory, of course, plants do notgrow in dark boxes with occasional flashes of red andfar-red light. Moreover, sunlight contains a mixtureof red and far-red wavelengths and, depending upontime of day and environmental conditions, the relativeproportions of red and far-red wavelengths in sunlightwill change. The result is that sunlight will also producean equilibrium mixture of Pr and Pfr and, because thequality of sunlight changes throughout the day, so willthe equilibrium mixture of Pr and Pf. The biologicalresponse in most cases will depend on the proportionof Pfr, or (�) in the system. The proportion of Pfr will,in turn, depend on at least three factors: the relativeproportions of red and far-red wavelengths in the lightsource, the forward and reverse rates of photoconversionbetween Pr and Pfr, and the rate of thermal reversion ofPfr to Pr.

22.2.4 PHYTOCHROME RESPONSESCAN BE GROUPED ACCORDINGTO THEIR FLUENCEREQUIREMENTS

Phytochrome-mediated responses are convenientlygrouped into three categories on the basis of theirenergy requirements. The classical red, far-red photo-reversible responses discovered by Hendricks and

Borthwick and their colleagues are known as lowfluence responses (LFRs). LFRs are stimulated bylight doses in the range of 1 μmole m−2 to 1000 μmolem−2. This is equivalent to about a 0.1 second exposureunder a dense plant canopy at the lower end and aboutone second of full daylight at the upper end. In addition,LFRs are FR-reversible.

Phytochrome responses stimulated by light levels inthe range of 10−6 to 10−3 μmole−2 are called very lowfluence responses (VLFRs). Typically, such low levelsof light (comparable to the light emitted from a firefly)convert only about 0.01 percent of the phytochrome.Several studies have indicated that dark-grown seedlingsare capable of responding to such very low levels of light.Red light, for example, promotes an increase in sensitiv-ity of cereal grain seedlings to a subsequent phototropicstimulus. But the red light fluence required to saturatethe response was found to be at least 100 times lessthan that required to induce a measurable conversion ofPr to Pfr! A low far-red fluence also promotes pho-totropic sensitivity just as red light does, indicating thatless than 1 percent of the pigment need be converted toPfr in order to saturate the response. Exposure to eventhe traditional dim green safelights is sufficient to elicitor even saturate elongation responses in dark-grownAvena seedlings. For example, as little as 0.01 percentPfr is required to elicit inhibition of mesocotyl elonga-tion. This extreme sensitivity to light obviously makesthe study of VLFRs technically difficult. VLFRs, forexample, are not photoreversible. The principal evi-dence that a VLF response is mediated by phytochromeis the similarity of its action spectrum to the absorp-tion spectrum of Pr. The phenomenon, however, raisesperplexing yet intriguing questions about experimentalphotocontrol of plant development.

In the natural environment, plants are exposedto long periods of sunlight at relatively high fluencerates. Under such conditions, characterized by relativelyhigh energy over long periods of time, the photomor-phogenic program achieves maximum expression, andresponses such as leaf expansion and stem elongationare far more striking. Such light-dependent responsesare known as high irradiance reactions (HIRs). Highirradiance reactions generally share the following char-acteristics: (1) full expression of the response requiresprolonged exposure to high irradiance, primarily witha high proportion of far-red light; (2) the magnitudeof the response is a function of the fluence rate andduration; (3) like VLFRs, HIRs are not red, far-redphotoreversible.

The HIR has been implicated in a wide range ofresponses that also qualify as LFRs, including stemgrowth, leaf expansion, and seed germination. How-ever, HIRs may exhibit strikingly different action spec-tra depending on the species or growth conditions.Etiolated seedlings, for example, respond to blue, red,

22.4 Phytochrome and Cryptochrome Mediate Numerous Developmental Responses 379

and far-red light. As de-etiolation progresses, there isa shift from a far-red-sensitive HIR to a red-sensitiveHIR. Not surprisingly then, light-grown, green tissuesare more responsive to red light rather than far-red.Some systems, such as anthocyanin synthesis in Sorghumseedlings, respond only to blue-UV-A light.

Genetic studies involving phytochrome mutants inArabidopsis have identified that phyA is responsible forthe VLFR and FR-sensitive HIR responses and phyB isresponsible for the LFR and red-sensitive HIR.

22.3 CRYPTOCHROME:RESPONDING TO BLUEAND UV-A LIGHT

Charles Darwin was one of the first to note that plantsrespond to blue light when he observed that heliotropicmovements were diminished in light passed through asolution of potassium dichromate, an effective absorberof blue light. It is now known that those portionsof the spectrum that constitute blue and UV-A lightregulate many aspects of growth and development inplants, fungi, and animals. Plant responses to blue andUV-A light include aspects of de-etiolation such asthe inhibition of hypocotyl elongation and stimulationof cotyledon expansion, the opening and closure ofstomata, gene expression, flowering time, the ‘‘setting’’of endogenous clocks (Chapter 24), and the growth ofshoots in response to light gradients, or phototropism(Chapter 23).

Most action spectra for blue-light responses hadpeaks in the blue and UV-A regions of the spec-trum and closely resembled the absorption spectra offlavin molecules, such as riboflavin. This promptedA. W. Galston to postulate as early as 1950 that aflavoprotein was involved in blue light responses. Oth-ers, however, mounted strong arguments in favor of acarotenoid-based photoreceptor, and for many years theflavin–carotenoid controversy was hotly debated. Thecontroversy was difficult to resolve because plants con-tain a bewildering array of flavoproteins and caroteno-proteins, a factor that seriously complicated any effortsto identify the one that might be involved specifically inblue-light responses. Because of this elusive, or ‘‘cryp-tic,’’ nature of the pigment and the pervasive blue-lightresponses in cryptogams, or nonflowering plants, thepigment was referred to as cryptochrome.

The first protein with the characteristics of ablue-light photoreceptor was isolated from Arabidopsisin 1993. Elongation of hypocotyls in wildtypeArabidopsis can be inhibited by red, far-red, or blue light.Several mutants (the hy mutants) were characterized byelongated hypocotyls because they had lost the capacityto respond to one or more regions of the spectrum. One

of these, hy4 (later named cry1), had lost the capacity torespond specifically to blue light. The protein encodedby the wildtype allele was subsequently isolated and, onthe basis of its photobiological and genetic properties,identified as cryptochrome 1 (cry1). The argumentbetween the flavin camp and the carotenoid camp wasfinally resolved after more than 40 years when it wasshown that cry1 binds two flavin-related chromophores,one of which is flavin adenine dinucleotide (FAD)(Section 22.5.3).

A second cryptochrome, cry2, has also been identi-fied. Cry2 mediates blue-light suppression of hypocotylelongation, cotyledon expansion, and anthocyanin pro-duction in Arabidopsis. In addition, cry2 has a role indetermining flowering time and is synonymous withFHA, the product of a flowering-time gene. The roles ofcry2 and FHA in flowering are examined in Chapter 25.

22.4 PHYTOCHROME ANDCRYPTOCHROMEMEDIATE NUMEROUSDEVELOPMENTALRESPONSES

Phytochrome and cryptochrome act both jointly andindependently to regulate a wide range of developmentalresponses. Some of the better understood responses aredescribed in the following sections.

22.4.1 SEED GERMINATION

The germination of many seeds is influenced by light asevident in the flush of germination in areas of cultivationor natural disturbance. Some seeds, known as positivelyphotoblastic seeds, are stimulated to germinate bylight. The germination of others, known as negativelyphotoblastic seeds, is inhibited by light. Some seeds,mostly agriculturally important species that have beenselected for high germinability, are not affected by light.Many seeds, such as lettuce, may require only briefexposure to light, measured in seconds or minutes, whileothers may require as much as several hours or even daysof constant or intermittent light (e.g., Lythrum salicaria,Epilobium cephalostigma). In all cases, the responsiblepigment appears to be phytochrome.

Most seeds that require light for germination tend tobe very small seeds that have comparably small embryosand limited endosperm. They need to be close to thesurface when they germinate so that the young seedlingcan reach the sunlight before the reserves are exhausted.Most soils attenuate light very quickly. A 1 mm thicknessof fine soil, for example, passes less than 1 percent ofthe incident light and then only at wavelengths longerthan 700 nm. As a result, most light-requiring seeds

380 Chapter 22 / Photomorphogenesis: Responding to Light

need not be buried very deeply for germination tobe held in check. However, some seeds (e.g., Sinapisarvensis) require very little Pfr (� = 0.05) to stimulategermination and may exhibit germination when coveredwith up to 8 mm of soil. Thus, the role of phytochromein both seed germination and seedling developmentappears to be one of conveying information to the seedor seedling about its position relative to the soil surface.

Interestingly, most common agricultural weedssuch as Amaranthus (pigweed), Ambrosia (ragweed),and Chenopodium (lambs quarters) produce prodigiousnumbers of very small light-sensitive seeds. These seedsaccumulate in the ‘‘seed bank’’ just below the surfaceof the soil where they will not germinate. Every timethe soil is disturbed, however, a new batch of seedsgerminates because they are exposed to light. Thisis a major factor in the competitive success of thesespecies.

Suppression of germination in negatively photo-blastic seeds, such as wild oats (Avena fatua), generallyrequires long-term exposures at high fluence rates.Far-red and blue light are most effective, although insome cases (e.g., Phacelia tanacetifolia) red light is alsoeffective. Photoinhibition of seed germination appearsto be an example of a high irradiance reaction. InArabidopsis, seed germination is controlled solely byphytochromes.

22.4.2 DE-ETIOLATION

Plants grown in darkness exhibit a distinct morphology.The details may vary from one species to another, but ina dicot such as bean (Figure 22.6) the hypocotyl is elon-gated and spindly, with a pronounced plumular hook,or recurve, just below the first leaves. The leaves them-selves undergo limited development and remain smalland clasping, as they were in the embryo. Chlorophyll isabsent and the seedlings appear white or yellow in color.In monocot cereal grains the first internode, or meso-cotyl, elongates excessively in the dark and the coleop-tile, which is a modified leaf, grows slowly. The primaryleaves remain within the coleoptile and stay tightly rolledaround their midvein. This general condition exhib-ited by dark-grown seedlings is called etiolation. Othercharacteristics of the etiolated condition include arrestedchloroplast development and low activities of manyenzymes. When exposed to light, etiolated seedlingsundergo de-etiolation, a process under control of bothphytochromes and cryptochromes. Hypocotyl growthis arrested, the plumular hook gradually straightens,and elongation of the epicotyl accelerates. Light alsostimulates the leaves to unfold, enlarge, and completetheir development. Chloroplast development also pro-ceeds and the leaves green up as chlorophyll accumulatesand the leaves become photosynthetically competent. In

FIGURE 22.6 The de-etiolation response in 7-day-oldseedlings of bean (Phaseolus vulgaris). The seedling onthe right was grown in darkness. Note the elongatedhypocotyl, recurved plumular hook, an absence of pri-mary leaf expansion, and absence of chlorophyll. Theseedling at left was grown under normal white light con-ditions. Note the shortened hypocotyl, unfolding of theplumular hook, expanded primary leaves, and accumula-tion of chlorophyll. The center seedling was exposed to5 minutes of weak red light daily for three days, which issufficient to initiate the de-etiolation response.

particular, light stimulates the synthesis of the chloro-phyll a/b light harvesting pigment-protein.

The developmental significance of etiolation andde-etiolation is not difficult to construct. Remember thata plant is fundamentally a photosynthetic organism. Aseed carries a limited amount of nutritive tissue that mustsupport development of the seedling until such time asits leaves are established in the light and photosynthesiscan take over the supply of energy and carbon. Inthe darkness experienced beneath the soil, the limitedreserves of a seed are committed to extension growthof the hypocotyl in order to maximize the possibilitythat the plumule, composed of the young leaves, willreach the light and be able to carry out photosynthesisbefore the reserves are exhausted. Once establishedin the light, the remaining reserves may be investedin development of photosynthetic tissue, such as leafexpansion and chloroplast development.

22.4 Phytochrome and Cryptochrome Mediate Numerous Developmental Responses 381

22.4.3 SHADE AVOIDANCE

Plants need light for photosynthesis and those that findthemselves growing in the shade of neighbors or undera canopy can adjust to the reduced availability of lightin two ways. They can (1) adjust their light-harvestingcapability by increasing the specific leaf area and theamount of chlorophyll a/b light-harvesting complex orthey can (2) adjust their morphology in order to positiontheir leaves out of the shade. Plants typically respond toshadelight with increased elongation of stem-like organs(including hypocotyls and leaf petioles, a more upwardorientation of leaves (hyponasty), reduced branching,and reduced tillering (in grasses). In the end, shad-ing leads to early flowering and seed set in an effortto ‘‘escape’’ shading by shortening generation time.These and other effects are collectively called the shadeavoidance syndrome.

Radiation within and below a canopy is markedlydeficient of red and blue light because these wavelengthsare largely absorbed by the chlorophyll in the overly-ing leaves. By contrast, chlorophyll is transparent tofar-red light; any attenuation of far-red is limited almostsolely to reflection. Plants therefore use phytochrome(specifically phyB, phyD, and phyE) and cryptochrometo detect these characteristic differences in compositionbetween shadelight and unfiltered daylight.

The effect of canopy shading can be described interms of the ratio of red to far-red fluence rates (R/FR,or ζ ; Gr. zeta). The value of ζ in unfiltered daylightis typically in the range of 1.05 to 1.25. The value inshadelight beneath a canopy will, of course, vary with thenature of the vegetation and the density of the canopy.Some representative values are listed in Table 22.2.These values fall well within the range where a smallchange in ζ would cause a relatively large change in theproportion of Pfr (Figure 22.7).

Shadelight can be mimicked in the laboratory orgrowth chamber by supplementing white fluorescentlight (ζ = 2.28) with various amounts of far-red lightthrough the entire photoperiod. This can be done insuch a way that the fluence rate of photosyntheti-cally active radiation (PAR) remains constant from one

TABLE 22.2 Approximate values of R/FR (ζ ) forcanopy filtered light.

Canopy R/FR

Wheat 0.5Maize 0.20Oak woodland 0.12–0.17Maple woodland 0.14–0.28Spruce forest 0.15–0.33Tropical rainforest 0.22–0.30

Under clear water1.15 Max.

Daylight 1.05 - 1.25

Twilight 0.65 - 1.15

Canopy shade 0.05 - 1.15

10 2 3 4

R:FR Ratio (ζ)

12 14 16

0.8

0.6

0.4

0.2

0E

stim

ated

phy

toch

rom

e ph

otoe

quili

briu

mP

fr/P

tota

l (φ)

FIGURE 22.7 Phytochrome photoequilibrium ( ) isrelated to the ratio of red to far-red light (ζ ). Blockedareas indicate the range of values for ζ observed underindicated ecological conditions. (Reproduced with per-mission from the Annual Review of Plant Physiology, Vol.33. Copyright 1982 by Annual Reviews, Inc.)

treatment regime to the next. This removes the impactof photosynthetic output and any differences in growth;morphogenesis can thus be attributed to the phy-tochrome photoequilibrium value (�), which can beestimated from the measured R/FR ratio for each reg-imen. When young plants of Chenopodium album weregrown this way, the logarithmic stem extension ratewas found to be linearly related to � (Figure 22.8).The response to light quality may be quite rapid. InChenopodium, for example, an increase in the stem exten-sion rate can be observed within seven minutes of addingFR light to the background fluorescent source. Evenwhen not directly shaded, plants are able to anticipateshading. The reduction in red to far-red ratio reflectedfrom the leaves of nonshading neighbors and propa-gated horizontally is often sufficient to initiate shadeavoidance responses.

22.4.4 DETECTING END-OF-DAYSIGNALS

There are also substantial changes in the spectral energydistribution of natural daylight on a daily basis. At bothdawn and dusk, when the sun sits low on the horizon,there are significant relative decreases in the value of ζ

382 Chapter 22 / Photomorphogenesis: Responding to Light

FIGURE 22.8 The growth ofChenopodium album seedlings in simu-lated shadelight. Seedlings were grownfor 14 days in shadelight simulated byproviding supplementary far-red lightsufficient to provide the indicated R:FRratios (ζ ). Note that internode elon-gation is inversely proportional to theamount of Pfr in the tissue. (Reproducedby permission of Dr. David Morgan andProfessor Harry Smith, University ofLeicester, UK. Photograph courtesy ofProf. Smith.)

compared with the main part of the day. In one study,for example, ζ values at dusk were reduced by 14 to 44percent of those at midday. A detailed examination ofthe response of pumpkin (Cucurbita pepo) to end-of-dayred or far-red light reveals that a reduction in the pro-portion of phytochrome maintained as Pfr at the endof the photoperiod is associated with drastic changes inthe developmental pattern. Lowering of � from a highvalue (ca. 0.65 to 0.75) to a very low value (ca. 0.03)accentuates stem and petiole extension, reduces leafexpansion and branching, and lowers the chlorophyllcontent. These experiments do not, of course, provea causal link between phytochrome photoequilibriumand morphogenic responses to changes in the radia-tion environment. They do, however, demonstrate thatplants have the capacity to respond to changes in spectralenergy distribution similar to those that occur naturallyin the environment. It also seems highly likely that phy-tochrome is the photoreceptor that detects end-of-daysignals, which could reflect an important role in timemeasurement (Chapters 24, 25)

22.4.5 CONTROL OF ANTHOCYANINBIOSYNTHESIS

Anthocyanins are the water-soluble red and blue pig-ments responsible for the color of many vegetables,fruits, and flowers. The biosynthesis of anthocyaninsis a classical high irradiance reaction, first revealed instudies of red cabbage seedlings. Like other responsesof etiolated seedlings, the initiation of anthocyanin accu-mulation is a classic phytochrome-dependent LFR. Thered, far-red photoreversibility, however, is limited to

brief irradiations. When longer-term irradiations areapplied, the action peak for continued anthocyaninaccumulation is shifted to the far-red, with reducedeffectiveness in the red. This effect of prolonged far-redirradiation has been interpreted as a requirement formaintaining a low level of Pfr over time—long enoughto avoid rapid depletion of the Pfr pool by degradation.

22.4.6 RAPID PHYTOCHROMERESPONSES

The response time for most phytochrome-mediateddevelopmental effects is measured in hours or even days,but there are some responses with response times mea-sured in minutes or seconds. Most, but not all, of theserapid responses appear to be related to membrane-basedactivities such as bioelectric potential or ion flux. Oneof the earliest indications that phytochrome influencedthe electrical properties of tissues was a curious effect onthe surface charge of root tips reported by T. Tanadain 1968. Tanada observed that dark-grown barley roottips would float freely in a glass beaker with a speciallyprepared negatively charged surface. Within 30 secondsfollowing a brief red irradiation the root tips wouldadhere to the surface. A subsequent far-red treatmentwould release the root tips from the glass. It was laterfound that adhesion and release was correlated withphytochrome-induced changes in the surface potentialof the root tips. A brief red treatment generated apositive surface potential, attracting the tips to the neg-atively charged surface. A far-red treatment generateda negative surface potential, thereby causing the tipsto detach. Similar effects of red and far-red light on

22.5 Chemistry and Mode of Action of Phytochrome and Cryptochrome 383

surface potential of Avena coleoptiles have also beendemonstrated.

Phytochrome-modulated transmembrane poten-tials have since been reported for a variety oftissues from several laboratories. The results are notcompletely consistent, but in most cases red lightinduces a depolarization of the membrane within5 to 10 seconds following a red light treatment. Asubsequent far-red treatment causes a slow return tonormal polarity or small hyperpolarization. At thispoint, it is not known whether such effects are due toa direct action of phytochrome on the membrane orwhether a second messenger system is involved.

22.4.7 PhyA MAY FUNCTION TODETECT THE PRESENCEOF LIGHT

What purpose does it serve the plant to have multipleforms of phytochrome? More specifically, of what ben-efit is it for etiolated plants to accumulate an apparentexcess of labile phyA, which is so quickly degraded inthe Pfr form? One possibility is that phyA functionsonly to detect the presence of light, rather than to dis-tinguish subtle differences in light quality. Note thatphyA accumulates in two particular situations: (1) inseeds that require red light to germinate and conse-quently do not germinate when buried deep in the soil,and (2) in germinated seedlings in which phytochromeis used to detect light as the seedling approaches thesoil surface. The large amount of phyA that accumu-lates under both of these conditions appears to functionas a sensitive antenna or photon-counter that detectsonly the presence of light. Once the seed or seedlingis exposed to adequate light, the excessive quantity oflabile phytochrome disappears. This allows the morestable phyB to monitor the R-FR ratio over time

and direct development accordingly. It may be diffi-cult to obtain direct evidence in support of such ascenario, but it is an important first step in taking phy-tochrome studies out of the laboratory and into the realworld.

22.5 CHEMISTRY AND MODE OFACTION OF PHYTOCHROMEAND CRYPTOCHROME

22.5.1 PHYTOCHROME IS APHYCOBILIPROTEIN

The phytochrome chromophore (Figure 22.9) is calledphytochromobilin because it is a linear tetrapyrrole thatis similar in structure to mammalian bile pigments. Phy-tochromobilin is virtually identical to phycocyanobilin,the chromophore of the pigment phyconcyanin foundin cyanobacteria and the red algae. Phytochromobilin isalso related to the chromophores of chlorophyll, the res-piratory cytochrome pigments, and hemoglobin, exceptthat these pigments are all cyclic tetrapyrroles.

The difference between the Pr chromophore andthe Pfr chromophore is a rotation (a cis–trans isomeriza-tion) of the double bond between rings C and D. Theabsorption of red light provides the energy requiredto overcome the high activation energy for rotationaround the double bond, a transition that is not nor-mally possible at ambient temperature. The pigment canbe returned to the more stable Pr configuration by eitherFR light or thermal-dependent reversion in darkness.There is also evidence that the change in the chro-mophore induces substantial conformational changes inthe protein, which would account for its activation whenconverted to Pfr.

Cys

S

A

B CO

COOH COOH

NH

D

NH

NH HN

Pr Pfr

C

COOHFar-red

Red

D

O

HN

HN

FIGURE 22.9 The structure of the phytochrome chromophore and its binding tothe apoprotein. The chromophore, a linear tetrapyrrole, is covalently linked to thepolypeptide chain at cysteine-321 via a sulfur bond. Photoconversion between Pr andPfr involves a rotation around the double bond linking the C and D rings.

384 Chapter 22 / Photomorphogenesis: Responding to Light

Pfr-Specific phosphorylationLight

N

N

PCB

PCB

P

P

S

S D

D HKRD C

CHKRD

Photosensory domain Regulatory domain

P1 Cys P4

P1 Cys P4

FIGURE 22.10 The structure of phytochrome. This blockdiagram of two phytochrome molecules (a dimer) showsthe principal domains and subdomains. The N-terminalphotosensory domain binds the phytochromobilin chro-mophore (PCB) and is linked by a hinge region to theC-terminal regulatory domain. The regulatory subunitcontains the dimerization domain (DD) and the histidinekinase-related domain (HKRD). One possible phospho-rylation site is shown (dashed line); a serine residue (S)located in the hinge region that is phosphorylated specif-ically in the Pfr form. Other possible phosphorylationsites are serine residues located in the P1 subdomain atthe N-terminus of the photosensory domain. The P4domain is believed to interact with the D-ring of thechromophore to stabilize the energetically unfavorablePfr form. The regulatory domain presumably initiatestransduction of the light signal.

Phytochrome apoprotein is a relatively small pro-tein with a molecular mass of 125 kDa. It consists oftwo structural domains of approximately equal size; aglobular N-terminal domain and an open, or extended,C-terminal domain (Figure 22.10). The A ring ofthe chromophore is covalently linked to the proteinthrough a cysteine residue in the N-terminal domain.The N-terminal end of the molecule is thus calledthe photosensory domain. The C-terminal domain,or regulatory domain, contains a subdomain that hasthe characteristics of a histidine kinase (the histidinekinase-related domain, HKRD). In this respect, phy-tochrome resembles a two-component system like thosepreviously encountered in osmoregulation and hor-mone sensing. In vivo, phytochromes exist as a dimer,with one chromophore on each monomer.

22.5.2 PHYTOCHROME SIGNALTRANSDUCTION

Phytochrome regulation of proteins or proteinfunction was first reported in 1960 by A. Marcus,who demonstrated red, far-red reversible control ofglyceraldehyde-3-phosphate dehydrogenase activity inbean seedlings. Since then, the list of enzymes andother proteins whose activities are known to be lightregulated, in most cases by phytochrome, has grown to

more than 60. These observations led to the obviousconclusion that phytochrome acted by controlling geneexpression. Only recently, however, have the results ofgenetic studies suggested mechanisms for the control ofgene expression by phytochrome. Although the detailedmechanism of phytochrome signaling is still not clear,at least three important factors have been identified:

1. the phytochromes are serine/threonine kinases thatboth autophosporylate and phosphorylate a numberof other proteins,

2. phytochromes are imported into the nucleus in theactive Pfr form, and

3. phytochrome responses are associated with signifi-cant changes in gene expression.

Early studies of the amino acid sequence of the phy-tochrome molecule revealed that its carboxy-terminaldomain had amino acid sequences homologous to bac-terial histidine kinase enzymes. Because of this similarityit was suggested in the 1980s that phytochromes mightbe protein kinases. There was, however, no directevidence of such activity. This all changed with thediscovery of phytochrome in the cyanobacteria. Thecyanobacterial phytochrome (cph1) has an N-terminaldomain similar to the chromophore-binding domainof plant phytochrome, has spectral properties similarto plant phytochromes, and, most importantly, exhibitslight-mediated histidine kinase activity. Shortly after-ward, it was shown that purified oat phytochrome Aexhibited light-regulated autophosphorylation ability.However, plant phytochrome autophosphorylates serineresidues, rather than histidine residues as its cyanobac-terial counterpart does. Phytochrome is preferentiallyphosphorylated in the Pfr form (Figure 22.10) whichmay then initiate signaling by transferring the phos-phate group to an appropriate substrate molecule. Itis well known that the addition of a phosphate groupor its removal has profound effects on the structureand stability of proteins, thereby influencing functionalproperties and intracellular location. Phytochrome mayregulate various aspects of development simply by phos-phorylating different substrates or initiating differentkinase cascades.

At least two putative phosphorylation substrateshave been identified. PKS1 (PHYTOCHROMEKINASE SUBSTRATE 1) is a cytoplasmic proteinthat is phosphorylated by phyA in vitro. PKS1 binds toboth phyA and phyB and its phosphorylation in vivo isstimulated by red light, suggesting that phytochrome isthe responsible kinase. Nucleoside diphosphate kinase2 (NDPK2) is an enzyme that also appears to interactwith phytochrome. NDPK catalyzes the synthesisof various nucleoside triphosphates (e.g., CTP,GTP, UTP) from ATP and the corresponding nucleo-side diphosphate. Its activity is significantly increasedin the presence of the Pfr form of phyA in vitro,

22.5 Chemistry and Mode of Action of Phytochrome and Cryptochrome 385

phosphorylation of NDPK2 is stimulated by red lightin vivo, and ndpk2 mutants exhibit altered responsesto both red and far-red light. Purified phyA canalso phosphorylate both cryptochromes in vivo andphosphorylation of cry1 is also stimulated by redlight in vivo. This is particularly interesting in viewof the known physiological interactions between redand blue light responses and raises the possibilitythat phyA might modulate blue light responses viaphosphorylation of cryptochrome (see Section 22.5.3).Unfortunately, a direct link between kinase activity andphytochrome signaling has yet to be established.

In order to control gene expression, the phy-tochrome signal chain must extend into the nucleuswhere the genes are located. In fact the signal chain inthis case is rather short, because phytochrome itself canmove into the nucleus. For a long time it was assumedthat phytochrome was strictly a cytosolic protein. Thisview changed when phytochrome was tagged by fusing itwith a green fluorescent protein (GFP) and the fusionproduct was then expressed in transgenic plants. TheGFP-fused phytochrome remains biologically active andthe location of the tagged phytochrome can be visuallyconfirmed by microscopic examination. These studiesmade it clear that phytochrome in its inactive Pr formdoes indeed accumulate in the cytoplasm, but conver-sion to the Pfr form unmasks a nuclear localizationsequence that allows phytochrome to be recognizedby the nuclear import machinery (Figure 22.11). Themechanism of nuclear import is not clear, but it hasbeen shown that the recognition sequence is locatedin the carboxy-terminal domain of the phytochromemolecule and that a protein identified as FHY1 (fromthe mutant fhy1, or far-red elongated hypocotyl 1) is specif-ically required. The rate of nuclear import varies. PhyA,

Cytoplasm

Nucleus

?

?

PrPr

PfrPr

PfrPfr

FIGURE 22.11 Import of phytochrome from the cyto-plasm into the nucleus is an important step in phy-tochrome signal transduction. Nuclear import requiresthat at least one of the two molecules in the dimer be inthe Pfr form.

for example, is transported into the nucleus within 15minutes of the onset of light, but phyB is not detectedfor at least two hours.

As multiple phytochromes arrive inside thenucleus, they appear to form aggregates referredto as nuclear bodies or speckles. Phytochrome alsophysically interacts, at least in vitro, with a family oftranscription factors called phytochrome-interactingfactors (PIF), an interaction that establishes a directlink between phytochrome and gene expression.Genetic analysis has established that PIF and PIF-like(PIL) proteins are predominantly negative regulatorsof phytochrome-dependent pathways. For example,members of the PIF family inhibit seed germination,control protochlorophyllide accumulation and repressphytochrome-regulated gibberellin biosynthetic genes.Most PIF proteins are stable in the dark, but arerapidly degraded in the light. The degradation of atleast two PIFs (PIF1 and PIF3) is mediated by the26S-proteosome system in a phytochrome-dependentmanner. Once the light is switched off, PIF proteindegradation stops and the PIF proteins rapidlyaccumulates.

Based on the above observations, a generalmodel for phytochrome action has begun to emerge

Cytoplasm Nucleus

Promotor

Promotor Gene

mRNA

Gene

Gene

Degradationby 26s proteosome

FR

Promotor

Red light

PIF

PIF

PIF

PfrPfr

Pr

Pr

Pfr

FIGURE 22.12 A model for the control of gene activa-tion by phy. Phytochrome interacting factor (PIF) in thenucleus is a negative regulator and probably repressestranscription by binding with the promoter region of aphotoresponsive gene. Red light converts cytosolic phyto its active Pfr form, which then is imported into thenucleus. The Pfr recruits the promoter-bound PIF fordegradation by the 26S-proteosome system, thus reliev-ing the repression of the gene. Far-red light will convertthe Pfr back to Pr, which immediately dissociates fromPIF, thus allowing PIF to reassociate with the promoterand reestablish gene repression.

386 Chapter 22 / Photomorphogenesis: Responding to Light

(Figure 22.12). In the dark, phytochrome accumulatesand remains in the cytoplasm as Pr, which is inactive.Meanwhile, nuclear PIF proteins inhibit the expressionof phytochrome-dependent genes. Upon irradiation,Pr is converted to Pfr, which is then imported intothe nucleus where it targets transcriptional regulatorsfor degradation, thereby activating transcription ofphytochrome-responsive genes. This model, however,does not explain all phytochrome-dependent responses.The rapid phytochrome responses described above(22.4.5) would seem to require a cytoplasmic, perhapseven membrane-located, phytochrome system. There issome evidence that these responses may involve cyclicGMP and/or calcium second messenger systems.

22.5.3 CRYPTOCHROME STRUCTUREIS SIMILAR TO DNA REPAIRENZYMES

Most plant cryptochromes are 70 to 80 kDa proteinswith two recognizable domains (Figure 22.13). TheN-terminal domain shares amino acid sequencehomologies with microbial DNA photolyase, a uniqueclass of flavoenzymes that use blue light to catalyzerepair of UV-induced damage to microbial DNA.The N-terminal domain of cryptochrome is thereforecalled the photolyase-related (PHR) domain. The PHRdomain of cryptochrome binds two chromophores.The first is flavin adenine dinucleotide (FAD), the sameredox cofactor encountered in respiratory metabolism.The second is 5,10-methenyltetrahydrofolate, orMTHF. MTHF is a member of another family of redoxcofactors called pterins (Figure 22.14). The second,or C-terminal, domain of cryptochrome has no knownhomologies with other proteins and its function is notclear.

The principal differences between photolyase andthe cryptochromes are that the cryptochromes have thedistinguishing carboxy-terminal extension not found in

Cryptochrome 1

MTHF

Photolyase-related domain

Photolyase

N

N C

C

FAD

FADpterin

FIGURE 22.13 Cry1 and cry2 share a similarphotolyase-related domain (the amino acid sequence isabout 58 percent identical) but differ primarily in thelength of the C-terminal domain. The two chromophoresare attached to the photolyase-related domain. TheC-terminal domain contains the amino acid sequencethat determines nuclear localization of the pigment.

C

O

O

AB

N N

NHN

CH3

CH3

R

Flavin

O

N

NHN

NH2N

R

Pterin

5-10-Methenyltetrahydrofolate (MTHF)

H2N

CH2

NO

HH

10

5H

H N

C H

R

N

N

N

FIGURE 22.14 The core ring structures of cryptochromechromatophores. Depending on the nature of the Rgroup, the flavin nucleus forms either flavin-adeninedinucleotide (FAD), flavin mononucleotide (FMN) orriboflavin. 5,10-Methenyltetrahydrofollate (MTHF) is apterin derivative. Note the similarity in the structuresof pterin and the B and C rings of flavin. The R groupof MTHF consists of 3 to 6 glutamate molecules. Bothflavins and pterins absorb strongly at the blue end of thespectrum.

the photolyases and cryptochromes exhibit no pho-tolyase activity. The similarity does, however, raise thepossibility of an evolutionary relationship between thephotolyases and the cryptochromes and it has been sug-gested that microbial photolyases are the evolutionaryprecursors for plant cryptochromes.

22.5.4 CRYPTOCHROME SIGNALTRANSDUCTION

Unlike phytochrome, the molecular consequences ofphotoexcitation of cryptochrome remain unknown.Other flavoproteins are known to participate almostexclusively in oxidation–reduction reactions and pho-tolyases repair damaged DNA by transferring electronsto pyrimidine dimers. It seems likely that the primaryphotochemical event when cryptochromes absorbblue light would involve a similar electron-transfermechanism.

22.7 De-Etiolation in Arabidopsis: A Case Study in Photoreceptor Interactions 387

The signal transduction chain for the crypto-chromes appears to be relatively short since thecryptochromes are located mainly in the nucleus. Cry1is located in the cytoplasm in the light but moves intothe nucleus in the dark while cry2 seems to residepermanently in the nucleus. In the nucleus, cryp-tochrome interacts directly with the COP1 (CONSTI-TUTIVELY PHOTOMORPHOGENIC 1), an E3ubiquitin ligase. COP1 is a major repressor of photo-morphogenic responses by constantly degrading anumber of transcription factors. The cop1 mutant there-fore displays all of the characteristics of a light-grownseedling (a constitutive photomorphogenic phenotype)in the dark. When irradiated with blue light, cryp-tochrome undergoes a conformational change whichleads to the deactivation of COP1 and accumulation oftranscription factors necessary for proper developmentin the light.

A number of experiments have shown thatcryptochromes are subject to light-dependent phospho-rylation. In one study, for example, it was shown thatthe C-terminal domain of cry1 was phosphorylated byphyA in vitro. Phosphorylation of cry1 by phyA occurredwith both red and blue light. In other experiments, bothcry1 and cry2 exhibited blue light-dependent phos-phorylation in vivo. The in vivo blue light-dependentphosphorylation of cry2 was also detected in a range ofphytochrome mutants—that is, in the absence offunctional phytochrome. Thus, while phytochromecan phosphorylate cryptochrome, blue light-dependentphosphorylation of cryptochrome is independent ofphytochrome. The implication is that cryptochromein the dark is not phosphorylated and, consequently,inactive. The absorption of blue light by cryptochromeenables its phosphorylation by some unknown kinase.The phosphorylated form of cryptochrome is activeand initiates signal transduction.

22.6 SOME PLANT RESPONSESARE REGULATED BY UV-BLIGHT

A number of plant responses are attributed to radiationin the UV-B region of the spectrum. A positive effect ofultraviolet light on anthocyanin accumulation has beenknown since the mid-1930s. Later it was recognized thatsunlight filtered through window glass, which absorbsultraviolet rays, was less effective than unfiltered sun-light. This effect was finally characterized when it wasdemonstrated that flavonoid biosynthesis in parsley (Pet-roselinum crispum) cell suspension cultures and seedlingswas induced by UV-B radiation (280–320 nm). Maxi-mum effectiveness was at 290 to 300 nm, with little or noactivity beyond 320 nm. By 1986, 11 species of higherplants were listed for which UV-B induced anthocyanin

and flavonoid biosynthesis in coleoptiles, hypocotyls,seedling roots, and cell culture.

In Sorghum bicolor the action spectrum for flavonoidbiosynthesis shows three peaks: 290 nm, 385 nm, and650 nm. Action at 385 nm and 650 nm could be reversedby a subsequent exposure to far-red, but the peak at290 nm could not. The 385 nm and 650 nm peaks havebeen attributed to phytochrome, leaving the 290 nmpeak due to a UV-B receptor. In parsley it appears thatflavonoid biosynthesis results from the coaction of threepigments: phytochrome, a separate blue receptor (prob-ably cryptochrome), and a UV-B receptor. The UV-Bsystem is a necessary prerequisite to flavonoid biosyn-thesis since neither the blue receptor nor phytochromeis effective unless preceded by a UV-B light treatment.

The UV-B receptor has yet to be isolated andits identity remains unknown. Phytochrome has beensuggested—the protein moiety does absorb UV-Blight—but results such as those described above wouldargue against it. In members of the Leguminoseaefamily, ultraviolet-induced flavonoid biosynthesis canbe reversed with blue light in a manner reminiscent ofphotoreactivation of UV damage in microorganisms.This could implicate DNA itself as a UV photoreceptor,but the action peak is shifted to wavelengths somewhatshorter than those normally characteristic of UV-Baction.

22.7 DE-ETIOLATION INARABIDOPSIS: A CASE STUDYIN PHOTORECEPTORINTERACTIONS

Arabidopsis is a typical dicot seedling in that growth inwhite light is accompanied by (1) arrested hypocotylelongation, (2) a straightening of the hypocotyl orplumular hook, (3) unfolding of the cotyledons, and(4) expansion of the cotyledons. Studies withphytochrome- and cryptochrome-deficient mutantshave confirmed that de-etiolation responses inArabidopsis seedlings involve complex interactionsbetween three different photoreceptors: phyA, phyB,and cry1. An experiment demonstrating some ofthese interactions is illustrated in Figure 22.15. In theexperiment illustrated here, elongation of the hypocotylin wildtype seedlings was suppressed by approximately68 percent in white light. The phyA-deficient singlemutant (phyA) had little effect on hypocotyl elongation,while in the absence of phyB, hypocotyl elongation wassuppressed by only 20 percent. These results indicatethat phyB is the principal photoreceptor for control ofhypocotyl elongation. phyA still has a role, however, afact which is uncovered in the double mutant, phyAphyB.In the absence of both phyA and phyB, not only ishypocotyl elongation not suppressed by white light,

388 Chapter 22 / Photomorphogenesis: Responding to Light

phyAphyBcry1

phyAphyB

phyA phyB cry1Light

Wildtype Mutants (in white light)

Dark

FIGURE 22.15 The effect of phytochrome- andcryptochrome-deficient mutations on Arabidopsishypocotyl elongation in white light. (Based on the data ofNeff, M. M., J. Chory. 1998. Plant Physiology 118:27–36.)

the hypocotyls are even longer than in dark-growncontrols. In other words, the phyA mutation appears tohave enhanced the effect of the phyB mutation.

Both straightening of the hypocotyl hook andunfolding of the cotyledons in white light wererelatively unaffected by any one of the three singlemutants, phyA, phyB, and cry1. In the phyAphyBcry1triple mutant, however, the hook failed to straightenout and the cotyledons did not unfold. These resultsindicate that hook straightening and cotyledonunfolding are controlled in a completely redundantmanner by all three photoreceptors. Only when allthree photoreceptors are missing are these two aspectsof de-etiolation significantly compromised.

Although not shown in Figure 22.15, cotyledonexpansion in Arabidopsis appears to be controlled prin-cipally by phytochrome B and cryptochrome. In oneexperiment, cotyledon area in the phyB and cry1 mutantswas reduced by 50 percent and 64 percent, respectively.However, in the phyBcry1 double mutant, cotyledon areawas not significantly greater than in the dark-grown con-trols. This additive effect of the phyB and cry1 mutantssuggests that each photoreceptor independently controlsa distinct aspect of cotyledon expansion.

The results described here represent only a briefsampling of the many interactions that occur betweenthe several photoreceptors, but one thing is clear.

Redundancy, or the possibility of evoking similareffects through different photoreceptors, is commonto light-induced phenomena. As well, numerousexamples of synergism and antagonism betweenphotoreceptors have been demonstrated. Whethersuch interrelationships are the results of parallel,independent signal pathways or of extensive crosstalkbetween pathways remains to be seen. Nevertheless,plants have clearly evolved a versatile system of multiplephotoreceptors that allows them to respond efficientlyand flexibly to dynamic changes in fluence rate andcomposition of light in their environment.

SUMMARY

The light in a plant’s environment contains a signifi-cant amount of information and the use ofthat information by plants, called photomorphogenesis,is a central theme in plant development. In orderto acquire the information provided by light, plantshave developed a sophisticated array of photorecep-tors and signal transduction pathways. There are threemajor classes of photoreceptors: the phytochromes thatrespond principally to red and far-red light, the cryp-tochromes that respond to blue light, and phototropin,which also responds to blue light.

Phytochrome is a family of photoreceptors; five areknown in Arabidopsis (phyA–phyE). They are chromo-proteins with a chromophore that consists of an lineartetrapyrrole similar to phycocyanin. Phytochrome invivo is a dimer of a 125 kDa polypeptide chain. Thehallmark of phytochrome action is photoreversibility:irradiation with red light converts the red-absorbingform of the pigment (Pr) to the far-red-absorbingform (Pfr). The physical presence of phytochrome wasestablished by demonstrating the predicted photore-versible absorbency changes in vivo. PhyA accumulatesin dark-grown seedlings in the Pr form, which is sta-ble. PhyAfr is unstable and is destroyed with a half-lifeof 1 to 1.5 hours. PhyB is expressed at low levels inboth light and dark. PhyBfr is light stable. A mixtureof red and far-red light (or white light) will establisha photoequilibrium mixture of Pr and Pfr. Pfr is thephysiologically active form.

Phytochrome-mediated effects are convenientlygrouped into three categories on the basis of theirenergy requirements: very low fluence responses(VLFR), low fluence responses (LFR), and high irradi-ance reactions (HIR). LFRs include the classicallyphotoreversible phytochrome responses such as seedgermination and de-etiolation. LFRs conveyinformation to the seed about its position relativeto the soil surface and maximize the potential for aseedling to become established in light and initiatephotosynthesis before the nutrient reserves of the

Further Reading 389

seedling are exhausted. VLFRs are not photoreversibleand are difficult to study because they saturate at lightlevels below those that cause a measurable conversionof Pr to Pfr. HIRs require prolonged exposure tohigh irradiance, are time dependent, and are notphotoreversible.

Under natural conditions, the phytochrome pho-toequilibrium value (Pfr/P) is related to the red tofar-red fluence rates. It is likely that phyB is the sen-sor that detects changes in red, far-red fluence ratiothat occur under canopies and as end-of-day signal. Inthis way, phytochrome mediates the shade avoidancesyndrome, provides a plant with information aboutthe proximity of its neighbors, and contributes to thetime-sensing mechanism.

Cryptochrome is also a chromoprotein. Cryp-tochrome has two chromophores—FAD and apterin—and bears similarity to the microbial DNArepair enzyme, photolyase. Cryptochrome mediateshypocotyl elongation, cotyledon expansion, and settingthe biological clock. Cryptochrome also has a role indetermining time of flowering.

The study of photoreceptor signal transductionis in its infancy, although some significant advanceshave been made. Phytochrome appears to operatein two modes. Phytochrome has serine-threoninekinase activity and at least two signaling partners areknown. PKS1 and NDPK2 are both phosphorylatedby phyA in vitro. phyA can also phosphorylate cry1,which may help to explain the extensive interactionsbetween phytochrome and cryptochrome in regulat-ing de-etiolation. Phytochrome also regulates geneaction through the translocation of phyB into thenucleus where it has a role in activating or suppress-ing transcription. The mode of action of cryptochromeis unknown, but its similarity to photolyase suggeststhat cryptochrome may have a redox function. Cryp-tochrome is also readily phosphorylated, which may besignificant in the signal transduction chain.

CHAPTER REVIEW

1. What unique character distinguishes phy-tochrome from all other plant pigments?

2. How is phytochrome uniquely suited tomonitor the natural light environment?

3. Compare and contrast the structures of phy-tochrome and cryptochrome.

4. Distinguish between low fluence responses andhigh irradiance responses.

5. Irradiation of phytochrome establishes aphotoequilibrium between Pr and Pfr. Canyou think of a situation in which only oneform of the pigment would be present?

6. What is a chromoprotein?7. What is meant by the statement that cryp-

tochrome is ‘‘homologous’’ with photolyase?What is the significance, if any, of this homology?

8. Two plant physiologists, H. Mohr andW. Shropshire, once wrote that ‘‘Normaldevelopment in higher plants is photomorphogenesis.’’What do you think they meant by this statement?

9. How does the experiment described in section 22.7demonstrate an interaction between phytochromeand cryptochrome in control of de-etiolation?

FURTHER READING

Banerjee, R., A. Batschauer. 2005. Plant blue-light receptors.Planta 220:498–502.

Briggs, W. R., M. Olney. 2001. Photoreceptors in plantphotomorphogenesis to date. Five phytochromes, twocryptochromes, one phototropin and one superchrome.Plant Phytisology 125:85–88.

Chen, M., J. Chorey, C. Frankhauser. 2004 Light signaltransduction in higher plants. Annual Review of Genetics38:87–117.

Kim, J-I., J-E. Park, X. Zarate, P-S. Song. 2005. Phy-tochrome phosphorylation in plant light signaling.Photochemical & Photobiological Sciences 4:681–687.

Lin, C. 2002. Blue light receptors and signal transduction.The Plant Cell. Supplement 2002:S207–S225.

Lin, C., D. Shalitin. 2003. Cryptochrome structure and signaltransduction. Annual Review of Plant Biology 54:469–496.

Lorrain, S., T. Genoud, C. Frankhauser. 2006. Let therebe light in the nucleus! Current Opinion in Plant Biology9:509–514.

Sage, L. C. 1992. Pigment of the Imagination: A History of Phy-tochrome Research. New York: Academic Press.

Shen, H., J. Moon, E. Huq. 2005. PIF1 is regulated bylight-mediated degradation through the ubiquitin-26Sproteosome pathway to optimize photomorphogen-esis of seedlings in Arabidopsis. The Plant Journal44:1023–1035.

Vandenbussche, F. et al. 2005. Reaching out of the shade.Current Opinion in Plant Biology 8:462–468.

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Ca2+ Ca2+

H+

NN

H+

PERCEPTION TRANSDUCTION TRANSMISSION RESPONSE

23Tropisms and Nastic Movements:

Orienting Plants in Space

The power of movement is not normally associated withplants. Yet movement pervades the life of the greenplant. Movement in higher plants does not involve loco-motion as it does in animals, nor is it so obvious. Plantmovement is mostly slow and deliberate, but it is akey factor in determining the orientation of plants inspace. Plants that have been inadvertently placed in thehorizontal position will reorient their root and shootto the vertical. House plants will bend, appearing toseek light coming through a window. Leaves may peri-odically rise and fall throughout the day and night,while others track the sun as it moves across the sky.Leaves of the Venus flytrap snap closed on a haplessinsect. While most plant movements are relatively slow,they nonetheless serve important functions by position-ing organs for the uptake of nutrients and water andoptimal interception of sunlight, or (in the case of theflytrap) obtaining nutrients such as nitrogen throughthe leaves.

There are two principal categories of movement inplants, based on the distinctiveness of their mechanisms.Growth movements are irreversible. They arise asthe result of differential growth within an organ orbetween two different organs. Turgor movementsare reversible, resulting from simple volume changes incertain cells—most often in a special organ called the

pulvinus. Within each group, we can further distinguishbetween nutation, tropism, and nastic movement.The term nutation (or circumnutations) denotes aregular rotary or helical movement of plant organs, mosttypically the stem apex, in space. Nutations are best de-monstrated by time-lapse photography. Tropic respon-ses are directionally related to the stimulus suchas light (phototropism), gravity (gravitropism), water(hydrotropism), or touch (thigmotropism). Nastic respon-ses are not obviously related to any vector in the stimu-lus. Directionality of nastic responses is inherent in thetissue and includes epinasty (bending down), hyponasty(bending up), nyctinasty (the rhythmic sleep movementsof leaves), seismonasty (response to mechanical shock),thermonasty (temperature), and thigmonasty (touch).

This chapter will focus on the three plant move-ments that have been explored most thoroughly. Theseare

• phototropism, particularly the nature of the pho-toreceptor and the role of auxin in the signal trans-duction chain,

• gravitropism, including a brief discussion of thenature of the gravitational stimulus and the mecha-nism of gravity perception, the particular characterof gravitropism in shoots and roots, and the role

391

392 Chapter 23 / Tropisms and Nastic Movements: Orienting Plants in Space

of auxin and calcium in the differential growth res-ponse, and

• nastic movements, including the structure of motororgans and the role of potassium flux in nyctinasticand seismonastic movements.

23.1 PHOTOTROPISM:REACHING FOR THE SUN

Most people are familiar with the sight of house plantsbending toward the light from an open window, aneveryday example of the phenomenon called pho-totropism (Figure 23.1). Phototropism is a classic plantphysiology problem that has attracted the interest ofbotanists since the middle of the nineteenth century.Darwin’s study of phototropism, published in his bookThe Power of Movement in Plants in 1881, is creditedwith overcoming the preoccupation of English-speakingbotanists with descriptive and taxonomic biology andstimulating an interest in the more dynamic aspects ofplant function. Cell elongation in phototropically stim-ulated grass coleoptiles also led to Went’s discovery ofplant hormones (Chapter 18).

Tropic responses may be either positive or negative.If a plant responds in the direction of the stimulus (e.g.,toward a light source) it is said to be positive. If itgrows away from the stimulus it is said to be negative.Whether the phototropic response is positive or negativedepends largely on the nature of the organ or its age.For example, coleoptiles, hypocotyls, and the elongatingportions of stems and other aerial organs are for themost part positively phototropic while the tendrils ofmost climbing plants are negatively phototropic. Leavesare normally plagiotropic, which means they orient at

FIGURE 23.1 The phototropic response in oat (Avenasativa) coleoptiles. Left: Dark-grown seedling placedin unilateral blue light, from the right, for 90 minutes.Right: Unlighted control. Note that the overall lengthof the coleoptile is approximately the same in the twoseedlings.

angles intermediate to the light. Roots, on the otherhand, are largely nonphototropic, although some mayexhibit a weakly negative response. The stems of ivy(Hedera helix) are negatively phototropic during theshade-loving juvenile stage, but older branches becomepositively phototropic. The stems of ivy-leafed toadflax (Cymbalaria muralis) become negatively phototropicfollowing fertilization. This interesting behavior helpsto place ripening seed pods into crevices in the walls onwhich the plant is normally found.

23.1.1 PHOTOTROPISM IS A RESPONSETO A LIGHT GRADIENT

Phototropism is often defined as a response to unilaterallight, and so it is in the laboratory. Under normal growthconditions, however, the bending response will occureven in plants that are receiving light from all sides.All that is required is that the fluence rate be unequallydistributed. In experiments with bilaterally illuminatedgrass coleoptiles, for example, as little as 20 percentdifference in fluence rate on the two sides of the organwill induce a bending response (Figure 23.2). Thus lightcan be presented unilaterally (as it is in most laboratoryexperiments), bilaterally, from all sides, and even fromabove, providing only that a gradient is created acrossthe organ. Phototropism is thus a growth response to alight gradient.

The magnitude of a light gradient across an organsuch as a coleoptile is dependent on optical propertiesof the tissue as well as differences in incident light.A light gradient across an organ, for example, can be

FIGURE 23.2 A phototropic response will occur wheneverthere is a light gradient established across the stem orcoleoptile axis. The upper part of the figure representsunilateral illumination, which is normal in experimentalsituations. The lower part of the figure illustrates bilat-eral illumination, with the highest fluence rate from theright. Phototropic curvature is the same in either case.

23.1 Phototropism: Reaching for the Sun 393

intensified by screening within the organ. Pigments,including but not limited to the photoreceptor itself,will attenuate the light as it passes through the organ.Light can also be attenuated by scattering, reflection,or diffraction within the cells or as it passes betweencells. Thus gradients across individual cells, measuredby using microfiberoptic probes, may vary from 5:1to 50:1. To further complicate matters, organs suchas coleoptiles appear to function as light pipes. Thismeans that light applied to the tip, for example, willbe transmitted through the coleoptile to cells fur-ther down the organ. Thus the phototropic stimulusis far from being a simple matter. These complexinteractions between light and the optical propertiesof tissue have led to significant difficulties in experimen-tal design as well as in interpretation of the resultingdata.

23.1.2 PHOTOTROPISM IS ABLUE-LIGHT RESPONSE

Since the 1930s, action spectra for phototropism havebeen repeatedly determined for a number of organisms,but have been most thoroughly documented for coleop-tiles of oats (Avena sativa) and maize (Zea mays) and forsporangiophores of the fungus Phycomyces. The actionspectra for oat coleoptiles and Phycomyces are virtuallyidentical, indicating that they share a homologous, if notcommon, photoreceptor. All other phototropic actionspectra are similar and consistently show two peaks inthe blue region of the spectrum near 475 nm and 450 nmand a small peak or shoulder at 420 nm (Figure 23.3).In addition there is a broad action peak in the UV-Aregion near 370 nm. The action spectra for both oat andPhycomyces phototropism show an additional peak in theregion of 280 nm, indicating that the photoreceptor isprobably a chromoprotein.

As early as the 1940s, it was suggested that thephotoreceptor could be a flavin molecule such as ribo-flavin. However, because of its action spectrum the pho-toreceptor for phototropism was subject to the sameflavin-carotenoid controversy described in the previouschapter for other blue-light responses now knownto be regulated by the cryptochromes. On the otherhand, there were several physiological results thatruled out carotenoids as the photoreceptor for pho-totropism long before the responsible pigment, photo-tropin, was finally discovered and shown to be aflavo-protein. Carotenoid biosynthesis, for example,can be blocked, either by mutation or by treatmentof seedlings with the herbicide norflurazon, whichinhibits the enzyme phytoene desaturase. Yetcarotenoid-deficient maize mutants, albino barleyseedlings, and norflurazon-treated seedlings all exhibita normal phototropic response to blue light.

Rel

ativ

e ab

sorp

tion

or

resp

onse

(A)

350 400 450 500

(B)

(C)

Wavelength (nm)

FIGURE 23.3 (A) The action spectrum for Avena coleop-tile phototropism. The action spectrum shows peakactivity in the blue and UV-A regions of the spectrum.For comparison, the absorption spectra for riboflavin, acommon flavonoid (B), and β-carotene (C) are shown.Although β-carotene ‘‘fits’’ in the blue region (around450 nm), riboflavin has the required absorption in boththe blue and the UV-A regions (320 nm–400 nm) of thespectrum.

23.1.3 PHOTOTROPISM ORIENTSA PLANT FOR OPTIMALPHOTOSYNTHESIS

The phototropic blue-light response is distinct fromthe blue-light responses mediated by phytochromeand cryptochrome that were discussed in the previouschapter. Phytochrome and cryptochrome responsesare morphogenetic responses—they alter the patternof growth and development. The singular impact ofphototropism, on the other hand, is that it orientsgrowth and leaf angle toward incident light in orderto maximize light interception for photosynthesis. Thebending of coleoptiles and hypocotyls is only the mostvisible part of a larger blue-light syndrome that plantsuse to optimize photosynthesis. Plants also use blue lightto control stomatal opening and facilitate gas exchangeas well as to relocate chloroplasts within the cell.

It has long been known that stomatal opening isunder the control of light. On the one hand, lightabsorbed by chlorophyll (i.e., red light) stimulates stom-atal opening and obviously depends on photosyntheticreactions in the guard cell chloroplasts. However, thereis a second, much more sensitive, system that is drivenby low levels of blue light. Most of the evidence pointsto a dominant role of the blue-light response in the early

394 Chapter 23 / Tropisms and Nastic Movements: Orienting Plants in Space

phases of stomatal opening, such as when the stomataopen at dawn, prior to the beginning of photosynthesis.

Plants also use blue light to control the high-lightavoidance response of chloroplasts in the mesophyllcells. In low light, the chloroplasts always gather alongthe cell walls that are parallel to the surface, (i.e., peri-clinal walls) that are perpendicular to the incident light(Figure 23.4). In high light, such as direct sunlight,the chloroplasts avoid potential damage by lining upalong the anticlinal walls (i.e., parallel to the incidentlight). The redistribution of chloroplasts appears to bein response to a light gradient through the cytoplasm,so the responsible photoreceptor is probably locatedin the cytoplasm, not the chloroplasts. The mecha-nism of redistribution has yet to be discovered, but thecytoskeleton is commonly involved in moving organelleswithin the cell and may be involved in the movement ofchloroplasts as well.

23.1.4 FLUENCE RESPONSE CURVESILLUSTRATE THE COMPLEXITYOF PHOTOTROPIC RESPONSES

Perhaps no aspect of phototropism has indicated thecomplexity of the process so much as attempts to definerelationships between fluence and response. Phototro-

pism is characterized by a rather curious fluence-response curve, quite unlike most photobiologicalresponses.

Fluence response curves are generally obtainedby monitoring the response of the organ to differ-ent total amounts of light (fluence), usually by using asingle fluence rate but varying the presentation time.Figure 23.5 shows a fluence-response curve determinedfor Avena coleoptile phototropism that illustrates theclassic response to increasing fluence. There is an initialrise to a first peak, which is called first positive curva-ture. With increasing fluence, curvature declines, to thepoint that this may even result in a bending away fromthe light source. This decline and negative response iscalled first negative curvature. Note that first negativecurvature is not necessarily ‘‘negative’’ in the sense ofbending away from the light. It may be simply a reducedpositive response. Following the region of first negativecurvature, the response curve again rises into what iscalled second positive curvature. In some cases, a sec-ond negative and even a third positive curvature havebeen reported. First positive curvature is also knownas tip curvature, because it is restricted to the apex ofcoleoptiles. Second positive curvature is also called basalcurvature because the curvature extends more towardthe basal region of the coleoptile.

B

A

FIGURE 23.4 The high-light avoidance response of chloroplasts in a typical meso-phyll cell. In low light conditions (left), the chloroplasts gather perpendicular to theincident light along the upper and lower (periclinal) walls in order to maximize lightinterception. In high-light conditions (right) the chloroplasts gather along the side(anticlinal) walls, or parallel to the incident light, in order to avoid damage due toexcessive light. Mesophyll cells are shown in cross-section (A) and surface view (B).

23.1 Phototropism: Reaching for the Sun 395

Deg

rees

cur

vatu

re

–13

0

–12 –11 –10 –9 –8

12

24

36

Log fluence (mol photons cm–2)

1st Positive

2nd Positive

1st Negative

FIGURE 23.5 A phototropic fluence-response curve forAvena coleoptiles. First positive, first negative, and sec-ond positive curvatures are indicated.

There is a fundamental law of photochemistry,called the Bunsen-Roscoe reciprocity law, which saysthe product of a photochemical reaction is determinedby the total amount of energy presented, regardless offluence rate or presentation time. In other words, thesame result is obtained with either a brief exposureto a high fluence rate or a longer exposure to a lowfluence rate. Numerous experiments have establishedthat the reciprocity law applies to first positive cur-vature but does not apply to first negative or secondpositive curvatures. Second positive curvature is insteadvery time-dependent. In other words, second positivecurvature is more dependent on presentation time thanon fluence rate. Failure of reciprocity for second pos-itive curvature suggests the possibility that more thanone photoreceptor might be involved. However, actionspectra have been determined for both first and secondpositive curvature and they are identical. Apparentlyboth first and second positive curvature are mediated bythe same photoreceptor and the complexities of secondpositive curvature are due to subsequent events in thesignal transduction chain.

23.1.5 THE PHOTOTROPIC RESPONSEIS ATTRIBUTED TO A LATERALREDISTRIBUTION OFDIFFUSIBLE AUXIN

At the same time F. W. Went and his contemporarieshad chosen to study the influence of the apex on coleop-tile elongation, parallel studies on the role of the rootapex were being conducted in Germany by N. Cholodny.The result was independent proposals by Cholodny, in1924, and Went, in 1926, that the apex was able toinfluence cell elongation in the more basipetal extensionregion of the organ. These ideas of these 2 investigators

were drawn together in the late 1920s in an attempt toexplain phototropism. The Cholodny-Went hypoth-esis states that unilateral illumination induces a lateralredistribution of endogenous auxin near the apex of theorgan. This asymmetry in auxin distribution is main-tained as the auxin is transported longitudinally towardthe base of the organ. The higher concentration of auxinon the shaded side of the organ stimulates those cellsto elongate more than those on the lighted side. It isthis differential growth that causes curvature toward thelight source.

The experimental basis for the Cholodny-Wenthypothesis is derived largely from agar-diffusion exper-iments originally conducted by Went and describedearlier in Chapter18. In Went’s experiments, oat coleop-tiles were first stimulated with unilateral light. Thecoleoptile apices were then excised, split longitudinally,and the two halves placed on agar blocks in order to col-lect the auxin that diffused out of the base. The amountof auxin collected in the agar blocks was then assayed bythe Avena curvature test (Chapter 18). Went reportedthat a significantly higher quantity of auxin was collectedfrom the shaded half of the coleoptile apex than from thelighted half, indicating that unilateral lighting caused agreater proportion of the auxin to be transported downthe shaded side of the coleoptile.

Doubts as to the validity of the Cholodny-Wenthypothesis arose from numerous unsuccessful attemptsto verify asymmetric auxin distribution by applying14C-labeled IAA to tropically stimulated coleoptiles.These problems, however, may be largely attributed topoor experimental technique. It is now evident that alarge proportion of the radioactive auxin taken up bythe tissue in those experiments did not enter the auxintransport stream. When care is taken to discount thisnondiffusible auxin, a clear differential in auxin transportcan be detected. For example, when maize coleoptile tipswere supplied with 14C-IAA, approximately 65 percentof the radioactivity was recovered from the shaded side.There was no significant asymmetry when subapicalsections were used, further evidence that the lateralredistribution of auxin occurs at the very apex of thecoleoptile.

In the 1960s, the Cholodny-Went hypothesis wassystematically reevaluated by W. R. Briggs and his col-leagues. Briggs repeated Went’s original split-tip exper-iments but, unlike Went, he excised the tips and placedthem on agar blocks before presenting the phototropicstimulus. The results (Figure 23.6) clearly demonstratethat when the tip is partially split, leaving tissue conti-nuity only at the very apex of the coleoptile, exposureto unilateral light causes an increase in the amount ofdiffusible auxin on the shaded side and a decrease onthe lighted side. The total amount of auxin recovered,however, remains effectively constant. When lateraldiffusion of auxin is prevented throughout the entire

396 Chapter 23 / Tropisms and Nastic Movements: Orienting Plants in Space

DARK CONTROLIntact tip

ILLUMINATEDIntact tip

ILLUMINATEDPartially split tip

ILLUMINATEDTotally split tip

A.

B.

C.

D.

25.6

30.7 16.2

22.3 23.0

25.8

hν

hν

hν

FIGURE 23.6 Phototropic stimulation establishes anasymmetric distribution of diffusible auxin in excised Zeamays coleoptile apices. (A, B) Intact control apices. A wasmaintained in darkness and B was provided light unilat-erally from the right. (C) Tips were partially split, leavingtissue continuity only at the very apex. A microscopecover slip was inserted to provide a barrier to lateraldiffusion. The tips were then presented with unilaterallight from the right. (D) Tips were totally split and thediffusion barrier passed through the apex before beingpresented with unilateral light from the right. Numbersindicate the amount of auxin collected in the agar blocksover a 3-hour period, based on degrees of curvature inthe Avena curvature bioassay. Values are for auxin col-lected from 3 tips (A, B) or 6 half-tips (C, D). (Data fromBriggs, W. R. 1963. Plant Physiology 38:237.)

length of the tip, no such asymmetric auxin distributionis observed. These results clearly support the princi-pal tenet of the Cholodny-Went hypothesis, namely,that unilateral light induces a preferential migration ofauxin down the shaded side of the coleoptile. Theirexperiments also confirmed that auxin production incoleoptiles of Zea mays is confined to the apical 1 to 2 mmand that lateral redistribution during phototropic stim-ulation probably occurs within the most apical one-halfmm.

Compelling support for the Cholodny-Went the-ory has been provided by a more recent study of auxinredistribution in Brassica oleraceae hypocotyls. The freeIAA concentration found on the shaded side of thehypocotyl was found to be at least 20 percent higherthan on the lighted side following phototropic stimula-tion. Moreover, the differential auxin concentration wasaccompanied by a several-fold increase in the expres-sion of auxin-regulated genes on the shaded side of thehypocotyls, including two members of the α-expansinfamily of genes that are necessary for cell wall extension(see Chapter 17). Finally, both the auxin differential and

the differential in auxin-regulated gene expression couldbe detected before there was any noticeable curvature ofthe hypocotyls.

23.1.6 PHOTOTROPISM AND RELATEDRESPONSES ARE REGULATED BYA FAMILY OF BLUE-SENSITIVEFLAVOPROTEINS

Two lines of study led to the discovery of the photore-ceptor for phototropism, now called phototropin. Inthe late 1980s, it was reported that blue light stimulatedthe phosphorylation of a 120 kDa plasma membraneprotein localized in the actively growing regions of etio-lated pea seedlings. This is the same region that is mostresponsive to the phototropic stimulus. After extensivebiochemical and physiological characterization, the pro-tein was found to be a kinase that autophosphorylates inblue light. There was also a strong suggestion that thiskinase was the photoreceptor for phototropism.

A short time later, a mutant characterized by afailure to respond to the phototropic stimulus (non-phototropic hypocotyl 1, or nph1) was isolated fromArabidopsis. Plants carrying the nph1 mutant not onlyfailed to exhibit phototropism, but coincidentally lackedthe 120 kDa membrane protein. When the NPH1 genewas cloned, it was found, as expected, to encode the120 kDa protein. The NPH1 holoprotein was subse-quently renamed phototropin 1 (phot1) because of itsfunctional role in phototropism.

Phototropin 1 is a flavoprotein with two flavinmononucleotide (FMN) chromophores (Figure 23.7). Ithas a carboxy-terminal domain with the characteristicsof a serine-threonine kinase. The photosensory domainat the N-terminus has two distinctive domains calledLOV domains, so named because they share charac-teristics with microbial proteins that regulate responsesto light, oxygen, or voltage. Not surprisingly, the twoLOV domains are the two sites that bind FMN and makephototropin responsive to light. A second phototropin,phototropin 2 (phot2), has since been discovered. Onthe basis of amino acid sequence, the two phototropinsare about 60 % similar, but the two LOV domains arevirtually identical and each phototropin binds two FMNchromophores.1

The two phototropins found in Arabidopsis, phot1and phot2, exhibit some overlapping functions. Eachalso appears to have unique physiological roles. Firstpositive curvature appears to be mediated solely by

1In 2001, the gene for photo2 was originally described asNPL1 (NPH-like1), a homolog of the gene NPH1 (non-phototropic hypocotyl 1) or PHOT1. NPL1 has since beenrenamed PHOT2 to bring the nomenclature in line withphototropin 1.

23.1 Phototropism: Reaching for the Sun 397

PHOTOTROPIN

N

N

NEOCHROME

LOV 1

LOV 1

LOV 2

LOV 2 Ser-Thr Kinase C

CSer-Thr Kinase

FIGURE 23.7 The domain structuresof phototropin and neochrome. Pho-toropin contains two LOV domainsthat are characteristic of proteinsactivated by light, oxygen, or volt-age. Each LOV domain carriesone flavin mononucleotide chro-mophore that absorbs in the blueregion of the spectrum. At thecarboxy-terminal end of the pro-tein (–C) is a serine-threonine kinasedomain. Note that neochrome isvirtually identical to phototropinexcept that it has a phytochrome pho-tosensory domain at the N-termi-nal end. Neochrome responds to bothblue and red/far-red light.

phot1, while second positive curvature is mediated byboth phot1 and phot2. Phot1 and photo2 contributeequally to stomatal opening, while the avoidance move-ment of chloroplasts under high light intensities ismediated only by phot2.

Recent evidence has also indicated a role forphototropins in the promotion of cotyledon and leafexpansion and the rhythmic sleep movements of kidneybean leaves (see Chapter 24).

23.1.7 A HYBRID RED/BLUE LIGHTPHOTORECEPTOR HAS BEENISOLATED FROM A FERN

A particularly interesting photoreceptor has recentlybeen isolated from the fern Adiantum capillus-veneris.Designated neochrome (formerly known as phy3),this photoreceptor has properties of both phytochromeand phototropin (Figure 23.7). The amino acidsequence of the amino-terminal domain shows asignificant homology to the chromophore-bindingdomain of phytochrome. Furthermore, when the genewas expressed in yeast and the purified protein wasreconstituted with a phycocyanobilin chromophore, itshowed typical phytochrome photoreversible behavior.But neochrome also has the two LOV domains, whichbind FMN, and a serine/threonine kinase domainat the C-terminus that are virtually identical tophototropin. Neochrome is required for phototropismin Adiantum, which is regulated by red as well as bluelight. Neochrome is clearly a hybrid photoreceptorthat mediates red, far-red, and blue-light responses.What is curious, however, is that Adiantum also hastwo fully functional phototropins like their higherplant counterparts and, again like their higher plantcounterparts, phot2 is solely responsible for mediatinghigh-light chloroplast avoidance movements.

Several other ferns, mosses, and algae have bothphototropins and neochromes, which raises interesting

questions regarding the evolution of photoreceptors aswell as their physiological action.

23.1.8 PHOTOTROPIN ACTIVITYAND SIGNAL CHAIN

Participants in the phototropin signal chain are onlynow just beginning to ‘‘come to light.’’ Some of themore important factors can be summarized as follows:

1. Autophosphorylation of phototropin plays a significantrole in the phototropic response, probably by initiatinga phosphorylation cascade. Studies employing mutantsof the PHOT1 gene have shown that the protein isapparently folded in such a way that the phospho-rylation site is blocked by the LOV2 domain in thedark. Absorption of blue light by the chromophoreinduces a change in the conformation of the pro-tein so that the phosphorylation site is available andactive. The role of the LOV1 domain is not clear.Mutants lacking the LOV1 domain have shown thatLOV1 is not necessary for phosphorylation but itspresence does increase kinase activity.

2. Phototropins may be involved in gene regulation. Nosubstrates directly phosphorylated by phototropinin planta have yet been identified, but there areseveral proteins that interact with the photore-ceptor and are necessary for a proper response.For example, the proteins NONPHOTOTROPICHYPOCOTYL 3 (NPH3) from Arabidopsis and ahomologous protein (called an ortholog) from rice,COLEOPTILE PHOTOTROPISM 1 (CPT1),include domains that are characteristic of transcrip-tional regulators or proteins that are involved inprotein degradation. The mutants nph3 and cpt1,in which these proteins are missing, show no pho-totropic response.

3. Phototropin disrupts polar auxin transport. One of thechallenges presented by phototropism is to establishwhether or not a link exists between the absorption

398 Chapter 23 / Tropisms and Nastic Movements: Orienting Plants in Space

of blue light by phototropin and the asymmetricalauxin distribution proposed by the Cholodny-Wenthypothesis. As described previously, asymmetricauxin distribution has been demonstrated experi-mentally. Recent experiments have focused on therelationship between phototropin and auxin effluxfacilitator PIN1. PIN1 is normally localized at thebasal ends of xylem-associated cells where itserves to facilitate the polar vertical flow of auxin(Chapter 18). When the location of PIN1 in Arabi-dopsis hypocotyls was monitored by immunofluore-scence microscopy following phototropic stimulus,the basal location of PIN1 in wildtype plantswas disrupted in the cortical cells on the shadedside of the hypocotyl. A similar disruption wasnot observed in phot1 mutants. These resultssuggest that phototropic bending is initiated bya phototropin-mediated decrease in the verticaltransport of auxin. This would lead to a retentionor sequestering of auxin, and consequent increasedgrowth, in those cells that are directly involved inphototropic bending.

23.1.9 PHOTOTROPISM IN GREENPLANTS IS NOT WELLUNDERSTOOD

A final area of concern is phototropism in light-grownplants, where relatively little is known about thephototropic process. As with phytochrome, discussed inChapter 22, most of what we know about phototropismis derived from laboratory studies with etiolated seed-lings. However, in light-grown cucumber (Cucumissativus) and sunflower (Helianthus annuus) seedlingssubjected to uniform lighting, curvature of thestem can be induced by simply shading one of the

FIGURE 23.8 Curvature in cucumber (Cucumis sativus)seedlings induced by shading cotyledons. The left-handcotyledon was covered with aluminum foil and theseedling uniformly irradiated with white light for 8hours. (After Shuttleworth, J. E., M. Black. 1977. Planta135:51.)

cotyledons (Figure 23.8), and the phototropic responseof sunflower seedlings is markedly decreased if theleaves are removed. The cucumber response, at least,differs from the classical phototropic response in thatit is induced by red light rather than blue light. Thisappears to be a phytochrome-mediated response and isrelated to inhibition of hypocotyl elongation below theirradiated cotyledon. Both the cucumber and sunflowerresponses may be attributed to the fact that the leavesare a prime source of auxin required for the growthresponse. Both cucumber and white mustard (Sinapisalba) also exhibit a classical phototropic responseinduced by irradiating the hypocotyls directly with bluelight. Clearly the control of stem growth in green plantsis an area where there is still much to learn.

23.2 GRAVITROPISM

Gravitropism is probably one of the most unfailinglyobvious and familiar plant phenomena to most people(Figure 23.9). Everyone is aware that shoots alwaysgrow ‘‘up’’ and roots always grow ‘‘down.’’ Or do they?A casual walk through the woods or garden shouldreveal how overly simplified this view is. The lateralbranches of most trees and shrubs do not grow up; theygrow outward in a more or less horizontal position.Stolons (or runners) of strawberry (Fragaria) plants andbuttercups (Ranunculus) also grow horizontally alongthe soil surface. Dig into the soil and you will findrhizomes (underground stems) and many roots growinghorizontally. Many pendulous inflorescences show nodirectional preference for growth, but hang down simplyof their own weight.

FIGURE 23.9 Gravitropism in maize (Zea mays) seedlings.Four-day-old dark-grown seedlings were placed in thehorizontal position for 3 hours. Note the shoot exhibitsnegative gravitropism and the root exhibits positive grav-itropism.

23.2 Gravitropism 399

Unlike most other environmental stimuli, the forceof gravity is omnipresent and nonvarying. It does notvary in magnitude as temperature does, for example.Gravity cannot be turned on and off, such as light atdawn and dusk. Moreover, gravity is not a unilateralstimulus—there is no gradient component in gravity.Cells on the lower side of a stem or root are subjected tothe same gravitational force as those on the upper side.Consequently, it is likely that gravity can be detectedonly by the movement of some structure or structureswithin the cells—a movement that establishes an initialasymmetry in the cell and is translated in terms ofpressure. The mass and movement of whatever structureis involved must be consistent with the sensitivity andspeed of the gravitational response and there must bea mechanism for transducing the pressure signal into abiochemical signal that can lead to a differential growthresponse.

23.2.1 GRAVITROPISM IS MORE THANSIMPLY UP AND DOWN

It is true that the root and shoot of the primary plantaxis do align themselves parallel with the directionof gravitational pull. Such an alignment is said tobe orthogravitropic. The primary root, which growstoward the center of the earth, exhibits positive gravit-ropism. The shoot, which grows away from the centerof the earth, exhibits negative gravitropism. Organssuch as stolons, rhizomes, and some lateral branches,which grow at right angles to the pull of gravity, aresaid to be diagravitropic. Organs oriented at someintermediate angle (between 0◦ and 90◦ to the vertical)are said to be plagiogravitropic. Lateral stems and lat-eral roots are commonly plagiogravitropic. Organs thatexhibit little or no sensitivity to gravity are said to beagravitropic.

The advantages to the plant of positive and nega-tive gravitropic growth responses are fairly obvious.Seeds may assume a random orientation in the soil, butin order to ensure survival, the shoot, with its photo-synthetic structures, must be above ground in orderto take advantage of sunlight. The root system mustremain in the soil in order to secure anchorage and areliable supply of nutrients and water. The primary rootmost often exhibits a strongly positive orthogravitropicresponse. Secondary roots (i.e., first-level branchroots), however, tend to grow more horizontallywhile tertiary roots are generally agravitropic. Thishierarchy of gravitational responses ensures that theroot system more effectively fills the available spaceand thus more efficiently mines the soil of waterand nutrients (Figure 23.10). In a similar fashion, ahierarchy of negative orthogravitropic, diagravitropic,and plagiogravitropic responses in the shoot system

Principal Shoot Axis(negative orthogravitropic)

Branches(plagiogravitropic)

(diagravitropic)

Soil level

Rhizome(diagravitropic)

Tertiary Roots(plagiogravitropic,or agravitropic)

Secondary Roots(plagiogravitropic)

Primary Root(positive orthogravitropic)

FIGURE 23.10 Diagram illustrating the range of gravit-ropic responses in shoots and roots. Note that differencesin the gravitropic behavior among different levels of bothshoot and root branches ensures the plant fills space.

helps to reduce mutual shading and ensures a moreefficient capture of sunlight to drive photosynthesis.

23.2.2 THE GRAVITATIONAL STIMULUSIS THE PRODUCT OF INTENSITYAND TIME

Gravitational stimulation (stimulus quantity or dose) isthe product of the intensity of the stimulus and the timeover which the stimulus is applied:

d = t · a

where a is the acceleration of mass due to gravity(in g), t is the time (in seconds) over which the stim-ulus is applied, and d is the dose in g seconds (g z s)(see Box 23.1: Methods in the Study of Gravitropism).The minimum dose required to induce gravitropic cur-vature is called the threshold dose. Threshold dose willvary depending on the organism or experimental con-ditions. Values of d in the range of 240 g · s (at 22.5◦C)to 120 g z s (at 27.5◦C) have been reported for Avenacoleoptiles, but more careful mathematical analyses sug-gest that less than 30 g · s (i.e., an acceleration of 1 g forless than 30 seconds) is sufficient to induce gravitropiccurvature in roots. Three other parameters are of inter-est when defining gravistimulation: presentation time,reaction time, and threshold intensity.

400 Chapter 23 / Tropisms and Nastic Movements: Orienting Plants in Space

S1S1

CI CI CI CI

QcQcQcQc

S2

S3

S3 S3 S3

S2S2S2

S1 S1

BOX 23.1METHODS IN THESTUDY OFGRAVITROPISM

A fundamental requirement for any form of scientificexperimentation is that of controlling the application ofthe stimulus in the form of intensity (or concentration)and duration. Since the gravitational field of earth cannotbe extinguished (except in the microgravity conditionsof space), experimentation on gravitational effects onplants and other organisms has required some uniqueapproaches.

Most experiments require a mass acceleration in therange of 1 g or less, which can easily be achieved bysimply orienting the organ (e.g., a coleoptile or primaryroot) away from the vertical. The force (at least forshort-term stimulation) is generally proportional to thesine of the angular deviation from the vertical. Thusthe force is greatest in the horizontal position since sine90◦ = 1 (the sine of angles less than 90◦ is less than 1).Seedling shoots generally must be oriented between 0.5◦

to 10◦ from the vertical in order to induce curvature.Forces greater than 1 g can be achieved by the use

of specially designed centrifuges. Similar centrifugeshave been used earlier in the century for studying theproperties of animal (egg) membranes, and so forth.Centrifugation has not been used extensively in thestudy of gravitropisn, but those experiments in which ithas been used have provided some useful insights.

The problem of extinguishing gravitational forceshas been approached in two ways: clinostats and spaceflight. A clinostat is a device that holds the plant axis in ahorizontal position while continuously rotating it about

the horizontal axis. This does not actually extinguish thegravitational field, of course, but the summated effect isa nondirected constant stimulation. With the clinostat,plants can first be subjected to a brief stimulus and thenrotated to, in effect, remove any further stimulus. Asmight be expected, continuous multilateral stimulationhas been found to influence a variety of physiologicalparameters. Avena seedlings, for example, respond withincreased growth rate and increased respiration. Someof these changes may be incidental, but others mayinfluence the gravitropic response. Results must alwaysbe interpreted with caution.

SPACE—THE FINAL FRONTIER

The advent of space flight in the 1950s has pro-vided plant scientists with unique opportunities to studyresponses to microgravity conditions. Since 1960, whenthe first wheat and maize seeds were carried aloft onSputnik 4, experiments with plants have been con-ducted on manned and unmanned spacecraft from boththe United States and the Soviet Union (now Rus-sia). Perhaps not unexpectedly, physiological effectsof microgravity are not limited to gravitropism butembrace a variety of other cellular events. Many ofthe effects are deleterious, including reduced growth,chromosomal aberrations, and other cytological abnor-malities. Death at the flowering stage was commonuntil, in 1982, Arabidopsis thaliana were successfully car-ried through a complete life cycle and produced viableseed. Many of the difficulties could be attributed simplyto the logistics of trying to maintain plants in space, buteven with improved methods, difficulties are still beingencountered. As yet the returns may be modest, but theuse of microgravity as an experimental tool is ripe forexploitation.

The minimum duration of stimulation required toinduce a curvature that is just detectable is known as thepresentation time. The intensity of stimulation shouldalso be defined, although a stimulus of 1 g at 90◦ is moreor less standard. A force of 1 g is easily obtained by simplyplacing the stem or root in a horizontal position. Presen-tation times of 12 seconds for cress roots and 30 secondsfor Avena coleoptiles have been determined, but a brief1-second stimulus will induce curvature in Avena coleop-tiles if the stimulus is repeated every 5 seconds. Thissuggests that some cumulative receptive process beginsthe instant the plant assumes a horizontal position.

Presentation time should not be confused with reac-tion time, which is the interval between the presentationof the stimulus and the actual development of curvature.Reaction times involve the complete signal transduction

sequence that leads to the asymmetric growth response.Typically, 10 minutes is required before curvature canbe visually detected, although reaction times may varyfrom a few minutes to hours, depending on the speciesand conditions. In experiments employing sensitive elec-tronic position-sensing transducers, curvature of maizecoleoptiles could be detected within about 1.5 minutesof horizontal placement, while bending of the mesocotylcould not be detected before 3.5 minutes.

The minimum stimulus intensity required to inducea response is known as the threshold intensity. Thresh-old intensities have been determined for a variety ofplant organs under different experimental conditions.The results are remarkably consistent and indicate thatroots are perhaps an order of magnitude more sensi-tive than shoots. In land-based clinostat experiments

23.2 Gravitropism 401

Ca2+ Ca2+

H+

NN

H+

PERCEPTION TRANSDUCTION TRANSMISSION RESPONSE

FIGURE 23.11 The four phases of root gravitropism. When the orientation of a rootchanges in a gravitational field, the change is perceived in the root cap by the settlingof amyloplasts against intracellular membranes such as the endoplasmic reticulum.The biophysical signal is then converted to a biochemical signal through second mes-sengers such as hydrogen ions, calcium ions, and the relocation of auxin transportfacilitators (red circles). The signal is then transmitted to the elongation zone of theroot via an altered flow of auxin (arrows) which results in the curvature response. N= nucleus.

(see Box 23.1), values for threshold intensity for Avenacoleoptiles and roots were found to be 1.4 × 10−3 g and1.4 × 10−4 g. Values calculated for lettuce seedlinghypocotyl and roots in experiments aboard the Salyut 7spacecraft were 2.9 × 10−3 g and 1.5 × 10−4 g, respec-tively. It is apparent that many plants are very sensitiveto gravitational stimulus.

23.2.3 ROOT GRAVITROPISM OCCURSIN FOUR PHASES

Virtually all of the studies on root gravitropism havefocused on primary roots and have identified four suc-cessive phases: perception, transduction, transmission,and growth response (Figure 23.11). Although the actualtiming may vary depending on the conditions of theexperiment, the initial perception phase occurs withinperhaps one second of orienting a root off the verticaland involves biophysical mechanisms (e.g., pressure) forsensing the direction of gravitational pull. The trans-duction phase, occurring between 1 and 10 secondsfollowing reorientation, involves the conversion of thebiophysical single to a biochemical signal. The transmis-sion phase occurs between 10 seconds and 10 minutesof reorientation and involves a redistribution of auxinwithin the root tip. The growth response, due to theunequal distribution of auxin, causes curvature of theroot toward a more vertical orientation.

23.2.3.1 Gravity is perceived by the columellacells in the root cap. Gravitropic perception in theroot is localized in the root cap, a thimble-like mass ofcells that covers the tip of the root. The root cap consistsof a central core of cells (the columella) arranged inregular tiers and one or more outer layers of peripheralcells. Traditionally, the function of the root cap wasthought to be twofold; it provides physical protectionfor the root apical meristem and its peripheral cellssecrete a mucilaginous polysaccharide that lubricatesthe path of the growing root.

A third function, that of gravity perception, hasbeen established by experiments in which the root capis wholly or partially surgically removed (Figure 23.12).Complete removal of the root cap does not interfere withthe elongation of the root but completely abolishes anygravitropic response. Decapped roots will recover sen-sitivity to gravity after about 24 hours, which correlateswith the regeneration of a new cap. Surgical experimentshave also indicated that removal of the central core, orcolumella, cells caused the strongest inhibition of theresponse to gravity. Individual cells or pairs of columellacells can be selectively removed or ablated (L. ablatus, totake away) with a nitrogen laser, in conjunction with anoptical microscope. Ablation of the innermost columellacells has the greatest impact on root curvature, withoutaffecting overall growth rate of the root (Figure 23.13).Laser ablation of the root cap peripheral cells, on theother hand, has no effect on the gravitropic response.

A.

B.

FIGURE 23.12 The role of the root cap in curvature ofvertically oriented roots. (A) Control root. Growth isuniform when the root cap is left intact. (B) When theroot cap is surgically removed from one-half of theroot, the root grows toward the side with the capremaining.

402 Chapter 23 / Tropisms and Nastic Movements: Orienting Plants in Space

S1S1

CI CI CI CI

QcQcQcQc

S2

S3

S3 S3 S3

S2S2S2

S1 S1

FIGURE 23.13 Diagram of an Arabidopsis root cap show-ing the quiescent center (QC), columella initials (CI),and three ranks of columella cells. Colored cells indicatethe cells that are sensitive to gravistimulation, based onlaser ablation experiments. Relative sensitivity is indi-cated by the intensity of color. (Based on Perbal andDriss-Ecole, 2003).

23.2.3.2 Gravity perception involves displace-ment of starch-filled amyloplasts. A response togravity must almost certainly involve sedimentationof some physical structure within the cell. F. Nollwas the first to suggest, in 1892, that plants mightsense gravity in a manner similar to some animals.Crustaceans, molluscs, and many other invertebrateshave gravity-sensing organs called statocysts, smallinnervated cavities lined with sensory hairs. Withinthe cavity are one or more statoliths, tiny granules ofsand or calcium carbonate that are pulled downwardby gravity. When the statocyst changes position, thestatoliths also shift position, bending the sensory hairsand sending an action potential to inform the centralnervous system of the change.

In 1900, G. Haberlandt and E. Nemec indepen-dently adapted the statolith theory to account for plantresponses to gravity. Based on careful cytological stud-ies, they proposed the starch-statolith hypothesis inwhich starch grains found in specialized tissues func-tion as statoliths. Statocytes are cells containing sedi-mentable starch grains. Tissues that contain statocytesare known as statenchyma. Support for the statolithhypothesis was found in earlier reports by Darwin andothers that removal of the root tip, where most of thestarch grains are found, resulted in a loss of gravit-ropic response. Nonetheless, the hypothesis was notuniversally accepted and over the decades a number ofinvestigators have attempted to prove or disprove it.

A statolith is not a naked starch grain, but a groupof starch grains contained within a membrane, called an

amyloplast (Figure 23.14). There may be 1 to severalindividual grains within an amyloplast and as many as adozen amyloplasts in each statocyte. This compares withthe single large grains characteristic of starch storageorgans. Not all amyloplasts in all cells are readily mobile.In fact, detection of putative statoliths, or readily mobileamyloplasts, appears to be largely confined to regions ofhigh gravitropic sensitivity. These include the mass ofcolumella cells in the central core of the root cap and, inhypocotyls, a zone of endodermal cells that sheath thevascular tissues (also referred to as the starch sheath).Mobile amyloplasts may also be found in the innercortical cells of aerial organs and the pulvini, or motororgans in the nodes of grass stems that are responsive togravity.

Any gravity-sensing mechanism involving particlesedimentation would have to operate with a speed andsensitivity consistent with the known speed and sensitiv-ity of the response. In the 1960s, L. J. Audus undertooka careful examination of various subcellular particles.Audus concluded that, of all the cellular organelles, onlystarch grains have the mass and density to move throughthe viscous cytoplasm within known presentation times.Ultrastructural examination has shown that other cel-lular organelles, such as the endoplasmic reticulum,may become shifted in cells subjected to gravitational

Nucleus

Cell wall

Mitochondria

Amyloplasts

Endoplasmicreticulum

FIGURE 23.14 A statocyte (columella cell) containingthree statoliths (amyloplasts). (Based on an electronmicrograph of Lepidium root, Volkmann and Sievers,1979. In W. Haupt, M. E. Feinleib (eds.), Encyclope-dia of Plant Physiology, NS, Vol. 7, pp. 573–600. Berlin:Springer-Verlag.)

23.2 Gravitropism 403

stimulus, but these movements are thought to be a con-sequence of starch grain sedimentation. Although thereis still no direct proof for the starch-statolith hypothesis,there is a large body of evidence that is more consistentwith that idea than any other that has been put forwardto date. This evidence is summarized below:

1. Gravitropism is generally absent in plant species thathave no starch grains or amyloplasts. In lower plantssuch as algae and fungi, excess carbohydrate maynot be stored as starch. In these cases, some othersubstance may function as statoliths. In the algaChara, for example, the role of starch grains isreplaced by granules of barium sulphate.

2. There is a strong correlation between the rate of starchsedimentation and presentation time. In sweet pea(Lathyrus odoratus), for example, there is a parallelincrease in both sedimentation time and presenta-tion time as the temperature is lowered from 30◦Cto 10◦C. The decline in both is presumably relatedto an increase in protoplasmic viscosity.

3. Loss of starch by hormone treatment or mutation isaccompanied by a loss of graviresponse. For example,roots of cress seedlings (Lepidium sativum) treatedwith cytokinin or gibberellin at 35◦C becomestarch-free in 29 hours. The growth rate of treatedroots is reduced only slightly (0.48 mm h−1 vs.0.64 mm h−1) but any response to gravity iscompletely eliminated. Transfer of the roots towater in the light results in a parallel recovery ofboth amyloplasts and gravitropic responsivenessafter 20 to 24 hours. In maize (Zea mays) theamylomaize mutant produces smaller amyloplaststhan the wildtype. In studies of the percentageand speed of amyloplast sedimentation the degreeof coleoptile curvature was strictly correlatedwith the size of the amyloplast. Another mutantof maize, hcf-3 (high chlorophyll fluorescence-3) isunable to carry out photosynthesis and thus canform no starch in the leaf base statocytes when theendosperm reserves have been exhausted. Suchseedlings do not respond to gravity unless fedsucrose, in which case recovery of both amyloplastsand sensitivity to gravity was noted.

4. Amyloplasts can be displaced by a high-gradient magneticfield in place of gravity. An intracellular magnetic fieldhas been used to displace statoliths laterally (calledmagnetophoresis) in both roots and hypocotyls. Themagnetic field also induces a curvature similar to thegravitropic response.

It is not known how the sedimentation of statolithscreates physiological asymmetry in the cell or tissue,although a number of models have been proposed. Mostmodels agree that it is not the change in position ofthe statolith or the process of movement per se that

is important. The preferred view is that the statolithexerts pressure on one or more membranes or other cel-lular components. Although there is no direct evidencefor pressure-sensitive membranes in plants, both theplasma membrane and the endoplasmic reticulum (ER)have been suggested as likely targets. Electron micro-graphs of gravistimulated root statocytes, for example,commonly show amyloplasts sedimented on the endo-plasmic reticulum against the lower side of the cell(Figure 23.14).

23.2.3.3 The transmission and response phasesinvolve a lateral redistribution of auxin in theelongation zone. Like phototropism, developmentof curvature in response to gravity ultimately involves adifferential growth response. It is not surprising, then,that the Cholodny-Went hypothesis of asymmetricauxin distribution has dominated thinking and researchinto gravitropism for more than 60 years. Accordingly,the hypothesis states that horizontal orientation of theshoot or roots induces a lateral translocation of auxintoward the lower side of the organ. Auxin redistributionwould bias the growth rate in favor of the lower sidesuch that negatively gravitropic organs (e.g., coleoptilesand shoots) would turn upward. In positively gravitropicorgans such as roots, the higher concentration of auxinis thought to inhibit elongation on the lower side relativeto the upper, causing the organ to grow downward.Exogenous auxin, for example, very effectively inhibitsroot growth when applied at concentrations of 10−6

M or greater, concentrations that normally stimulateelongation of coleoptiles and shoots. In addition,gravitropic curvature is prevented by inhibitors of auxintransport (TIBA, NPA).

Auxin flow in the root is described by the ‘‘auxinfountain model’’ (Figure 23.15). Auxin synthesized in theshoot is transported basipetally through the shoot andinto the root. There it continues to move acropetallytoward the root tip through cells associated with thecentral vascular tissues, or stele. In the columella regionof the root cap, the auxin flow is reversed and the auxinmoves basipetally into the cortical cells of the elongationzone (Figure 23.15A). In a vertical root the flow ofauxin into the cortical region is distributed uniformlyaround the root. When the root is displaced horizontally(Figure 23.15B), the flow of auxin from the columella isredistributed laterally (i.e., downward). In other words,auxin flow in a horizontal root is biased toward the lowerside of the root. The implication is that the higher auxincontent on the lower side of the root inhibits elongationrelative to the upper side and the root curves downward.

23.2.3.4 Gravitropic induction in roots involvesseveral second messengers and redistribu-tion of auxin transporters. An early event ingravity-stimulated roots is a change in the membrane

404 Chapter 23 / Tropisms and Nastic Movements: Orienting Plants in Space

FIGURE 23.15 The path of auxin flow in roots.Auxin produced in the shoot flows into theroot through the central vascular tissue. In thecolumella (yellow square), the auxin stream isdiverted into the epidermal and cortical cell files,where it flows up toward the elongation zone.When the root is displaced from the vertical,events in the columella divert the main auxin flowtoward the lower side of the root.

Elongation

C

C

S C

C

Auxin inhibitselongation

A. B.

S

potential of the columella cells. Within seconds ofreorienting Lepidium sativum roots to a horizontalposition, the columella cells on the lower side of theroot depolarize (the potential becomes less negative)and those on the upper side hyperpolarize (the potentialbecomes more negative). These changes in membranepotential appear to be in some way related to thecytoskeleton because they are inhibited by cytochalasinD, a drug that disrupts the cytoskeleton by bindingto rapidly elongating actin filaments. There is alsoevidence that amyloplasts are connected to the plasmamembrane by actin filaments. These observationshave given rise to the hypothesis that displacement ofamyloplasts stimulates stretch-activated ion channels.Stretch-activated channels are so named because theyare activated in patch-clamp experiments when themembrane is stretched by applied suction. According tothis hypothesis, stretch-activated ion channels wouldbe responsible for the observed changes in membranepotential in the columella cells, which in turn wouldlead to the asymmetric distribution of auxin.

Changes in the pH of columella cells have alsobeen observed in gravity-stimulated roots. By usingpH-sensitive fluorescent proteins to monitor changesin pH during gravistimilation of Arabidopsis root, it hasbeen shown that pH of the root cap apoplast decreasesfrom pH 5.5 to 4.5 within 2 minutes of gravistimu-lation. Conversely, the cytoplasmic pH of columellacells increases slightly, from pH 7.2 to pH 7.6. ThesepH changes in the root cap precede auxin-related pHchanges in the elongation zone by about 10 minutes.

Several investigators have highlighted a role forcalcium in the gravity response of both coleoptilesand roots. Radio-labeled calcium (45Ca) accumulatedin the upper half of sunflower hypocotyls and maizecoleoptiles within one hour of stimulation by gravity.Calcium redistribution also occurred followingasymmetric application of exogenous IAA and could be

prevented in horizontal organs treated with the auxintransport inhibitor NPA. Histochemical techniqueshave demonstrated calcium localization in cells ofthe upper epidermis and underlying parenchymaof Avena coleoptiles within 10 minutes of gravis-timulation. In addition, both calcium redistributionand gravitropism are prevented by prior treatmentwith EGTA (ethyleneglycol-bis-(β-aminoethyl ether)-N,N′-tetraacetic acid), a chelator that ties up freecalcium, and the graviresponse of coleoptiles isprevented by treatment with chloropromazine, aninhibitor of the calcium-binding protein calmodulin.This suggests that the response to gravity might at onestage involve a Ca2+/calmodulin complex.

Experiments with exogenously applied calcium haveprovided equally convincing evidence of a role for cal-cium in root gravitropism. For example, asymmetricallyapplied agar blocks containing 10 mM CaCl2 will inducecurvature of maize roots, but only if the block is appliedto the root tip (Figure 23.16). Migration of calcium inroots appears to be restricted to the root cap—migrationis prevented if the cap is removed—and is directedtoward the lower side of the horizontal root. More-over, the calcium appears to move not through rootcap cells but through the thick mucilaginous layer thatcoats the root cap. The importance of this mucilaginouscoating is reflected in the observation that its contin-ual removal by washing renders the root insensitive togravity. Note that the direction of calcium asymmetryrelative to auxin is opposite in gravistimulated coleop-tiles and roots. In both organs, calcium migrates towardthe potentially concave side. Thus in a horizontally ori-ented root, calcium moves downward to accumulate onthe lower side of the root cap but it moves toward theupper side of a coleoptile. The source of this calciumis unknown, but it could be released from the ER asthe result of an interaction between the amyloplasts andthe ER.

23.3 Nastic Movements 405

A.

B.

C.

FIGURE 23.16 Calcium-induced curvature of the primaryroot of maize (Zea mays). (A) An agar block containingcalcium placed on the side of the root in the elongationzone has no effect. (B) When the agar block contain-ing calcium is applied to the root tip, the root growstoward the source of calcium. (C) An agar block contain-ing EGTA, a calcium chelator, causes the root to grow inthe opposite direction.

Finally, and perhaps not unexpectedly, recent exper-iments in which the PIN proteins were linked to a greenfluorescent protein and examined microscopically havedemonstrated that the auxin fountain and root grav-itropism depend on the coordinated distribution andactivities of the auxin efflux facilitators PIN1, PIN2,and PIN3 (Figure 23.17). PIN1 is localized at the apicalend of cells in the stele and is responsible for deliver-ing the auxin stream to the root apex. PIN3 is locatedalong the lateral wall of the columella cells in a verti-cally oriented primary root, where it diverts the flow ofauxin laterally, or centrifugally, toward the peripheralroot tissues. PIN2 is located primarily on the basal wallsof the peripheral root cap, epidermal, and cortical cellswhere it mediates the basipetal stream of auxin towardthe cell elongation zone. The importance of PIN2 ingravitropism is indicated by the observation that pin2mutants fail to establish the required lateral auxin dis-tribution following gravistimulation and do not exhibita normal gravitropic response.

When a vertical root is rotated to the horizon-tal position, two significant events occur. First, PIN3becomes redistributed, accumulating along the lowersidewalls of the columella cells. This redistribution ofPIN3 presumably diverts the incoming auxin toward thelower side of the root. Second, PIN2 becomes asym-metrically distributed between the upper and lower sidesof the root. The evidence suggests that the PIN2 pro-tein is rapidly turned over due to ubiquitination and

A B

FIGURE 23.17 The expression and distribution of theauxin transport facilitator PIN2 in vertical (A) and grav-istimulated (B) roots. In vertical roots PIN2 is locatedmore or less symmetrically throughout the peripheralroot cap, epidermis, and cortex, mediating the basipetalflow of auxin toward the cell elongation zone uniformlyaround the root. In gravistimulated roots, enhanceddegradation of PIN2 at the upper side of the root biasesPIN2 location to the lower side.

proteasome-dependent degradation. Gravistimulationapparently leads to an increased rate of PIN2 degra-dation in the upper side of the root. At the same time,higher auxin levels were found to promote increasedretention of PIN2 in the lower side of the root. Thisunequal distribution of PIN2 would thus help to main-tain the auxin gradient originally established by PIN3in the columella cells.

But how does a root straighten out when it onceagain approaches a vertical orientation? It turns out thatauxin can also promote the degradation of PIN2. It hasbeen proposed that as auxin levels continue to buildup in the lower side of the root, concentrations even-tually reach a threshold where auxin-mediated PIN2degradation begins to reduce auxin transport toward theelongation zone, thus reducing further curvature.

The big unresolved question at this time, of course,is how the settling of amyloplasts in response to gravityinduces changes in pH, calcium flux, and other secondmessengers and how these changes are all linked tochanges in the distribution and activity of the auxintransport proteins.

23.3 NASTIC MOVEMENTS

In addition to the directed movements of tropisms,many plants and plant parts, especially leaves, exhibitnastic movements, in which the direction of movementis not related to any vectorial component of the stim-ulus. Nastic responses may involve differential growth,in which case the movement is permanent. Alterna-tively the movement may be reversible, caused by turgorchanges in a specialized motor organ.

406 Chapter 23 / Tropisms and Nastic Movements: Orienting Plants in Space

Epinasty and thermonasty are examples of nasticresponses involving differential growth. Epinasty is thedownward bending of an organ, commonly petiolesand leaves whose tips are inclined toward the ground.It is not a response to gravity, however, but appearsto depend on an unequal flow of auxin through theupper and lower sides of the petiole. Epinasty is also acommon response to ethylene or excessive amounts ofauxin. The reverse response, called hyponasty, is lesscommon but can be induced by gibberellins. A typicalexample of thermonasty is the repeated opening andclosure of some flower petals, such as tulip and crocuses.In spite of their repeated nature, however, thermonasticmovements are permanent and result from alternatingdifferential growth on the two surfaces of the petals.

The most dramatic nastic movements are all turgormovements, which may be broadly separated intothree categories: (1) leisurely rhythmic leaf movementsin nyctinastic plants, (2) very rapid seismonasticmovements in a limited number of species, and(3) thigmonastic or thigmotropic curling of threadlikeappendages in climbing plants and vines. Nyctinasticand seismonastic responses depend on differentialturgor movements in specialized motor organs, calledthe pulvinus (pl. pulvini). The pulvinus is a bulbousstructure most often encountered in plant familiescharacterized by compound leaves, such as the Legumi-noseae and Oxalidaceae (Figures 23.18 and 23.19). It

Secondarypulvinus

Pinnule

Rachis

Tertiarypulvinus

Rachilla

Pinna

Primarypulvinus

Open Closed

FIGURE 23.18 A leaf of Samanea samanan, illustratingthe location of primary and secondary pulvini. Activationof the primary pulvinus causes leaflets to fold upward,parallel to the rachis. Activation of the secondary pulv-inus causes the rachilla to fold downward. (Reproducedfrom the Journal of General Physiology 40:413–430, 1974,by copyright permission of the Rockefeller UniversityPress.)

occurs at the base of the petiole (primary pulvinus),the pinna (secondary pulvinus), or the pinnule (tertiarypulvinus). The pulvinus contains a number of large,thin-walled motor cells, which alter the position of theleaf by undergoing reversible changes in turgor.

23.3.1 NYCTINASTIC MOVEMENTSARE RHYTHMIC MOVEMENTSINVOLVING REVERSIBLETURGOR CHANGES

Nyctinastic movements (Gr. nyctos, night + nastos = clo-sure) are most evident in leaves that take up a differentposition in the night from that taken during the day.Typically leaves or leaflets are in the horizontal, or open,position during the day and assume a more vertical, orclosed, orientation at night. The primary leaf of com-mon bean plants exhibits particularly strong nyctinasticmovements but this can also be seen in Coleus, prayerplants, and other common garden and house plants.Observations of nyctinastic movements can be tracedback as far as the writings of Pliny in ancient Greece.The Swedish botanist C. Linnaeus (in 1775) coined theterm ‘‘plant sleep’’ to describe nyctinastic movementsand they are commonly referred to as sleep movementstoday. The sleep movements of bean were prominent inthe discovery of endogenous biological clocks and aredescribed further in Chapter 24.

Sleep movements have been studied by severaleighteenth- and nineteenth-century botanists, includ-ing Darwin. The process, however, has been studiedmost extensively by Ruth Satter and her colleagues inSamanea samanan, a member of the Leguminoseae withdoubly compound leaves (Figure 23.18). In Samanea thepaired pinnae and pinnules are normally separated andspread apart, while in closing they fold toward eachother. In Samanea, the paired pinnules fold basipetally(i.e., downward), but in other species, such as Mimosapudica and Albizzia julibrissin, closure of paired pinnulesis upward, or acropetal. The doubly compound leavesof Mimosa, Albizzia, and Samanea all have three pulvini,but the simple leaf of Phaseolus (bean) has only two. It isthe secondary pulvinus that generally exhibits the morerapid or dramatic change and has consequently beenstudied most extensively (Figure 23.19). They are alsorelatively large (2 to 3 mm diameter, 4 to 7 mm longin Samanea) and the changes in curvature are readilyvisible to the naked eye.

All nyctinastic responses depend on reversible tur-gor changes in the pulvinus. The pulvinus is typicallycylindrical in shape, with prominent transverse furrowswhich facilitate bending, on the adaxial and abaxial sides(Figure 23.19). It contains a central vascular core withboth xylem and phloem surrounded by sclerenchymatissue. The vascular tissue assumes a linear arrangement

23.3 Nastic Movements 407

A.

Vascular bundle

Flexor cells

Corticalparenchyma

EpidermisExtensor cells

B.

Adaxial surface

Abaxial surface

FIGURE 23.19 The pulvinus. (A) Secondary pulvini of bean (Phaseolus vulgaris). Thesecondary pulvinus is the swollen area (arrows) at the juncture of the petiole withthe stem, just below the axillary bud. In bean, the primary pulvinus is located at thedistal end of the petiole, where it attaches to the leaf. During closure in bean, thepetioles fold toward the stem axis while the leaf blade drops from the horizontalto the vertical position. (B) Schematic diagram of the bean secondary pulvinus incross-section, showing the enlarged cortical parenchyma region. During closure, themotor cells of the extensor region lose turgor, while the motor cells of the flexorregion gain turgor and the petiole moves toward the stem axis. During opening, theextensor cells gain turgor and the flexor cells loose turgor, thus driving the petioleaway from the stem axis.

as it passes through the pulvinus, apparently enhanc-ing the flexibility of the pulvinar region. Outside thevascular core is a cortex comprised of 10 to 20 layersof parenchyma cells. The cells of the outer cortex havethin, elastic walls and exhibit large changes in size andshape during movement. These are called motor cells.Changes in the size and shape of the motor cells areresponsible for leaf movement.

The opposite sides of the pulvinus are known asthe extensor and flexor regions (Figure 23.19B). Theextensor region is formed by motor cells that lose turgorduring the bending movement, or ‘‘closure.’’ Motorcells in the flexor region gain turgor during closureand lose turgor during opening. Thus, the swellingof extensor motor cells and shrinkage of flexor motorcells straightens the pulvinus and opens or spreads apartleaves or leaflets. The relative positions of extensorand flexor regions in the pulvinus (whether adaxial orabaxial, for example) will be reversed, depending onwhether closure is basipetal or acropetal.

Nyctinastic movements are sensitive to blue light,the physiological status of phytochrome, and endo-

genous rhythms. Although the mechanism of signalperception and how these three stimuli interact is notknown, it is clear that both the receptors and theresponding system (the motor cells) are located in thepulvinus—at most a few cells apart. It is known thatphytochrome can ‘‘reset’’ the endogenous clock thatregulates nyctinastic leaf oscillations. The relationshipbetween phytochrome and endogenous rhythms will bediscussed in the next chapter.

23.3.2 NYCTINASTIC MOVEMENTSARE DUE TO ION FLUXESAND RESULTING OSMOTICRESPONSES IN SPECIALIZEDMOTOR CELLS

Regardless of the nature of the stimulus, motor cellvolume changes are due to osmotic water uptake (orloss) as a result of ion accumulation (or loss) acrossthe cell membrane. What ions are exchanged, how arethey transported, and what is the driving force for ion

408 Chapter 23 / Tropisms and Nastic Movements: Orienting Plants in Space

movement? These questions have been approachedwith a variety of techniques, including histochemi-cal and radiochemical methods, and scanning electronmicroscopy coupled with X-ray analysis. The resultsshow that leaf movement in all nyctinastic plants stud-ied thus far is associated with a massive redistribution ofpotassium ion (K+) between the symplast and apoplastin both the extensor and flexor regions of the pulvinus(Figure 23.20). Swollen extensor cells are characterizedby high protoplasmic K+ and low apoplastic K+. InPhaseolus vulgaris, fully 30 percent of the osmotic poten-tial change can be accounted for by K+ movement. Thecharge carried by K+ is compensated primarily by chlo-ride and possibly small organic anions such as malateand citrate.

Beyond the central role of K+ flux in nastic move-ments, it is difficult to pin down the specific sequenceof molecular and biophysical events in the signal chain.Patch-clamp experiments with isolated Samanea motorcell protoplasts have established that K+ exchange acrossthe plasma membrane occurs through K+ channels andthat these channels can be regulated by changing themembrane polarity. Depolarization of the membraneopens the channels and allows K+ to move out of thecell down its electrochemical gradient. It has also beenestablished that there are pH gradients across the plasmamembranes of motor cells. In the case of open Samaneapulvina, the pH of the apoplast is about 5.5 in theflexor region and 6.2 in the extensor region. The cyto-plasmic pH is approximately neutral in both regions.Upon a light/dark transition (leading to closure), thepH gradient in the extensor region dissipates whilein the flexor region the gradient increases. Althoughquantitative relationships between H+ flux and K+ fluxhave yet to be tested, H+ extrusion could contributeto the electrochemical gradient necessary to drive K+uptake. Any observed changes in membrane potentialare undoubtedly the consequence, not the cause, of thecross-membrane traffic in osmotically active ions.

The prominent roles of K+ channels and H+pumps have been incorporated into the current modelfor motor cell movement. A simplified version of themodel is shown in Figure 23.21. In this model, thelight signal activates phytochrome (or cryptochrome),which accelerates inositol phospholipid turnover.Recent experiments have shown that light thatstimulates opening of Samanea pulvini also decreasesthe level of phosphotidylinositol 4,5-bisphosphate(PIP2) and increases the level of the second messengerinositol 1,4,5-triphosphate (IP3). There is a transientstimulation of diacylglycerol (DAG). These changesare qualitatively similar but quantitatively smallerthan those normally detected in animal systems. Theassays, however, involved whole pulvini, which containvascular, collenchyma, and epidermal tissues as well as

0

10

20

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K+ A

ctiv

ity

(mM

)

30

40

50

60

70

80

0

10

20

30

0 20 40 60 80 100

FIGURE 23.20 Changes in K+ activity in the apoplast ofextensor (upper curve) and flexor (lower curve) cellsduring closure of Samanea leaflets. Loss of K+ from theprotoplasts is followed by loss of water and turgor. Clo-sure and opening were stimulated by dark and whitelight periods as indicated by the bar between the twographs. (Adapted from Lowen, C. Z., R. L. Satter. 1989.Light promoted changes in the apoplastic K+ activityin the Samanea samanan pulvinus. Planta 179:412–427,Figure 1. Copyright Springer-Verlag.)

the motor cells. The changes could be appropriatelygreater if restricted to the smaller population of motorcells. If inositol phospholipid metabolism functions inplants as it does in animals, DAG would be expectedto activate a protein-kinase C (or its plant equivalent)to phosphorylate certain proteins. IP3 would beexpected to release free calcium—exogenous IP3 doesliberate calcium—although from which compartmentis not known. Both the phosphorylated protein and/ortransient increases in free calcium stimulate protonextrusion by activating the proton pump. The resultingelectrochemical gradient energizes the uptake of K+and other ions, which in turn stimulates the osmotic

23.3 Nastic Movements 409

+

−] ]

K+ C1–

Ca2+

Plasmamembrane

Cytosol

Protein + ATP Protein ~ P

hν

H+

Pr

?

Pfr PIP2

[DAG]

[IP3 [Ca2+

FIGURE 23.21 A proposed model for the interaction of phytochrome, biologicalclocks, and the inositol triphosphate system in leaf movements of nyctinastic plants.Light, mediated by phytochrome and modulated by the endogenous clock, acceler-ates inositol phospholipid turnover and increases the level of the second messengersinositol-1,4,5-triphosphate (IP3) and diacylglycerol (DAG). The second messengersstimulate a release of Ca2+ into the cytosol and phosphorylation of various proteinswhich in turn stimulate the extrusion of protons from the cell. K+ diffuses into thecell in response to the proton motive force. An active transport pump extrudes Ca2+

as an aid to restoring Ca2+ homeostasis.

uptake of water and motor cell swelling. The presenceof a calcium pump that extrudes Ca2+ would help toensure the restoration of Ca2+ homeostasis.

Many details of this model remain to be described,especially the function of the inositol phospholipid cyclein plants. Still, plant cells are known to contain virtuallyall the required components and the model is consistentwith what has been observed in pulvini thus far. Signifi-cant advances are to be expected in the future, especiallynow that patch-clamp techniques—long a mainstay ofelectrophysiology research for animal cells—can beapplied to plant cell protoplasts. This state-of-the-arttechnique has been in use for plant cells only sinceabout 1984, but has proven invaluable for the study ofion channels.

23.3.3 SEISMONASTY IS A RESPONSE TOMECHANICAL STIMULATION

A limited number of leguminous plants that possesspulvini and exhibit nyctinastic movements also exhibit aresponse to mechanical stimulation. This phenomenonis known as seismonasty. Since seismonastic plantsrespond to touch, they are sometimes consideredthigmonastic. However, seismonastic plants respond to

a wider variety of stimuli including shaking or wind,falling raindrops, wounding by cutting, and intense heator burning.

The best known example of seismonastic plants isthe tropical shrub Mimosa pudica (Figure 23.22). Thesurvival advantage of such a response is not certain.Some have suggested that since these plants grow inarid, exposed areas where they are exposed to dryingwinds, folding of the leaves may be a means of reducingwater loss. Others suggest that it is a means of protectionfrom large herbivores or insects. However, one thing isclear—the response is very rapid. When the pulvinusis stimulated directly, bending begins in less than onesecond!

The ultimate response, leaf movement, of courseinvolves movement of pulvini motor cells just as in nycti-nastic movements. However, there are three essentialcharacteristics of the seismonastic response that haveserved to focus attention on the early steps of signaltransduction. The first of these is the rapidity of theresponse. Second, seismonasty follows the ‘‘all-or-noneprinciple,’’ which means that there is no obvious rela-tionship between the intensity of the stimulus and theextent of the response. Third, excitation is propagatedfrom the place of stimulation. The similarity of thesecharacteristics to animal nerve transmission has given

410 Chapter 23 / Tropisms and Nastic Movements: Orienting Plants in Space

A. B.

FIGURE 23.22 Seismonasty in the sensitive plant Mimosa pudica. The plant is shownin the open (A) and closed (B) positions. The plant on the right (B) was photographedabout 10 seconds after closure was stimulated by a sharp tap to the stem with a pencil.

rise to the expectation that plants may also be capableof transmitting stimuli in the form of potential changes.Indeed it has now been well established that virtuallyany part of the Mimosa plant can perceive stimuli andtransmit them as electric pulses to the pulvini. Althoughplants do not have discrete nerve tissue, it appears thatphloem sieve tubes can and do function as conduits forsignal transmission. Stimulation of the petiole results in arapid depolarization that is propagated basipetally alongthe sieve tube at a rate of about 2 cm s−1. The uniquestructure of the sieve tube with its protoplasmic conti-nuity through the sieve plates appears to be well suitedfor transmission of electrical signals. The appearance ofthe action potential is correlated with a rapid uptake ofprotons, suggesting that proton flux is responsible forthe depolarization. When the action potential reachesthe pulvinus, it appears to stimulate a rapid unloading ofboth K+ and sugars into the apoplast. Water would fol-low and the resulting loss of turgor would cause collapseof the motor cells.

Other investigators have found that substances iso-lated from phloem sap of Mimosa and other species willstimulate closure of Mimosa pulvini when applied to thecut end of the stem. The active substance has been iden-tified as a glycosylated derivative of gallic acid (4-0-β-d-gluco-pyranosyl-6′-sulphate)). Called ‘‘turgorin,’’ this

substance has been isolated from 14 higher plants thatexhibit nyctinastic movements. It has been suggestedthat turgorin may give rise to action potentials in amanner similar to the animal neurotransmitter, acetyl-choline.

SUMMARY

Plant movements serve to orient the plant body inspace. Thus roots exhibit positive gravitropism, grow-ing down in order to mine the soil for mineral nutrientsand water. Shoots exhibit negative gravitropism andpositive phototropism in order to optimize the inter-ception of sunlight for photosynthesis.

There are several categories of plant movements.Growth movements involve cell division and elonga-tion and are consequently irreversible. Turgor move-ments involve changes in turgor pressure and cellvolume, and are reversible. Tropisms are direction-ally related to the stimulus whereas the directionalityof nastic movements is inherent in the tissue and arenot related to any vector in the stimulus. Nutations arerotary or helical movements that are best observed withtime-lapse photography.

Further Reading 411

Under natural conditions, phototropism is agrowth response to a light gradient, although in thelaboratory it is usually studied by subjecting organsto unilateral light. Organs may either grow toward(positive phototropism) or away from (negativephototropism) the higher irradiance. Phototropismis a response to blue and UV-A light; mediated by aflavoprotein called phototropin, located in the plasmamembrane.

The phototropic response is characterized by dif-ferential growth on the lighted and shaded sides of theresponding organ. The most generally accepted the-ory to account for differential growth in coleoptilesand stems is the Cholodeny-Went theory. This the-ory proposes a lateral redistribution of auxin as it flowsbasipetally from the apex where it is synthesized. In thecase of positive phototropism, the higher concentrationof auxin flows down the shaded side of the organ, caus-ing cells on the shaded side to elongate more rapidlythan those on the lighted side.

Unlike most stimuli to which plants are exposed,gravity is omnipresent and nonvarying. There is nogravitational gradient. Gravity can be sensed only bymovement of cellular structures (statoliths), whichthen establishes an asymmetry that is translated interms of pressure. Although there is no direct proof,the weight of evidence indicates that statoliths are thestarch-containing plastids, amyloplasts.

Sensitivity to gravity in the root is localized inthe columella, a group of cells in the central coreof the root cap. The primary transducer that sensesthe pressure of the statoliths and initiates the sig-nal transduction chain remains unknown. Some evi-dence suggests it might be the endoplasmic reticulum.Another theory proposes that the sedimenting amy-loplasts activate stretch-activated ion channels in theplasma membrane. As in phototropism, the gravit-ropic response involves differential growth that canbe explained by redistribution of auxin transport. Thesteps between pressure sensing and auxin redistribu-tion are unknown, but experiments indicate that pHchanges, calcium ions, and inositol triphosphate mayall be involved. Ultimately, the signal chain results inthe relocation of auxin transport facilitators of the PINfamily.

Plants exhibit a variety of nastic responses. Oneof the most prominent is the periodic movement ofleaves known as sleep movements, or nyctinasty. Leafmovement is mediated by turgor changes in special-ized motor cells located in structures called pulvini,found at the distal end of the petiole. Turgor changesare mediated by a flux of potassium ion induced by aninteraction between phytochrome, biological clocks,and the inositol triphosphate system. Another nastic

response is illustrated by seismonasty in the sensi-tive plant Mimosa. Seismonasty involves similar turgorchanges in response to physical disturbance.

CHAPTER REVIEW

1. Define phototropism. Is phototropism restri-cted to unilateral lighting? In what way(s) can alight gradient be established across a plant organsuch as a stem?

2. What pigment(s) function as the photoreceptorfor phototropism? List the evidence that supportsyour conclusion.

3. Review the experimental basis for the Cholodny-Went hypothesis.

4. Shoots and roots express various levels of gravit-ropic response. What are the physiological advan-tages to be gained by such variation in response?

5. Review the statolith theory for gravitropismas it applies to roots. How is the gravita-tional stimulus perceived by a root and howdoes it respond? What is the evidence thatcalcium is involved in root gravitropism?

6. Describe nyctinasty. What might be the physio-logical significance or survival value of nyctinasty?In what ways is the seismonastic response similarto nyctinasty? In what ways is it different?

FURTHER READING

Abas, L. et al. 2006. Intracellular trafficking and proteolysisof the Arabidopsis auxin-efflux facilitator PIN2 areinvolved in root gravitropis. Nature Cell Biology8:249–256.

Blancaflor, E. B., J. M. Fasano, S. Gilroy 1998. Mappingthe functional roles of cap cells in the response ofArabidopsis primary roots to gravity. Plant Physiology116:213–222.

Brown, A. H. 1993. Circumnutations: From Darwin to spaceflights. Plant Physiology 101:345–348.

Celaya, R. B., E. Liscum 2005. Phototropins and associatedsignaling: Providing the power of movement in higherplants. Photochemistry and Photobiology 81:73–80.

Christie, J. M. 2007. Phototropin blue-light receptors.Annual Review of Plant Biology 58:21–45.

Darwin, C. 1881. The Power of Movement in Plants. New York:Appleton-Century-Crofts.

Esmon, C. A. et al. 2006. A gradient of auxin and auxin-dependent transcription precedes tropic growthresponses. Proceedings of the National Academy of Sciences,USA. 103:236–241.

Fasano, J. M., S. J. Swanson, E. B. Blancaflor, P. E. Dowd,T. Kao, S. Gilroy 2001. Changes in root cap pH are

412 Chapter 23 / Tropisms and Nastic Movements: Orienting Plants in Space

required for the gravity response of the Arabidopsis root.The Plant Cell 13:907–921.

Haga, K., M. Iino 2006. Asymmetric distribution of auxincorrelates with gravitropism and phototropism but notwith autostaightening (autotropism) in pea epicotyls.Journal of Experimental Botany 57:837–847.

Iino, M. 2006. Toward understanding the ecological func-tions of tropisms: Interactions among and effects of lighton tropisms. Current Opinion in Plant Biology 9:89–93.

Jarillo, J. A. et al. 2001. Phototropin-related NPL1 con-trols chloroplast relocation induced by blue light. Nature410:952–954.

Kimura, M., T. Kagawa 2006. Phototropin and light-signaling in phototropism. Current Opinion in PlantBiology 9:503–508.

Morita, M. T., M. Tasaka 2004. Gravity sensing and signal-ing. Current Opinion in Plant Biology 7:712–718.

Perbal, G., D. Driss-Ecole 2003. Mechanotransduction ingravisensing cells. Trends in Plant Science 8:498–504.

Satter, R. L., H. I. Gorton, T. C. Vogelmann 1990. The Pul-vinus: Motor Organ for Leaf Movement. Rockville, MD:American Society of Plant Physiologists.

24Measuring Time: ControllingDevelopment by Photoperiod

and Endogenous Clocks

Two hundred and fifty years ago, Carl v. Linne, betterknown for his development of the binomial system ofnomenclature, designed a flower clock based on theopening and closing of the petals at specific but differenttimes of the day. The plants were arranged in a circleand one could tell the time of day by simply notingwhich flowers were open and which were closed. It isoften difficult for the layman to understand that plantscan tell time without a Timex™, but many aspects ofplant behavior can be interpreted in no other way. Oneexample is the consistent flowering of various speciesat particular times of the year. Roses always bloom inthe summer and chrysanthemums in the fall. Indeed,the flowering of many plants is so predictable fromone year to the next that gardeners have for centuriesincorporated them into their gardens as floral calendars,unerringly marking the progress of the seasons. In thenorthern latitudes, perennial plants sense the short daysof autumn as a signal to induce bud dormancy, thusanticipating the unfavorable conditions of winter. Themost reliable indication of the advancing season is thelength of day, and an organism’s capacity to measurethe proportion of daylength in a 24-hour period is

known as photoperiodism. However, photoperiodismis only one of the more outward manifestations of a farmore fundamental timekeeping mechanism, known asthe biological clock.

In this chapter we will examine

• photoperiodism; including the distinction betweenshort-day plants, long-day plants, and otherresponse types; the central role of the dark period;the nature of photoperiodic perception; and adiscussion of current hypotheses to account for theelusive floral stimulus,

• vernalization—the low-temperature requirementfor flowering in winter annuals and biennialplants,

• the biological clock, with an emphasis on endoge-nous rhythms with a 24-hour periodicity and theirrole in photoperiodic time measurement,

• the molecular genetic basis for the circadian clockand the search for the central oscillator, and

• a brief discussion of the significance of photoperi-odism in nature.

413

414 Chapter 24 / Measuring Time: Controlling Development by Photoperiod and Endogenous Clocks

Flowering genes

Shoot ApicalMeristem

PhytochromeCryptochrome

FT

CO

FT FT FT

BOX 24.1HISTORICALPERSPECTIVES:THE DISCOVERYOFPHOTOPERIODISM

Although it had earlier been suggested that latitudinalvariations in daylength contributed to plant distribu-tion, the first efforts at controlled experimentation wereconducted by a French scientist in 1912. J. Tournoisfound that both Humulus (hops) and Cannabis (hemp)plants flowered precociously during the winter in thegreenhouse. Tournois eliminated temperature, humid-ity, and light intensity as environmental cues and in1914 concluded that the changing of either daylength ornightlength was responsible for early flowering. Unfor-tunately, World War I intervened and Tournois didnot live to continue his experiments. At the same time,H. Klebs was studying flowering of Sempervivum funkii(commonly known as ‘‘hens-and-chickens’’). Semper-vivum grows as a vegetative rosette in the wintergreenhouse. By supplementing normal daylight withartificial light, Klebs was able to break the rosette habit,stimulate stem elongation, and induce flowering. Fromhis experiments, Klebs concluded that length of daytriggered flowering in nature. However, it remained forW. W. Garner and H. A. Allard to demonstrate thefull impact of daylength on flowering and coin the termphotoperiodism.

W. W. Garner and H. A. Allard were scientists withthe U.S. Department of Agriculture near Washington,D.C. The initial focus of their work was a mutantcultivar of tobacco (Nicotiana tabacum), called MarylandMammoth. In the field, Maryland Mammoth plantsgrew to be very tall with large leaves. Such characteristicswould obviously be advantageous to the tobacco industryat the time (in the early 1920s), but breeding effortswere frustrated by the fact that the plants would notflower in the field during the normal growing seasonat that latitude. In the greenhouse, however, even verysmall plants flowered in the winter and early spring.Clearly, flowering was not simply a matter of the age of

the plants. Another problem of interest to Garner andAllard concerned flowering in soybean (Glycine max).When the cultivar Biloxi was sown over a 3-monthperiod from May to August, all of the plants floweredwithin a 3-week period in September (Figure 24.1).The earliest seeded plants thus took 125 days to flowerwhile those seeded last required only 58 days. Again itappeared that all plants, regardless of age, were simplyawaiting some signal to initiate flowering.

Like Tournois, Garner and Allard eliminated avariety of environmental conditions (such as nutrition,temperature, and light intensity) as the ‘‘signal,’’ com-ing finally (and with some reluctance) to the conclusionthat flowering was controlled by the relative length ofday and night. Using a crude but effective system ofrolling plant benches in and out of darkened garagelikebuildings at predetermined times, Garner and Allardproceeded to describe the flowering characteristics ofscores of different species with respect to daylength.They went on to suggest that bird migration mightalso be keyed to daylength—a phenomenon that is nowwell documented. We now know that photoperiodiccontrol is not limited to flowering, but is a basic regu-latory component in many aspects of plant and animalbehavior.

May June July August Sept

Days toFlowering

58

69

77

92

94

125

F

F

F

F

F

F

FIGURE 24.1 September soybeans. Soybeans (Glycinemax, cv. Biloxi) sown over a three-month period allflower within a three-week period in September.

24.1 PHOTOPERIODISM

The switch from the vegetative state to the floweringstate is arguably one of the most dramatic and mysteriousevents in the life of a flowering plant. Photoperiodisminfluences many aspects of plant development such

as tuber development, leaf fall, and dormancy, but thecontrol of flowering by photoperiod has attracted themajor share of interest. Indeed, it was the failure of atobacco (Nicotiana tobacum) mutant to flower under fieldconditions that led to the discovery of photoperiodism(Box 24.1).

24.1 Photoperiodism 415

24.1.1 PHOTOPERIODIC RESPONSESMAY BE CHARACTERIZED BY AVARIETY OF RESPONSE TYPES

Photoperiodic responses fall into one of three generalcategories. They are: short-day plants (SD plants),long-day plants (LD plants), and daylength-indifferentor day-neutral plants (DNP) (Table 24.1 andFigure 24.2). Short-day plants are those that floweronly, or flower earlier, in response to daylengthsthat are shorter than a certain value within a 24-hourcycle. Long-day plants respond to daylengths that arelonger than a certain value, while day-neutral plantsflower irrespective of daylength. Within the long- andshort-day categories, we also recognize obligate andfacultative requirements.1 Plants that have an absoluterequirement for a particular photoperiod before theywill flower are considered obligate photoperiodic types.The common cocklebur (Xanthium strumarium), forexample, is an obligate short-day plant. Xanthium willnot flower unless it receives an appropriate shortphotoperiod. On the other hand, most spring cerealssuch as wheat (Triticum sp.) and rye (Secale cereale) arefacultative long-day plants. Although spring cereals willeventually flower even if maintained under continuousshort days, flowering is dramatically accelerated underlong days. The popular research object Arabidopsisis also a facultative long day plant. However, thedistinction between obligate and facultative responseis not always hard and fast for a particular species orcultivar. Photoperiod requirement is often modified byexternal conditions such as temperature. A particularspecies may, for example, have an obligate requirementat one temperature but respond as a facultative plant atanother temperature.

In addition to these three basic categories, thereare a number of other response types that flowerunder some combination of long and short days. Var-ious species of the genus Bryophyllum are, for example,long-short-day plants (LSD plant)—they will floweronly if a certain number of short days are precededby a certain number of long days. The reverse is trueof the short-long-day plant (SLD plant) Trifoliumrepens (white clover). A few plants have highly special-ized requirements. Intermediate-daylength plants, forexample, flower only in response to daylengths of inter-mediate length but remain vegetative when the day iseither too long or too short. Another type of behavioris amphophotoperiodism, illustrated by Madia elegans(tarweed). In this case, flowering is delayed under inter-mediate daylength (12 to 14 hours) but occurs rapidlyunder daylengths of 8 hours or 18 hours. There aremany, often subtle, variations to the three basic response

1The terms obligate and facultative may be interchanged withthe terms qualitative and quantitative, repectively.

TABLE 24.1 Representative plants exhibitingthe principal photoperiodic response types.

Short-Day Plants

Chenopodium rubrum red goosefootChrysanthemum sp. chrysanthemumCosmos sulphureus yellow cosmosEuphorbia pulcherrima poinsettiaGlycine max soybeanNicotiana tobacum tobacco (Maryland Mammoth)Perilla crispa purple perillaPharbitis nil Japanese morning gloryXanthium strumarium cocklebur

Long-Day Plants

Anethum graveolens dillBeta vulgaris Swiss chardHyoscyamus niger black henbaneLolium sp. rye grassRaphanus sativus radishSecale cereale spring ryeSinapis alba white mustardSpinacea oleracea spinachTriticum aestivum spring wheat

Day-Neutral Plants

Cucumis sativus cucumberGomphrena globosa globe amaranthHelianthus annuus sunflowerPhaseolus vulgarus common beanPisum sativum garden peaZea mays corn

types, encompassing a large number of flowering plants.However, most of what is known about the physiol-ogy of photoperiodism in plants has been learned froma relatively small number of short-day and long-dayplants.

24.1.2 CRITICAL DAYLENGTH DEFINESSHORT-DAY AND LONG-DAYRESPONSES

It is important to understand that the distinctionbetween SD plant and LD plant is not based on theabsolute length of day. Consider, for example, that bothXanthium and Hyoscyamus niger (black henbane) willflower with 12 to 13 hours of light per day (Figure 24.3).Yet the former is properly classified as a SD plant andthe latter as a LD plant. Whether a plant is classified

A.

Pharbitis nil

SHO

RT

DA

YS

LO

NG

DA

YS

Hyoscyamus niger

B.

C. D.

FIGURE 24.2 Flowering response of the SD plant Japanese morning glory (Pharbitisnil) and the LD plant black henbane (Hyoscyamus niger) to short days and long days.Note the prominent flowers (arrows) in Japanese morning glory under short days andin black henbane under long days. Note also that black henbane remains as a rosetteunder short days. Plants of each species under both photoperiod regimes are of thesame age.

416

24.1 Photoperiodism 417

Length of photoperiod (hrs)

Rel

ativ

e fl

ower

ing

Hyoscyamus

6

Xanthium

12 18 24

50

100

FIGURE 24.3 A diagram to illustrate theconcept of critical daylength in popula-tions of Xanthium strumarium (cockle-bur), a short day plant, and in Hyoscyamusniger (black henbane), a long day plant.Critical daylengths are indicated by thevertical dotted lines. Note that Xanthiumflowers when the daylength is shorterthan its critical daylength and Hyoscya-mus flowers when the daylength is longerthan its critical daylength.

as a SD plant or LD plant depends instead on itsbehavior relative to a critical daylength. Plants thatflower when the daylength is shorter than some criticalmaximum are classified as SD plant. Those that flowerin response to daylengths longer than a critical minimumare classified as LD plant. Thus the critical daylengthfor the SD plant Xanthium is 15.5 hours, meaning thatit will flower whenever the daylength is less than 15.5hours out of every 24. The critical daylength for theLD plant Hyoscyamus is 11 hours and it will flowerwhen the daylength exceeds that value. In the absence offurther information, the actual daylength under whicha plant will flower is no indication of its response type.A corollary to this observation is that, although SDplants tend to flower in the spring and fall and LDplants tend to flower in midsummer, the classification asa SD plant or LD plant is not necessarily an indicationof the time of year that species will flower.

Many plants require more or less continuous expo-sure to the appropriate photoperiod, at least until floralprimordia have been developed, in order to flower suc-cessfully. Others will proceed to flower even if, onceexposed to even a single proper photoperiod, the plantis returned to unfavorable photoperiods. Such plantsare said to be induced and the appropriate photope-riod is referred to as an inductive treatment. Thephenomenon of induction raises intriguing, thoughunresolved, questions about the physiological propertiesof the induced state. Clearly a physiological change hastaken place in induced plants and this change persists,even though no anatomical or morphological change isevident at the apex where flowers will appear.

Induction can also be experimentally useful. Oneof the reasons Xanthium has been so widely used forstudies of photoperiodism is that a single short-daycycle will irreversibly lead to flowering, even in plantsthat are returned to long days. Such an extreme sensi-tivity to induction is not widespread, but it has beendemonstrated in other SD plants such as Japanese

morning glory (Pharbitis nil), duckweed (Lemna pur-pusilla), and lambs quarters (Chenopodium rubrum), andin the LD plants dill (Anethum graveolens) and rye grass(Lolium temulentum).

Induction is not an all-or-none process, but can beachieved in degrees. Although Xanthium will respond toa single inductive cycle, the initiation of floral primordiais more rapid and more prolific if multiple cycles aregiven. Other plants may exhibit fractional induction—asummation of inductive photoperiods despite inter-ruption with noninductive cycles. Plantago lanceolata(plantain), for example, normally requires a thresholdof about 25 long days to induce flowering. Plants givenonly 10 long days will remain vegetative, but a scheduleof 10 long days followed by 10 short days and thenanother 15 long days will induce flowering. The plantsare apparently able to sum the long days in spite of theintervening short days. Examples of summation withup to 30 intervening noninductive periods have beenreported.

24.1.3 PLANTS ACTUALLY MEASURETHE LENGTH OF THE DARKPERIOD

In their original publications, Garner and Allard sug-gested that plants responded to the relative lengthsof day and night and coined the term photoperiodism,which combines the Greek roots for light and dura-tion. Photoperiodism turns out to be a misleadingterm, however, because it implies that plants mea-sure the duration of daylight. In fact, plants measureneither the relative length of day and night nor thelength of the photoperiod—they measure the lengthof the dark period. This was elegantly demonstratedby the experiments of K. C. Hamner and J. Bonnerin 1938 (Figure 24.4). Under 24-hour cycles of lightand dark, Xanthium flowered with dark periods longerthan 8.5 hours but remained vegetative on schedules of

418 Chapter 24 / Measuring Time: Controlling Development by Photoperiod and Endogenous Clocks

VegA

B

C

D

16 (8)

8 (16)

4 (8)

16 (32)

Flower

Veg

Flower

FlowerE 15 (9)

VegF 16 (8)

VegG 15 (9)

VegH 16 (8)

FIGURE 24.4 The central role of dark period in Xan-thium strumarium, a SD plant. The photoperiod regimeis shown to the left. The number enclosed in bracketsindicates the length of the dark period. Note that theplants flower whenever the dark period is uninterruptedfor 9 hours or more.

16 hours light and 8 hours darkness (Figure 24.4A,B).On schedules of 4 hours light and 8 hours darkness,plants remained vegetative even though the 4-hourphotoperiod is much shorter than the 15.5-hour criticalphotoperiod (Figure 24.4C). On the other hand, sched-ules of 16 hours light and 32 hours darkness inducedrapid flowering even though the photoperiod exceededthe critical daylength (Figure 24.4D).

The results of the Hamner and Bonner experimentallow two conclusions. First, the relative length of dayand night is not the determining factor in photoperi-odism, because the ratio of light to dark is the same inschedules B, C, and D (Figure 24.4) but with differentresults. Second, it is the length of the dark period that isimportant. The consistent feature within the experimentis that Xanthium will flower whenever the dark periodexceeds 8.5 hours and will remain vegetative wheneverthe dark period is less than 8.5 hours. The critical role ofthe dark period was confirmed by interrupting the darkperiod with brief light exposures (Figure 24.4 E–H).The flowering effect of an inductive 9-hour dark periodcan be nullified by interrupting the dark period with abrief light-break (Figure 24.4G), but a ‘‘dark interrup-tion’’ of a long light period has no effect (Figure 24.4H).Experiments with LD plants give similar results; LDplants require a dark period shorter than some criticalmaximum. With LD plants, a light-break in the mid-dle of an otherwise noninductive long dark period willshorten the dark period to less than the maximum and

permit flowering to occur. At this point it is clear thatphotoperiodism has relatively little to do with daylengthper se. It is instead a response to the duration and timingof light and dark periods. Thus, the critical daylengthfor a SD plant actually represents the maximum lengthof day in a normal 24-hour regime that will allow a darkperiod of sufficient length. In the case of LD plants, longdark periods are inhibitory and the critical daylength isthe minimum in a 24-hour regime that will keep thedark period short enough to allow flowering.

The fluence given during a light-break need not bevery high to be effective. As little as one minute of incan-descent light at a low fluence will prevent flowering inXanthium. Even bright moonlight is sufficient to delayflowering in some SD plants. This raises an interestingpossible relationship between nyctinastic leaf move-ments, discussed in the previous chapter, and controlof flowering. It is possible, at least in some cases, thatnyctinastic leaf movements could serve to reorient theleaves parallel to incident moonlight and thus reduce theimpact of moonlight on the time-sensing mechanism.

24.1.4 PHYTOCHROME ANDCRYPTOCHROME ARE THEPHOTORECEPTORS FORPHOTOPERIODISM

Photoperiodism is a response to the length of a darkperiod, but the length of a dark period is defined bythe interval between photoperiods. In other words, thelength of a dark period is determined by the timing oflight-off and light-on signals. In the case of a SD plantsuch as Xanthium, a light-break given in the middleof an otherwise inductive long dark period, as shownearlier in Figure 24.4, may be construed as a prematurelight-on signal that interferes with the timing process.Light-breaks are useful because they are effective withshort exposures of low-fluence-rate light and can beapplied to a single induced leaf. The light-break thusprovides an opportunity to explore the nature of thepigment involved in photoperiodism by determining anaction spectrum.

Early action spectra on several SD plants in the late1940s indicated that red light was most effective as alight-break, with a maximum effectiveness near 660 nm.Then, at the same time that photoreversibility of seedgermination was demonstrated, H. A. Borthwick andhis colleagues also reported that red light inhibition offlowering in Xanthium was reversible with far-red. Later,similar results were demonstrated for other SD plants.Red, far-red photoreversibility of the light-break clearlyimplicates phytochrome in the photoperiodic timingprocess. The situation with LD plants, however, is notas clear. A light-break with red light in the middle ofan otherwise noninductive long night should promoteflowering. It does in Hyoscyamus and some others, but

24.1 Photoperiodism 419

many LD plants are not sensitive to light-breaks andsome are inhibited by red light. The role of phytochromeis far from clear at this point, but based on recent workwith phytochrome mutants in Arabidopsis, it has beensuggested that phyA is required to promote flowering ofan LDP under certain conditions. PhyB, on the otherhand, seems to inhibit flowering.

Blue light is also effective in controlling photope-riod. In Arabidopsis, for example, blue light is effectiveat promoting flowering when applied in a light-breakexperiment, suggesting that cryptochromes are involvedas well. It is not yet clear what role cry1 may have inphotoperiodic control of flowering. On the other hand,cry2 is known to promote flowering, but is believed tointeract directly with the biological clock. More will besaid about the role of cry2 later in this chapter, when weturn our attention to the biological clock.

24.1.5 THE PHOTOPERIODIC SIGNAL ISPERCEIVED BY THE LEAVES

The actual change from the vegetative to reproductivegrowth occurs, of course, in apical meristems—usuallybeginning at the shoot apex and appearing later inthe axillary buds. Contrary to expectations, however,the photoperiodic signal is perceived not by the stemapex but by the leaves. This has been demonstratedin a variety of ways. Some of the earliest experiments

C.

B. A.

Vegetative Flowering

Flowering

SD

SD

SD

LD

LD

LD

FIGURE 24.5 The role of the leaf in perception of the photoperiodic stimulus in theshort-day plant Perilla. (A) Plants remain vegetative when the shoot apex is coveredto provide short days and the leaves are maintained under long days. (B) Plants flowerwhen the leaves are given short days but the meristem is maintained under long days.(C) Flowering will occur when only a single leaf is provided short days.

were conducted by a Russian plant physiologist M.Chailakhyan. He reported flowering in Chrysanthemummorifolium, a SD plant, in which the apical, defoliatedportion was kept on long days but the leafy portionwas subjected to short days. When conditions werereversed, with the upper, defoliated portion kept onshort days and the leafy portion on long days, the plantsremained vegetative (Figure 24.5). Although the plantsin this experiment still contained axillary buds, floweringhas been successfully induced in plants from which theaxillary buds have been removed. In later experimentsit was shown that plants such as Perilla and Xanthiumstripped of all but one leaf could be induced to flowerif the remaining leaf were provided the appropriatephotoperiod. Leaves may also be removed from inducedplants and grafted to noninduced receptors where theywill induce a flowering response. Finally, leaves neednot even be attached to the plant in order to be induced.When excised leaves of Perilla (SD plant) were exposedto short days and grafted back to noninduced plants, theplants flowered even when maintained under long days.

The sensitivity of the leaf may vary with age.In Chrysanthemum, Perilla, and soybean (Glycine), theyoungest fully expanded leaf was found to be most sen-sitive. In experiments with Xanthium in which the plantsare stripped of all but the most sensitive leaf, it has beenshown that peak sensitivity is reached during the periodof most rapid expansion, when the leaf is about half its

420 Chapter 24 / Measuring Time: Controlling Development by Photoperiod and Endogenous Clocks

final size. These observations lead to two conclusions:first, that the leaf is independently responsible for per-ceiving the phototropic signal, and second, that the leafinitiates a signal chain, probably involving a diffusiblefloral stimulus, that communicates this information tothe shoot apical meristem. Grafting experiments werealso used to estimate velocity of movement of the stimu-lus, which for most species was found to be in the rangeof about 2.5 to 3.5 mm hr−1.

24.1.6 CONTROL OF FLOWERING BYPHOTOPERIOD REQUIRES ATRANSMISSIBLE SIGNAL

The spatial separation between the site of perceptionof the photoperiodic signal (the leaves) and the site offlowering (the shoot apical meristem) logically requiresa transmissible signal that carries the information fromthe leaf to the shoot apex. Over the past 70 years, severalhypotheses have been advanced in order to explain thetransmission of a floral stimulus, but the most persistenthas been the hypothesis of a floral hormone.

Russian plant physiologist M. Chailakhyan was thefirst to suggest, in 1936, that the floral stimulus might bea hormone, which he proposed be called florigen. Theconcept of a flowering hormone was based primarilyon the fact that the stimulus was transmissible acrossa graft union. For example, when several Xanthiumplants are approach-grafted in sequence, all can bebrought to flower if only the first is induced by shortdays (Figure 24.6). Members of the same family, suchas the SD plant tobacco (Nicotiana tobacum) and the LDplant black henbane (Hyoscyamus niger), both in the fam-ily Solanaceae, can be grafted with relative ease. In sucha partnership, Hyoscyamus will flower under short days ifthe tobacco is also maintained under short days, but notif the tobacco is maintained under long days. Conversely,tobacco will flower under long days when grafted toHyoscyamus maintained under long days. A numberof successful interspecific and intergeneric grafts haveyielded similar results. These results have led to the con-clusion that the final product of photoperiodic inductionappears to be physiologically equivalent in plants of dif-ferent photoperiodic classes and is probably identical tothe constitutive floral stimulus in day-neutral plants.

Given the universal nature of other plant hormones,it should not be too surprising that the same floral stim-ulus operates in all photoperiodic classes. The majorunanswered question that remains, however, is withregard to the chemical identity of the florigen. Oneapproach toward answering this question is to preparefrom flowering plants an extract that will evoke flower-ing in noninduced plants. Subsequent fractionation ofthe extract should lead to identification of the active sub-stance. Unfortunately, although several attempts havebeen made, they have met with limited success. In one

Induced leaf

SD

FIGURE 24.6 Transmission of the floral stimulus ingrafted plants. Several plants are ‘‘approach’’ grafted andthe terminal plant is induced to flower. All plants willflower, indicating that the floral stimulus has been trans-mitted from the single induced leaf through all of theplants.

of the most successful attempts to date, a methanolextract from freeze-dried Xanthium plants evoked aweak flowering response when applied to leaves of otherXanthium plants kept under long days. Unfortunately,few other attempts have been successful and none hasbeen consistently repeated.

Of the several classes of plant hormones, onlygibberellins have been shown consistently to evoke flow-ering in a wide variety of species. Repeated applicationsof dilute gibberellin solutions (containing principallyGA3) to the apex of annual Hyoscyamus, Samolus parv-iflorus, and Silene armeria (all LD plants) elicited aflowering response under short days. Does this meanthat gibberellin is equivalent to florigen? The answer tothis question is clearly negative. To begin with, evoca-tion of flowering in response to gibberellin applicationis almost entirely restricted to LD plants that normallygrow as rosettes under short days. This includes annualLD plants and biennial species that require an overwin-tering cold treatment before flowering as LD plants (seeChapter 25). For example, carrot (Daucus carota), Chi-nese cabbage (Brassica pekinensis), and biennial strains ofblack henbane (Hyoscyamus niger) all grow as rosettesand remain vegetative during the first growing season.The meristems are then subjected to an over winter-ing cold treatment. The following spring, the stemsundergo rapid internode elongation and the plants willflower in response to long days. In the absence of anycold treatment but under long days, exogenously appliedgibberellin will stimulate stem elongation and flower-ing. It thus appears that gibberellin will substitute forthe cold requirement of biennial species or the long-dayrequirement of annual LD plants. But gibberellin willnot evoke flowering in most SD plants (such as Xanthiumor Biloxi soybean) kept under long days.

24.1 Photoperiodism 421

Molecular genetic studies with Arabidopsis, a fac-ultative long-day plant, have recently cast a differentlight on the nature of florigen. A number of experi-ments conducted over the last decade have indicated aprominent role for the gene CONSTANS (CO). It wasfirst observed that plants carrying mutant alleles of COflower late under long days, but flower normally undernoninductive short days. It was also found that COmRNA accumulated in leaves under long days as wellas in vegetative meristems and leaf initials. However,in spite of these and other observations, it was notclear whether CO stimulated flowering by acting inthe leaves where the photoperiod signal is perceivedor in the meristem where flowering occurs. The keyproved to be experiments with transgenic Arabidopsisplants in which the CO gene was linked to a pro-moter from melon (Cucumis melo)—a promoter thatis active only in the phloem companion cells in theminor veins of mature leaves. The transgenic plantsexpressing the CO gene flowered early under nonin-ductive short-day conditions. Early flowering was alsoobserved in transgenic co mutant plants that were engi-neered to constitutively express the CO gene. Finally, itwas demonstrated that when scions homozygous for theco mutation were grafted to CO-expressing stock, earlyflowering was observed in the scions. Other experimentsdesigned to test this idea have provided no evidence thatCO itself is translocated out of the leaf cells Theseresults strongly indicate that the CO gene participates ingenerating a phloem-mobile transmissible floral signal.

A likely candidate for the mobile floral signalis FLOWERING LOCUS T (FT), a small protein(ca. 20 kD) that is known to promote flowering in adosage-dependent manner. The CO gene encodes atranscription factor that induces the expression of FTmRNA in leaves. The presence of RNA and peptides inphloem sap is well documented and in a recent study ofBrassica napus, FT was one of 140 proteins identified inthe phloem sap. According to one current model, COresponds to an inductive photoperiod by activating FTmRNA transcription and FT protein synthesis in leafphloem parenchyma cells (Figure 24.7). The FT proteinthen moves through the phloem sieve tube elements tothe shoot apical meristem where it induces a group ofgene known as floral meristem identity genes. Therole of floral identity genes will be discussed further inChapter 25.

24.1.7 PHOTOPERIODISM NORMALLYREQUIRES A PERIOD OF HIGHFLUENCE LIGHT BEFORE ORAFTER THE DARK PERIOD

Although a SD plant will flower in response to a singlelong dark period, to be most effective the inductivedark period must be preceded by a period of light. The

Flowering genes

Shoot ApicalMeristem

PhytochromeCryptochrome

FT

CO

FT FT FT

FIGURE 24.7 A model for the long-distance transport offloral stimulus. The photoperiod is measured in the leafphloem parenchyma cells where CO is activated by phy-tochrome and cryptochrome. Activated CO induces thetranscription of the FT gene. FT protein is translocatedthrough the phloem sieve tube cells to the shoot apicalmeristem. Once in the meristem, FT interacts with tran-scription factors to induce the transcription of floweringgenes.

function of the pre-dark light requirement is not clearsince the requirements vary markedly depending on thespecies and conditions of the experiment. For maximumflowering with a single inductive dark period, Xanthiumrequires 8 to 12 hours of light. Pharbitis requires atleast 6 hours. In others such as Kalanchoe, a few secondsof light per day were sufficient to induce flowering.Where longer light periods are necessary, it may bebecause photosynthetic products are required for theprocesses initiated in the dark period. Where very briefperiods of light are effective, clearly photosynthesiscannot be involved and some other explanation for thelight requirement must be sought.

In many cases it has been demonstrated thathigh-fluence light following the inductive dark periodis also important. Again the experimental details aresketchy and the requirements seem to vary, but twoexplanations have been offered. One possibility is thatthe postinductive light period provides a stream ofphotosynthate that enhances translocation of the floralstimulus out of the leaf. However, some of the resultsare not readily explained as translocation effects. Thereis also evidence suggesting that the floral stimulus issubject to destruction or inactivation in the leaf if thedark period is too long. Consequently, it has beensuggested that light may be required to stabilize orprevent inactivation of the floral stimulus in the leaf.

422 Chapter 24 / Measuring Time: Controlling Development by Photoperiod and Endogenous Clocks

BOX 24.2HISTORICALPERSPECTIVES:THE BIOLOGICALCLOCK

The possibility that an internal, or endogenous, time-keeper might be involved was first raised by the Frenchastronomer M. De Mairan in 1729. De Mairan foundthat leaf movements in the sensitive plant (Mimosa) per-sisted even when the plants were placed in darkness forseveral days. Subsequently studies by J. G. Zinn (in 1759)and A. P. De Candolle (in 1825) confirmed DeMairan’sfindings (see Sweeny, 1987). Curiously, DeCandollefound that under continuous light the time betweenmaximum opening of the leaves was closer to 22 or 23hours rather than the 24 hours under natural conditions.It would be a full century before the significance of thisfinding was fully appreciated!

The study of leaf movements continued to interestbotanists and plant physiologists through the latter halfof the eighteenth and the early nineteenth centuries.Much of the work simply confirmed the widespreadoccurrence of leaf movements and that they persistedunder either continuous light or continuous darkness.In 1863, J. Sachs reported no correlation between leafmovements and temperature fluctuations, thus elimi-nating temperature as a cause. One difficulty, from anexperimental perspective, was that studies of periodicphenomena required around-the-clock monitoring. In1875, W. Pfeffer devised an apparatus for automatic andcontinuous recording of leaf position. Pfeffer attachedthe leaf, via a fine thread, to a stylus, which in turnrecorded the position of the leaf on a rotating drumcoated with carbon (lampblack). With some improve-ments, the same apparatus is occasionally used eventoday (Figure 24.8).

Over a period of 40 years, from 1875 to 1915, Pfeffercontributed several papers devoted to leaf movementsin Phaseolus vulgaris, the common garden bean. At onepoint, he showed that plants that had lost their rhythmicleaf movements (as they will under prolonged continu-ous light or darkness) will regain them if exposed to anew light–dark cycle. If the new cycle is inverted withrespect to natural day and night, leaf movement willalso be reversed. Pfeffer concluded that persistent leafmovements under continuous light or darkness were a‘‘learned’’ behavior. Others showed that regardless ofany previous light–dark cycle, under continuous illumi-nation sleep movements clearly reverted to a 24-houroscillation. In the end, Pfeffer was forced to concludethat leaf movements were an endogenous, and probablyinherited, behavior.

SL

FIGURE 24.8 A diagram illustrating the principle of thedrum recording apparatus used by Bunning and oth-ers for recording leaf movements. The recording stylus(S) is attached to a finely balanced lever (L), which is inturn tied to the midvein of the leaf. As the leafchanges position, the stylus describes a tracing on therotating drum.

During the 1920s, improvements in the technol-ogy for maintaining constant environments, especiallywith respect to light and temperature, set the stage forsignificant advances in understanding leaf movements.One key observation was made by Rose Stoppel inGermany. Maintaining bean plants in a dark room atconstant temperature, Stoppel observed that maximumnight position of the leaves (i.e., near vertical orienta-tion) occurred at the same time (between 3:00 and 4:00A.M.) every day, exactly 24 hours apart. She reasoned thatan endogenous, biological timer could be that accurateonly if some environmental factor acted to, in effect,reset the clock on a daily basis. Stoppel referred to thisfactor as factor X.

The endogenous nature of the biological clockfinally became evident through the work of two youngbotanists who had been given the task of determin-ing whether subtle atmospheric factors might influenceplants. E. Bunning and K. Stern became interested inStoppel’s factor X and, in order to achieve satisfactoryconstant temperature conditions, set up their bean plantsand recording devices in Stern’s potato cellar. LikeStoppel, Bunning and Stern found that the maximumnight position came every 24 hours, but, surprisingly,some 7 to 8 hours later than in Stoppel’s experiments!Bunning and Stern quickly recognized that the key wasthe very weak red light used when watering the plantsand tending the recording equipment. Stoppel visitedher experiments early in the morning while Bunning andStern, because Stern’s potato cellar was some distance

24.2 The Biological Clock 423

A.

B.

FIGURE 24.9 Leaf movement inbean (Phaseolus vulgaris) is a mani-festation of an endogenous rhythmgenerated by the biological clock.(A) A 6-day record of primary leafposition with alternating light anddark periods. Light and dark barsrepresent light and dark periods,respectively. The vertical lines indi-cate midnight solar time. Periodlength is 24 hours. (B) Free-runningrhythmicity under continuouslight. After the first period, theperiod length is extended to 25.7hours. Note also that the ampli-tude (dashed lines) diminishes withtime under continuous light. (Fromthe data of Bunning, E. M. Tazawa1957. Planta 50:107.)

from the laboratory, made their visits in the late after-noon. Interestingly, at that time the textbook dogmawas that red light had no effect on plant morphogene-sis. Bunning and Stern concluded that the dogma waswrong—even weak red light apparently synchronizedleaf movement so that the maximum night position

always occurred about 16 hours later. Indeed, when thered light was eliminated, the period between maximumnight positions was no longer 24 hours but 25.4 hours(Figure 24.9). Bunning and Stern had identified factorX and lay the groundwork for the eventual discovery ofthe role of phytochrome in circadian rhythms.

Actually, most of the data fit a combination of these twoideas and the effect of the postinduction light require-ment may be to quickly move the stimulus, such as FTprotein, away from a site of inactivation in the leaf.

24.2 THE BIOLOGICAL CLOCK

Photoperiodism is inextricably linked to an internaltime-measuring system known as the endogenous bio-logical clock (Box 24.2). Another long-observed anddramatic manifestation of the biological clock is the diur-nal rise and fall of leaves (known as nyctinastic, or sleep,movements) (Figure 24.10 and Chapter 23). Superfi-cially, nyctinastic movements appear to be subject toexternal, or exogenous, control—namely the daily pat-tern of light and dark periods. However, under the rightcircumstances, it can be shown that the oscillations inleaf position are independent of external factors. Onthe other hand, photosynthetic carbon uptake describesa periodicity because it is light-driven and daylight isperiodic over time. Photosynthesis is thus diurnal. Itis active only during daylight hours and thus mirrors thedaily light-dark cycle.

There are three criteria that serve to distinguishbetween simple diurnal phenomena and rhythms drivenby an endogenous clock. (1) A clock-driven rhythm

persists under constant conditions (that is, in the absenceof external cues). (2) A clock-driven rhythm can be‘‘reset’’ by external signals, such as light or temperature.(3) There is no lasting effect of temperature on thetiming of the clock-driven rhythm. These three criteriawill be addressed in turn.

24.2.1 CLOCK-DRIVEN RHYTHMSPERSIST UNDER CONSTANTCONDITIONS

The key to an endogenous rhythm is that it persists,for at least several cycles, under constant conditions(usually constant light or constant darkness). The rhyth-micity expressed under constant conditions is describedas free-running. The time required to complete a cycleis referred to as the period (τ; tau) (Figure 24.11A).Period is conveniently described as the time from peakto peak, but it applies equally well to any two compara-ble points in the repeating cycle. Biological rhythms aretraditionally classified according to the length of theirfree-running period. Thus a circadian rhythm has aperiod of approximately 24 hours (circa, about + diem,day). Bean leaf movement is a circadian rhythm becauseits free-running period length is about 25.4 hours.A period of about 28 days, the time between one fullmoon and the next, describes a lunar rhythm and a

424 Chapter 24 / Measuring Time: Controlling Development by Photoperiod and Endogenous Clocks

A.

B.

FIGURE 24.10 Circadian clock-induced movements ofbean leaves. Seedlings of bean (Phaseolus vulgaris) areshown with the primary leaf in the horizontal day posi-tion (A) and vertical night position (B).

FIGURE 24.11 Examples of cir-cadian oscillations. (A) Tworhythms with period (τ ) andamplitude (A). Although theperiod and amplitude for bothrhythms are the same, thetwo are slightly out of phase.(B) Free-running rhythm andentrainment. A free-runningrhythm with a period of 28 hours(dashed line) is entrained to 24hours by a daily light-on signal.The open and closed bars repre-sent light and dark conditions.

B.

A.

A

period of one year is an annual rhythm. Also of interestare rhythms in metabolic activity with periods substan-tially less than 24 hours (measured in minutes or hours).These are known as ultradian rhythms.

The difference between the maximum and mini-mum, or peak and trough, of a rhythm is known asthe amplitude (A) (Figure 24.11A). The amplitude of afree-running circadian rhythm usually diminishes withtime until it eventually disappears altogether. In somecases this is probably due to declining energy reservesin prolonged darkness since the amplitude can be main-tained, at least for a while, by feeding sucrose. Moreoften, however, the clock seems to run down and anexternal signal is required to start up the rhythm again.The term phase has two slightly different but relatedusages. Any point of the cycle that can be identified byits relationship to the rest of the cycle can be consid-ered a phase. The position of the peak (maximum nightposition of leaves, maximum flowering, etc.) is the mostcommon reference point for phase relationships becauseit is usually most readily identified. In Figure 24.11A,for example, the two rhythms are displayed out of phaseby approximately 6 hours. Phase may also be used todescribe an arbitrary part of the cycle, such as nightphase or day phase.

Discussions of endogenous circadian rhythms aresometimes complicated by the fact that two time framesare involved: solar time and circadian time (CT).Solar time is an external time, based on a normal24-hour day. Circadian time, on the other hand, is aninternal time and is based on the free-running period.One cycle is considered to be 24 hours long, regardlessof its actual length in solar time. Each hour of circadiantime is therefore 1/24 of the free-running period. Thusif the free-running period is 30 hours, events that occurat 0, 15, and 30 hours of darkness will have occurred atcircadian times CT:0, CT:12, and CT:24, respectively.The circadian time scale is useful in assessing phase

24.2 The Biological Clock 425

relationships within experiments or between rhythmswith different periods. Finally, the phase of thefree-running cycle that corresponds to day in a normallight–dark environment is known as subjective dayand that which corresponds to normal night is thesubjective night. In the case of sleep movements inbean, for example, the phase of the free-running cycleduring which the leaves are in the horizontal positionwould be considered the subjective day. The phaseduring which the leaves are in the vertical positionwould be subjective night.

The rhythmic movements of bean leaves are nor-mally coupled, or synchronized, to the solar day–nightcycle (Figure 24.11B). The same coupling was evidentin both Stoppel’s and Bunning’s experiments when therhythm was coupled to the red light signal (Box 24.2).Such a coupling of a circadian rhythm to a regular exter-nal environmental signal is known as entrainment. Thesignal that synchronizes or entrains the rhythms is oftenreferred to as a zeitgeber (Ger. zeit, time + geben, togive).

A large number of circadian rhythms have now beendescribed in a wide range of organisms. The list includessingle-celled flagellates, algae, fungi, crustacea, insects,birds, and mammals (including humans), in additionto flowering plants. A full list of known rhythms inflowering plants alone would cover several pages. Anabbreviated list is provided in Table 24.2. A more exten-sive list, with references, is provided by Sweeny (1987).

24.2.2 LIGHT RESETS THE BIOLOGICALCLOCK ON A DAILY BASIS

The action of the biological clock or endogenousrhythms is to ensure that certain functions occur at aparticular time of day. For example, the oscillations ofthe clock in beans determines that the leaves rise duringthe day and fall at night. The period of the endogenousrhythm is fixed, but it may be ‘‘fast’’ or ‘‘slow’’ relativeto the 24-hour solar period. Moreover, the dailyduration of light and dark within the 24-hour solar

TABLE 24.2 Examples of circadian rhythmicphenomena in higher plants

Rhythm Organism

Sleep movements Many speciesStomatal opening Banana, tobacco, ViciaStem growth Tomato, ChenopodiumCO2 production Orchid flowersGas uptake Dry onion seedsMembrane potential Spinach leavesmRNA expression Pea

period changes steadily throughout the season. Howdoes the organism reconcile a nonvarying endogenousperiodicity with these daily and seasonal changes indaylight? How are these rhythms kept in phase? Toanswer this question, we go back to the concept ofentrainment. Entrainment is, in effect, a means formoving the oscillations of the clock forward or back intime on a daily basis, just as you might reset the time ofan alarm clock every night before retiring. Entrainmentis also useful to the experimentalist because it is theone significant way in which circadian rhythms can bemanipulated in the laboratory.

Entrainment is not limited to solar periodicity.Within limits, rhythms can be entrained to light–darkcycles either shorter (18 to 20 hours) or longer (upto 30 hours or more) than 24 hours. Entrainment toextremely short or long cycles is rare. More useful infor-mation, however, can be obtained by studying whetherthe rhythm is moved forward or back in time and byhow much. The experiment normally entails givingbrief light pulses (usually 1 hour or less) at various timesduring an established free-running rhythm in constantdarkness. The timing of the next peak is then comparedwith controls that have not been given a light pulse. Forexample, when populations of Chenopodium seedlingsare exposed to single dark periods of various lengths,their capacity to flower fluctuates rhythmically for atleast three cycles (Figure 24.12). In this case, a commonlight-off signal sets the rhythm in motion and the timingof the light-on signal (i.e., the length of the dark period)determines whether the plants will flower. If, however,relatively brief pulses of light are given at various timesduring the dark period, the pulses will reset the clockor shift the phase of the rhythm. The result is a phaseresponse curve such as that shown in Figure 24.13.This curve demonstrates that a light pulse given earlyin the subjective night causes a delay of the first andsubsequent peaks relative to the control. Somewherenear the middle of the subjective night there is a phasejump such that pulses given in the latter half of thesubjective might cause subsequent peaks to be advancedrelative to the controls. Note that light pulses givenduring the subjective day have very little effect on phaserelationships. Similar phase response curves have beendemonstrated for a variety of circadian rhythms and helpto explain entrainment. It seems to be a character of thesystem that re-phasing is accomplished in such a way asto require the least net displacement of the rhythm. Thus,during the early part of the subjective night, the lightpulse is apparently interpreted as a delayed light-off, ordusk, signal and the phase of the endogenous rhythmis adjusted accordingly. As the pulse arrives later, thedelay is increased until at some point the pulse is nowinterpreted as an early light-on, or dawn, signal. Thiscauses the rhythm to be advanced. Phase-shifting in thisway constantly adjusts or entrains the rhythm to local

426 Chapter 24 / Measuring Time: Controlling Development by Photoperiod and Endogenous Clocks

Length of dark period (hours)

20 40 60 820

20

40

60

80

100

% o

f po

pula

tion

flo

wer

ing

FIGURE 24.12 Flowering in the SD plant Chenopodiumrubrum responds rhythmically to the length of a singledark period. Populations of C. rubrum seedlings wereexposed to a single dark period of varied length as indi-cated. The free-running periodicity (τ ) is approximately30 hours. The amplitude diminishes because the youngseedlings are depleting their carbon supply over theextended dark period. Amplitude can be maintained bysupplying the seedlings with glucose. (From King, R. W.Time measurement in photoperiodic control of flower-ing, Ph.D. thesis, University of Western Ontario, 1971.With permission of R. W. King.)

solar time. The observation that similar phase responsecurves can be described for such different phenomena asinsect pupal eclosion, bioluminescence in the dinoflag-ellate Gonyaulax, and CO2 evolution and flowering inhigher plants indicates similar properties, if not mech-anisms, for the circadian clock in a variety of differentorganisms.

Although light plays an obvious role in resettingcircadian rhythms, the photoreceptor involved is clearlynot the same in all cases. Action spectra for resettingrhythms in Gonyaulax, the protozoan Paramecium, fungi,and insects all share a large peak in the blue region ofthe spectrum, suggesting that a flavoprotein blue-lightreceptor such as cry2 might be involved. In higher plants,however, both phytochromes and cryptochromes appearto be involved. For example, establishment of rhythmsin dark-grown bean plants and phase-setting of leafmovement in Samanea both show a classic phytochromephotoreversibility with brief red and far-red light treat-ments. Others, however, such as the CO2-evolutionrhythm in Bryophyllum leaves, can be reset with red lightbut the effect is not reversible with far-red. The effect offar-red light in the Bryophyllum system is to abolish therhythm altogether! This might also be a phytochromeeffect, but these and other experiments make it clear that

Pha

se a

dvan

ce o

r de

lay

(hou

rs)

Subjective night

0 6 12 18

–10

–5

+5

+10

24

Subjective day

Time of light pulse

FIGURE 24.13 Phase response curve. Light-on signalsduring the early part of the subjective night cause a delayin the timing of the next peak; given late in the subjectivenight, the result is an advance of the next peak. Light-onsignals given during the subjective day have little or noeffect on the phase of the rhythm. (Redrawn from King,R. W. Time measurement in photoperiodic control offlowering, Ph.D. thesis, University of Western Ontario,1971. With permission of R. W. King.)

photocontrol of phase-setting is not a straightforwardprocess.

24.2.3 THE CIRCADIAN CLOCK ISTEMPERATURE-COMPENSATED

Most chemical reactions, and thus growth and otherbiological responses, respond to temperature with a Q10near 2. This means that a 10◦C increase in tempera-ture will approximately double the rate of the process.A decrease in temperature leads to a decrease in the rateby the same amount. While such temperature sensitivitymay be advantageous to the organism in some cases, theaccuracy of a biological clock would be severely compro-mised if it were sensitive to often-random temperaturefluctuations brought about by local environments. Asit turns out, the period length of circadian rhythms isrelatively insensitive to temperature. A classic exampleis again bean leaf movement. When seedlings are raisedin the dark from seed, leaf movements tend to be smalland unsynchronized. A single flash of light (the zeitgeber)initiates larger, synchronized movements.

In Bunning’s experiments, the synchronized move-ments had a periodicity of 28.3 hours at a constant15◦C and 28 hours at 25◦C. Although these data would

24.2 The Biological Clock 427

seem to suggest that the circadian rhythm is insen-sitive to temperature, this is not strictly true. Whenseedlings were shifted from 20◦C to 15◦C, the initialperiod was 29.7 hours. Seedlings shifted from 20◦C to25◦C had a period of 23.7 hours. These periods, how-ever, lasted only for the first cycle or two—later cyclesreturned to periods of approximately 28 hours. Clearlythe circadian rhythm is temperature-sensitive, but somemechanism quickly compensates for variations in tem-perature. Consequently, the Q10 for most circadianrhythms is near 1. Amplitude may be affected by tem-perature, but temperature-compensation is clearly acharacteristic of the period for most circadian rhythms.

24.2.4 THE CIRCADIAN CLOCK IS ASIGNIFICANT COMPONENTIN PHOTOPERIODIC TIMEMEASUREMENT

Endogenous rhythms of all kinds are fundamentally aquestion of time measurement, a concept that is noteasily imagined within the framework of conventionalbiochemistry. Moreover, the clock is exclusively internaland, except for resetting by light and temperature, isnot generally subject to manipulation from the outside.This means the clock is not amenable to traditionalexperimental strategies, since these normally requirethat the investigator be able to control or manipulate thesystem in some way. Another difficulty is our inabilityto distinguish between oscillations that are part of thetimekeeping mechanism and those that are simply the‘‘hands’’ responding to the output of the basic oscillator.

In 1936, E. Bunning first proposed that photoperi-odism was tied to circadian rhythms. Bunning proposedthat the rhythm was comprised of two phases—thephotophile, or light-loving phase, and the scotophile,or dark-loving phase—which alternated about every12 hours. According to Bunning’s hypothesis, lightfalling on the plant during the photophile phase wouldpromote flowering while light during the scotophilephase would inhibit flowering. In most experimentalsituations, when the plant is placed under continuousconditions, the photophile phase would probably beequivalent to subjective day and the scotophile phaseequivalent to subjective night. In order to demonstraterhythmicity in flowering and test Bunning’s hypoth-esis, several novel experimental strategies have beenemployed. The difficulty is that, unlike leaf movement orcarbon dioxide evolution, flowering is not a continuousprocess.

One strategy to demonstrate rhythmicity inflowering is illustrated by the SD plant Chenopodiumrubrum, described in the previous section. Seedlings ofChenopodium can be induced to flower with a single darkperiod when the seedlings are only 41/2 days old. Beforeand after the dark period, the seedlings are maintained

under continuous light. As shown in Figure 24.12, whenthe length of the dark period is varied, light duringthe first 8 to 10 hours inhibits flowering as would beexpected with a SD plant. Thereafter, the capacity of theseedlings to flower as a function of the length of the darkperiod expresses a rhythmic pattern for at least threecycles, with a free-running period of 30 hours. Withminor variations in the experimental strategy, essentiallythe same results have been demonstrated by other inves-tigators using a variety of plants including cocklebur,soybean, Japanese morning glory, and duckweed.

A second example of the close dependency ofphotoperiodic time measurement on rhythmic phasescomes from a series of elegant experiments conductedby W. S. Hillman with the aquatic plant Lemna pur-pusilla (Figure 24.14). Lemna purpusilla is a SD plantwith a critical daylength of 12 hours. As an experi-mental organism, Lemna offers the unique advantageof growing heterotrophically in darkness when sup-plied with glucose. This means that long light periodscan be eliminated and timing can be controlled byskeleton photoperiods—short pulses of light that serveto mark the beginning and end of dark periods. Allthat is required to induce flowering in Lemna aretwo 15-minute (0.25 h) light periods every 24 hours(Figure 24.15). Thus, when taken from continuous light toa schedule of 13 hours dark: 0.25 hours light: 10.5 hoursdark: 0.25 hours light, Lemna will flower. On a scheduleof 10.5 hours dark: 0.25 hours light: 13 hours dark:

FIGURE 24.14 Duckweed (Lemna sp.). The genus Lemnais the smallest flowering plant. Each leaflike frond isabout 1 to 2 mm across. Flowers arise at the point of thefrond and consist of male and female parts only (no petalsor sepals). There are short-day and long-day species ofLemna.

428 Chapter 24 / Measuring Time: Controlling Development by Photoperiod and Endogenous Clocks

FIGURE 24.15 Skeletal photoperiods andthe control of flowering in Lemna pur-pusilla, a SD plant. Lemna is arrhythmicunder continuous light. The first light-offsignal (arrow) starts up the rhythm.Because the first phase is the scotophilephase, Lemna interprets the first darkperiod, regardless of its length, as night.(A) If the ‘‘night’’ dark period exceeds thecritical value, the plants will flower; if not(B), the plants remain vegetative. (C) Thehypothetical scotophile and photophilephases of the rhythm.

"Night"

0

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B.

(13)

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"Night" "Day"

(10.5) (13) (10.5) (13)

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12 24 36 48 60

Scotophilephase

Photophilephase

0.25 hours light, however, the plants remain vegetative.Note that the only difference between the two schedulesis the length of the first dark period following continuouslight. Even though both schedules contain a 13-hourinductive dark period, flowering is induced only whenthe long dark period comes first.

Clearly, Lemna recognizes the brief light pulses asthe beginning and end of two different dark periods,but how does the plant know which is which? As withother plants under continuous light, Lemna becomesarrhythmic; that is, the amplitude of the rhythm dampsout until the rhythm disappears. Transfer of the plants todarkness starts up the rhythm. Since flowering occurredonly when the first dark period exceeded the criticalnight length, the first dark period (and every alternateperiod thereafter) must have been interpreted as night.This appears to be a general rule—the light-off signalfollowing a period of continuous light starts up therhythm and the first dark period (CT 0 to 12) is alwaysthe night phase. The second phase (CT 12 to 24)is the day phase. In subsequent experiments, Hillmanconfirmed that flowering occurs only when a dark periodlonger than the critical night length coincides with thenight phase of its circadian rhythm. It is interesting tonote that this interpretation is not limited to floweringplants—skeleton photoperiods elicit similar responseswith respect to photoperiodic effects in insects and birds.

24.2.5 DAYLENGTH MEASUREMENTINVOLVES AN INTERACTIONBETWEEN AN EXTERNAL LIGHTSIGNAL AND A CIRCADIANRHYTHM

Several models have been proposed to explain theintegration of photoperiodic time measurement withthe biological clock. The model that is most consistentwith both the earlier physiological studies and morerecent genetic evidence is the external coincidencemodel. This model is essentially an updated version

of Bunning’s scotophile and photophile hypothesis(Figure 24.16). Keep in mind that this model has beendeveloped primarily for Arabidopsis, which is a facultativelong-day plant. In its simplest form, the externalcoincidence model separates time measurement intotwo interacting components: the circadian clock itselfand a clock-regulated daylength measuring mechanism.According to this model, the circadian clock sets alight-sensitive phase within the light–dark cycle. It doesthis by controlling the cyclic output of a key regulator

Regulator

FloweringGenes

Long day Short day

FIGURE 24.16 The external coincidence model explainsdaylength measurement in long-day plants. A criticalregulator gene is expressed maximally in late afternoon.Because the function and stability of the regulator pro-tein, whose function is to turn on flowering genes, isregulated by light, the downstream expression of flow-ering genes will occur only under long days when themaximum expression of the regulator coincides with thepresence of daylight. The vertical dashed line representthe day to night transition. The model also applies toshort-day plants if it assumed that the regulator inhibitsflowering.

24.2 The Biological Clock 429

such that it reaches a maximum concentration in lateafternoon. In a long-day plant, the regulator is ulti-mately responsible for turning on flowering genes in theshoot apical meristem but in order to initiate this chainof events, the regulator must first be activated by light.Consequently, flowering will be accelerated only whenthe late-afternoon expression of the regulator coincideswith the presence of daylight, i.e., under long days.Under short days, the regulator does not reach max-imum concentration until well after the beginning ofthe dark period. Consequently, the regulator can not belight-activated and is thus unable to initiate the cascadeof events that leads to transcription of the floweringgenes.

Although relatively little information is available forshort-day plants, the external coincidence model wouldapply if it is assumed that the regulator inhibits, ratherthan promoting, flowering in short day plants.

24.2.6 THE CIRCADIAN CLOCK IS ANEGATIVE FEEDBACK LOOP

A basic model for the circadian clock system in plantsrequires three components: input pathways, a centraloscillator, and output pathways (Figure 24.17). In plants,the principal input pathway is mediated by the photore-ceptors phytochrome and cryptochrome, which set thephase of the oscillator in response to an external timecue (e.g., daylength). The output pathways connect theoscillator to other physiological processes whose overtrhythms reflect the timing of the central oscillator.Most output pathways are believed to be mediated byclock-controlled genes.

The strategy for the genetic dissection of the circa-dian clock is first to identify genes whose transcriptionis controlled by the clock and thus exhibits circadianoscillations. One then looks for mutants that influencethe expression of the clock-controlled genes. If a mutantcan be identified that influences timing at any level,the wildtype gene can be isolated and its gene productanalyzed for clues to its role in the timing mechanism.

The first examples of clock-controlled genes inplants were those involved in photosynthesis, in particu-lar those encoding the chlorophyll a/b–binding proteinsfound in the light-harvesting complex. Others includethe small subunit of Rubisco, the enzyme Rubisco acti-vase, and, in crassulacean acid metabolism (CAM) plants,the enzyme PEP carboxylase kinase. The chlorophylla/b–binding proteins are encoded by a small family ofgenes (CAB). Expression of CAB, as measured by mRNAlevels, is cyclic. CAB transcription begins to increaseshortly before dawn and reaches a peak a few hourslater. This cyclic expression of CAB has been exploitedto develop a sensitive, rapid, and automated system formonitoring circadian rhythms at the molecular level.This has been accomplished by fusing a reporter gene,

Inputs(light, temperature)

centraloscillator

(the clock)

Rhythmicoutputs

(Flowering andother rhythms)

FIGURE 24.17 The three principal components of a sim-ple circadian system include input pathways, a centraloscillator, and output pathways. In plants, the inputpathway(s) originate with light signals, mediated by phy-tochrome and cryptochrome, that entrain (or, reset) thecentral oscillator and may activate clock components.The oscillator is localized within individual cells and reg-ulates the expression of clock-controlled genes, which inturn regulate overt rhythms. The output rhythms are the‘‘hands’’ of the clock. Multiple-output pathways allow asingle oscillator to control overt rhythms with the sameperiodicity but different phases.

the bioluminescent firefly luciferase (luc), with the pro-moter for one of the chlorophyll a/b–binding proteingenes, CAB2. Luciferase has a sufficiently short life spanso that its activity will accurately reflect the activity ofthe promoter. Because the CAB2 promoter is linked tothe central oscillator, transgenic plants containing theresulting CAB2::luc reporter system will thus emitlight rhythmically and these emissions can be moni-tored with sensitive light-monitoring equipment. Useof the CAB2::luc reporter system has played a large rolein facilitating genetic analysis of the clock.

A number of genes closely related to the clock havenow been described and the list continues to grow. Manyof these were originally identified as flowering-timemutants in Arabidopsis and only later were found tobe associated with the clock. One of the first circa-dian timing mutants to be identified was toc1 (timingof CAB). The gene product TOC1 is localized in thenucleus where it has a role in transcription. The toc1mutation shortens period length for a wide range ofclock-controlled processes. TOC1 transcripts exhibit a24 hour periodicity with a peak in the evening. Twoother clock-associated genes are CIRCADIAN CLOCKASSOCIATED 1 (CCA1) and LATE ELONGATEDHYPOCOTYL (LHY ). CCA1 is a transcription factor

430 Chapter 24 / Measuring Time: Controlling Development by Photoperiod and Endogenous Clocks

that was discovered because of its capacity to bindwith the CAB promoter in Arabidopsis. It appears tobe a key element in the signal transduction pathwaylinking phytochrome with the expression of CAB. How-ever, expression of CCA1 itself cycles with a 24-hourperiod, peaking in the morning soon after dawn. Thereare several reasons for believing that CCA1 is closelyassociated with the circadian oscillator. In transgenicplants that constitutively express high levels of CCA1,other circadian outputs, such as leaf movement, losetheir rhythmicity and flowering is delayed. The circa-dian rhythms of other clock-regulated genes, includingCAB, CAT3 (a gene encoding the enzyme catalase), andan RNA-binding protein (CCR2) are also eliminated intransgenic plants that over express CCA1.

PhytochromeCryptochrome

Resetting clock

CentralOscillator

CCA1

OtherRhythms

Flowering

CCA1

CO

FT

TOC1

TOC1

ELF3

FIGURE 24.18 A simplified molecular model of the Ara-bidopsis circadian clock. The input signal, light, is per-ceived by phytochrome and cryptochrome. The centraloscillator is a negative feedback system. TOC1 is a posi-tive regulator of the CCA1 gene while CCA1 is a negativeregulator of the TOC1 gene. As TOC1 levels increase, anincrease in the levels of CCA1 follows. CCA1 in turnshuts down the production of TOC1. The result is thatTOC1 and CCA1 levels alternately rise and fall. ELF3,itself a product of the oscillator, blocks the input signalin the evening in order to ensure that the clock is resetby the morning light-on signal. One output of the clockis the transcription of CONSTANS (CO), which initiatesthe flowering process.

Another clock-associated mutant is elf3 (early flow-ering 3). The elf3 mutation not only advances floweringunder short days, but also renders both leaf movementand CAB expression arrhythmic when entrained plantsare shifted to continuous light. However, CAB2::lucactivity still oscillates when entrained plants are shiftedto continuous dark. ELF3 protein levels are regulatedby the clock and peak in the evening. ELF3 functions torepress (or, gate) the light input pathway in the eveningby binding to PHYB and inhibiting its activity. Theresult is that ELF3 makes the clock insensitive to lightin the evening and ensures that clock is reset by themorning light-on signal.

According to the current model, CCA1 and TOC1are part of a negative feedback loop that makes upthe central oscillator (Figure 24.18). TOC1 acts as apositive regulator of the CCA1 gene, thus stimulatingits transcription and the accumulation of CCA1 protein.CCA1 protein, however, acts as a negative regulatorof the TOC1 gene, so as CCA1 accumulates, one ofits actions is to repress TOC1 transcription. In thismanner, TOC1 and CCA1 levels alternately increaseand decrease, or oscillate, over a 24-hour period. Theoscillations are kept in phase by the dawn light-on signalreceived by phytochrome and cryptochrome and gatedby ELF3, itself a product of the oscillator.

The circadian oscillator generates multiple out-put rhythms, possibly through the regulatory action ofTOC1, CCA1, or related proteins on different genes.One of these certainly is the rhythmic output of CON-STANS (CO) in the leaf, which initiates the floralsignaling cascade as described earlier. Other multipleoutputs with different free-running periods could beexplained in several ways. The clock is more complexthan the simplified model shown in Figure 24.17. Asthe information accumulates, it is becoming increas-ingly clear that many additional genes and proteins areinvolved, forming multiple feedback loops. These addi-tional layers of complexity could easily lead to multiplerhythms with different periodicities. Alternatively, therecould be multiple oscillators within each cell, each linkedto different genes, or there could be different oscillatorswithin different cells in the same tissue.

24.3 PHOTOPERIODISMIN NATURE

Photoperiodism almost certainly reflects the need forplants to synchronize their life cycles to the time of year.Outside of the tropics, daylength is the most reliablepredictor of seasonal change (Figure 24.19). Not sur-prisingly, photoperiodism is more important to plantsin the subtropical and temperate latitudes where sea-sonal variations in daylength are more pronounced. Buteven many tropical plants respond to the small changesin daylength that occur within 5 or 10 degrees of the

Summary 431

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J F M A M J J A S O N DMonth of year

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f da

ylig

ht

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50º

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FIGURE 24.19 Daylength as function of latitude andmonth of year. Daylength is plotted as the time betweensunrise and sunset on the 20th of each month. (Datafrom The American Ephemeris and Nautical Almanac, U.S.Naval Observatory, 1969.)

equator. Does this mean that the photoperiod responseties a species to one particular latitude? Probably not,since the critical photoperiod only sets the upper (forSD plant) or lower (for LD plant) limits for daylength.Beyond that, flowering and other responses to pho-toperiod can usually occur within fairly broad limits.Moreover, there is evidence that populations of plantsare able to genetically adapt to latitude, thus givingrise to physiological ecotypes. In a variety of species,including Betula (birch), Chenopodium (lambs quarters),Oxyria digyna (mountain sorrel), and Xanthium, there areknown ecotypes or photoperiodic races characterized bydifferent critical daylengths. As a rule, the length ofthe critical day is longer as the individuals are collectedat more northerly latitudes. In most cases, the criticaldaylength seems to key flowering to a consistent timeinterval before the arrival of damaging autumn frosts atthat latitude.

Photoperiodism also helps to ensure that plantsflower in their temporal niche, reducing competitionwith others as well as ensuring that reproductive devel-opment is completed before the onslaught of unfavor-able winter conditions. In many species, germination,for a variety of reasons, may not be uniform. If flower-ing relied solely on plant size, nonuniform germinationwould be expected to spread flowering out in time aswell. To the extent that cross-pollination is required oradvantageous, flowering synchronized by photoperiodwould serve to ensure the maximum pollinating popu-lation or to coordinate flowering with the appearance ofa particular pollinating insect.

Photoperiod and its effects on geographical distri-bution of plants can have a direct impact on humans,as illustrated by the case of common ragweed (Ambrosiaartemisifolia). Ragweed is an annual SD plant, with acritical daylength of about 14.5 hours. The further northone goes, the longer the summer daylength. At the lat-itude of Winnepeg, Canada, (50◦N), for example, thedaylength exceeds 16 hours through most of June andJuly and doesn’t drop below 14.5 hours until mid-August(Figure 24.19). Ragweed induced to flower at that timeof the year would have insufficient time to flower andproduce mature seed before the arrival of killing frosts inearly fall. Since common ragweed can reproduce only byseed, it is abundant only in the more southerly regionsof Ontario, Quebec, and the maritime provinces. It israrely found, and then only on scattered patches of agri-cultural land, throughout most of western and centralCanada. Hay-fever sufferers in these regions are at leastspared the inconvenience of highly allergenic ragweedpollen, which plagues their neighbors to the south. Thisis not to suggest that photoperiod is the only factor thatlimits the distribution of plants such as ragweed, but itis clearly a significant part of the equation.

SUMMARY

Photoperiodism is a response to the duration and tim-ing of light and dark periods. There are three basicphotoperiodic response types: short-day (SD) plants,long-day (LD) plants, and day-neutral (DN) plants.Other response types are variations on the three basictypes and may be modified by environmental condi-tions such as temperature. A photoperiod requirementmay be qualitative, in which case the requirement isabsolute, or quantitative, in which case the favorablephotoperiod merely hastens the response. The distinc-tion between LD plants and SD plants is based on theirresponse to daylengths greater than or shorter thanthe critical daylength. The absolute critical daylengthvaries from one species to another and the criticaldaylength for a LD plant may be shorter than thecritical daylength for a SD plant.

432 Chapter 24 / Measuring Time: Controlling Development by Photoperiod and Endogenous Clocks

Plants actually measure the length of a dark inter-val between the light-off and light-on signals. Actionspectra of the light-break, which interrupts an other-wise inductive dark period, indicate that phytochromeis involved in the light signals. Photoperiodic lightsignals are perceived in the leaf but the response ulti-mately occurs elsewhere in the plant. This separation ofperception and response suggests the logical necessityfor a transmissible stimulus. In the case of flowering,the stimulus was proposed to be a hormone called flori-gen. It now appears that transmissible signal may bea small protein, FT, which is synthesized in the leafphloem parenchyma and carried through the sievetubes to the shoot apical meristem where it turns onthe flowering genes.

Many aspects of plant development, including pho-toperiodism and nyctinasty, are controlled by an inter-nal circadian clock. The circadian clock is difficult tostudy by traditional methods because, other than shift-ing the phase (equivalent to setting the clock), theclock is not readily manipulated by external influence.Three criteria that distinguish between simple periodicphenomena and clock-driven rhythms are (1) a clock-driven rhythm persists under constant conditions, (2) aclock-driven rhythm is reset or phased by environmen-tal signals such as light and temperature, and (3) clock-driven rhythms exhibit temperature compensation.

Time measurement in photoperiodism involves aninteraction between phytochrome and the endogenousbiological clock. The nature of the clock and the phy-tochrome/clock interactions is beginning to yield togenetic studies. The use of a CAB2::luc reporter systemhas enabled identification of several clock-associatedgenes in plants. The central oscillator in plants is anegative feedback system similar to the oscillator pre-viously found in insects, animals, and cyanobacteria.

Because changing daylength is the most reliablepredictor of seasonable change, photoperiodism almostcertainly reflects the need for plants to synchronizetheir life cycle to the time of year. Photoperiodismhelps ensure that plants flower in their temporal niche,reducing competition with others, or that reproductionis complete before the onslaught of unfavorable winterconditions.

CHAPTER REVIEW

1. You have discovered a new plant whose photope-riod characteristics are not yet described. Howwould you go about determining whether thisplant were a SD plant, a LD plant, or a day-neutralplant?

2. Describe the three hypotheses that have beeninvoked to explain the photoperiodic floralstimulus. What is the basis for each hypothesis?

3. Many metabolic processes appear to varythroughout a normal day. How would youdetermine whether these processes wereregulated by an endogenous, circadian clock?

4. What is the physiological significance ofphysiological ecotypes, or photoperiodicraces within a species that are charac-terized by different critical daylengths?

5. How does the response of Lemna to skele-tal photoperiods lend support to Bunning’snotion of a photophile and scotophile phase?

6. Distinguish between diurnal, circadian, andfree-running rhythms.

7. Endogenous circadian rhythms have beendescribed extensively, but their underlyingmechanism is only now yielding to genetic infor-mation. Why is it so difficult to experimentallyunlock the secrets of the circadian clock?

8. What has recent genetic evidence added toour understanding of the circadian clock?

FURTHER READING

Ayre, B. G., R. Turgeon. 2004. Graft transmission of a floralstimulant derived from CONSTANS. Plant Physiology135:2271–2278.

Gardener, M. J. et al. 2006. How plants tell time. BiochemicalJournal 397:15–24.

Hayama, R., G. Coupland. 2004. The molecular basis ofdiversity in the photoperiodic flowering responses ofArabidopsis and rice. Plant Physiology 135:677–684.

Hayama, R., G. Coupland. 2003. Shedding light on the cir-cadian clock and the photoperiodic control of flowering.Current Opinion in Plant Biology 6:13–19.

Imaizumi, T., S. A. Kay. 2006. Photoperiod control of flow-ering: Not only by coincidence. Trends in Plant Science11:550–558.

Hillman, W. S. 1962. The Physiology of Flowering. New York:Prentice-Hall.

Kondo, T., M. Ishiura. 1999. The circadian clocks ofplants and cyanobacteria. Trends in Plant Science 4:171–176.

Putterill, J., R. Laurie, R. Macknight. 2004. It’s time toflower: Genetic control of flowering time. Bioessays26:363–373.

Salisbury, F. B. 1963. The Flowering Process. New York: Perg-amon Press.

Sweeny, B. 1987. Rhythmic Phenomena in Plants. 2nd ed.New York: Academic Press.

Thomas, B., D. Vince-Prue. 1997. Photoperiodism in Plants.2nd ed. San Diego: Academic Press.

Wigge, P. A. et al. 2005. Integration of spatial and temporalinformation during floral induction in Arabidopsis. Science309:1056–1059.

Stamens

Carpels

Petals

Sepals

25Flowering and Fruit Development

Flowering and fruit development have long held theinterest of developmental biologists and physiologistsbecause they represent a dramatic change in the patternof shoot development and have significant economicimplications. The switch of the shoot apical meri-stem from vegetative to floral organs and the subse-quent development of fruit is a critical step in thedevelopmental history of a plant and must be regulatedprecisely in order to ensure reproductive success. Inthe previous chapter it was shown that synchronizationof flowering time with an environmental cue such asphotoperiod is not a simple event. It is possible onlythrough the interactions of many genetic and biochem-ical pathways: pathways involved in signal perceptionand transduction; pathways involved in the regulationof the circadian clock; and pathways involved in thedevelopment of the floral primordia.

Over the years, flowering in a number of species,including maize (Zea mays), petunia (Petunia sps.), snap-dragon (Antirrhinum sps.), tobacco (Nicotiana tobacum),and annual ryegrass (Lolium temulentum), has been thesubject of molecular and genetic studies. More recently,focus has shifted to the model plant Arabidopsis, wherea number of genes that influence flowering have beenidentified and a model proposed to account for thegenetic specification of floral organ initiation. In manycases, especially winter cereals and biennials, a periodof low temperature can significantly alter the flowering

response. The low-temperature treatment, called ver-nalization, influences flowering time under long days.Finally, flowering is followed by the development of aspecialized organ, the fruit, which ensure the properenvironment for seed maturation and dispersal of themature seed.

In this chapter we will examine

• the molecular genetic control of flower initiationand development in the shoot apical meristem,

• the phenomenon of vernalization and its relation-ship to other flowering time pathways, and

• the basic principles of fruit set and fruit develop-ment.

25.1 FLOWER INITIATION ANDDEVELOPMENT INVOLVESTHE SEQUENTIAL ACTIONOF THREE SETS OF GENES

As noted in the previous chapter, flowering in manyplants is influenced by environmental factors such asphotoperiod and temperature and involves the synthesisof a mobile floral stimulus in the leaves. Other plants donot require external inputs and the signal is generatedin the leaves when the plant simply reaches a minimum

433

434 Chapter 25 / Flowering and Fruit Development

stage of development. In any case, flower developmentis initiated when that signal arrives at the shoot apicalmeristem. During the vegetative state, the shoot apicalmeristem is programmed to produce leaf primordial.When the floral signal arrives from the leaf, the meri-stem acquires floral identity and secondary inflorescencemeristems, or floral primordia, arise in the axils of theuppermost leaf primordia.

The use of genetic mutants that alter flowering-timeand the initiation of floral primordia has been a pow-erful tool for identifying many of the genes involvedand dissecting the various pathways that lead to flower-ing (Table 25.1). As a general rule, mutants that causeearly flowering indicate a wildtype gene that normallyrepresses flowering while mutants that cause late flow-ering point to a wildtype gene that normally promotesflowering.

Extensive research with Arabidopsis has identifiedthree sequential stages to the flowering process, eachwith its own set of genes. The first set of genes com-prises the flowering-time genes. Flowering-time genesdetermine when the plant initiates flowering, either inresponse to the appropriate environmental signal orby monitoring the developmental state of the plant.Most mutations of the flowering-time genes cause theplants to flower later than normal, although a few willcause flowering to advance. One role of flowering-timegenes is to activate the expression of floral-identitygenes. Floral-identity genes commit undifferentiated

primordia to the production of floral rather than veg-etative structures. Mutations in floral-identity genescause primordia that would normally develop as flowersto produce structures with vegetative characteristics.The floral-identity genes in turn activate a set oforgan-identity genes that serve to control the sub-sequent development of floral organs such as sepals,petals, stamens, and carpels. Expression of floral- andorgan-identity genes, however, is not strictly linear.Some identity genes overlap in both the time of theirexpression and their function. As we shall see later, muta-tions in the organ-identity genes may cause abnormaldevelopment of any or all of the floral organs.

25.1.1 FLOWERING-TIME GENESINFLUENCE THE DURATIONOF VEGETATIVE GROWTH

Flowering-time genes provide the connection betweenflorigen, or the floral induction signal, and the transi-tion to the production of floral organs. Flowering-timemutants may therefore interfere with the production ofthe signal in the leaf (including the timing mechanism, orcircadian clock), translocation of the signal to the apex,or its activity in the apex. Most flowering-time mutantsidentified thus far cause plants to flower later than nor-mal, indicating that the mutants interfere with pathwaysthat normally promote flowering. Note that floweringtime, as it is used here, refers to a developmental time

TABLE 25.1 Some principal genes involved in flowering.

Gene Name Gene product Pathway/function

AG AGAMOUS (Unknown) Organ identityAP1 APETALA 1 (Unknown) Floral/organ identityAP2 APETALA 2 (Unknown) Organ identityAP3 APETALA 3 (Unknown) Organ IdentityCCA1 CIRCADIAN CLOCK ASSOCIATED 1 Transcription factor Circadian clockCO CONSTANS Transcription factor Long-day pathwayELF3 EARLY FLOWERING 3 Novel protein Circadian clockELF4 EARLY FLOWERING 4 Novel protein Circadian clockFCA FCA RNA-binding protein Autonomous pathwayFLC FLOWERING LOCUS C Transcription factor Floral repressorFT FLOWERING LOCUS T Lipid-binding protein Floral promoterGA1 GIBBERELLIC ACID 1 (Unknown) Gibberellic acid pathwayLD LUMIDEPENDENS Nuclear protein Autonomous pathwayLFY LEAFY Transcription factor Floral identity geneTOC1 TIMING OF CAB 1 Transcription factor Circadian clockVRN1 VERNALIZATION 1 DNA-binding protein Vernalization pathwayVRN2 VERNALIZATION 2 Repressor protein Vernalization pathwayVRN3 VERNALIZATION 3 Equivalent to FT Vernalization/LD pathway

25.1 Flower Initiation and Development Involves the Sequential Action of Three Sets of Genes 435

rather than chronological time (e.g., days to flower-ing). For example, Arabidopsis is a facultative long-dayplant with a critical photoperiod of 8 to 10 hours.The vegetative Arabidopsis plant grows as a rosette andflowering is preceded by stem elongation. Thus, underlong days Arabidopsis flowers with 4 to 7 leaves in therosette (about 3 weeks) but under short days floweringis delayed until 20 leaves have formed (7 to 10 weeks).Note that flowering is not related to elapsed time, butto the number of rosette leaves produced before theflowering stem appears. Late-flowering mutants simplyextend vegetative growth and increase the number ofleaves in the rosette before the stem elongates and theflowers develop.

It is interesting to note that, although a large num-ber of late-flowering mutants have been described, nosingle Arabidopsis mutant that remains vegetative indef-initely has yet been identified. This fits with the generalassumption that there are multiple pathways controllingflowering time with a certain amount of built-in redun-dancy. Redundancy provides that inactivation of genes inone pathway is at least partially compensated for by othergenes or complementary pathways. In Arabidopsis, atleast five separate, but interacting, pathways for control-ling flowering time have been identified (Figure 25.1).

Several flowering-time mutants, including fca, ld,and fve, flower later than wildtype plants under bothLD and SD conditions but remain sensitive to ver-nalization. Because flowering in the mutants is equallyaffected under both LD and SD conditions, the cor-responding wildtype genes are thought to be activein an autonomous pathway that monitors develop-mental stage and initiates flowering in response to

Long-daypathway

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FIGURE 25.1 Five separate genetic pathways controlflowering time in Arabidopsis. All pathways appear toconverge on LEAFY (LFY ), a floral-identity gene in theshoot apical meristem that mediates the switch from avegetative meristem to a floral meristem.

internal developmental signals. Such a signal is com-monly reflected in a minimum leaf number that mustbe achieved before flowering can proceed. Flower-ing of a second group of mutants, including constans(CO) and gigantea (GI), is delayed under LD condi-tions, but not under SD conditions. These mutants alsoshow no response to low-temperature treatments, orvernalization, which is normally linked to a LD flow-ering response and not a SD response. As described inChapter 24, the CO gene is believed to be a centralcomponent in the photoperiodic or long-day pathwayand is responsible for promoting the mobile floral stim-ulus FT. Both CO and GI have been cloned and studiedin some detail. It appears that GI operates before COin the same pathway and that floral promotion underlong days depends on the amount of CO protein and,subsequently, FT protein that is produced. The actionof FT at the shoot apical meristem is at least in partmediated by a transcription factor FD. At this point itappears that the combination of FT and FD proteinsis responsible for initiating flowering under long daysby activating floral identity genes in the shoot apicalmeristem.

A single mutant identified as ga1, which flowerslate under LD conditions and does not flower at allunder SD conditions, is thought to represent a sepa-rate pathway mediated by gibberellin, the gibberellicacid or GA pathway. Finally, there is a small group ofgenes that appear to act primarily as floral repressors.This conclusion is based on the observation that theirloss-of-function mutants [e.g., elf3 (early flowering 3) andphytochrome B (phyB)] cause early flowering. Some, suchas the phyB mutants, retain a response to photoperiodwhile others, such as elf3, do not. The loss of photope-riod response in elf3 appears due to disruption of thecircadian rhythm component of photoperiodic timing.Other early-flowering mutants, such as emf1 and emf2(embryonic flower 1-2), flower almost immediately fol-lowing germination, without forming any rosette leaves.Instead, the plants form reproductive structures on thesurface of their cotyledons. Because wildtype plantseventually flower, it appears that the repressor activityof EMF genes must be intended to prevent precociousflowering. The activity of repressor genes must declineduring development or at some point be turned off,eventually allowing one or more of the promotory path-ways to take precedence. In addition to the above fourgenetic pathways, a possible fifth pathway that mediateslow-temperature effects on flowering has been suggestedby recent studies.

One of the challenges that remain is to sort out towhat extent known flowering-time genes are involved inthe production, transmission, and perception of thefloral induction signal. In this regard, an interest-ing late-flowering mutation has recently been isolatedfrom maize (Zea mays). Flowering in maize follows

436 Chapter 25 / Flowering and Fruit Development

an autonomous pathway and wildtype plants normallyflower after producing a fixed number of leaves. Maizeplants carrying the mutant id1 (indeterminate 1), how-ever, continue to produce only leaves long after thewildtype plant has produced ears and tassels. When theyeventually do flower, the floral structures are aberrant.However, unlike many of the Arabidopsis flowering-timegenes, which are expressed in the shoot apical meristemand activate floral-identity genes, id1 is expressed only inthe young leaves. This pattern suggests that the functionof ID1 may be more closely related to the synthesis ofthe floral stimulus (or repression of floral inhibitors) inthe leaves.

25.1.2 FLORAL-IDENTITY GENESAND ORGAN-IDENTITYGENES OVERLAP IN TIMEAND FUNCTION

While the principal effect of flowering-time mutantsis on the duration of vegetative development, muta-tions in the floral-identity genes disrupt the transitionof the undifferentiated primordia to floral meristems.At least four floral-identity genes have been isolatedfrom Arabidopsis: LEAFY (LFY ), APETALA1 (AP1),APETELA 2 (AP2), and CAULIFLOWER (CAL). Floral-identity genes are expressed in the apical meristem priorto the formation of floral organs but their expressionis up-regulated rapidly following floral induction bylong days or application of gibberellin. However, theirindividual roles have been difficult to study becauseof extensive redundancy (i.e., a loss of function dueto one mutation is readily compensated by one of theother wildtype genes). The LEAFY gene appears toplay a key role in floral meristem identity. This canbe demonstrated by placing the gene under the controlof a strong gene promoter (designated 35S) from thecauliflower mosaic virus. Transgenic plants that con-tain the 35S::LEAFY combination (called a construct)bypass the requirement for a floral induction signaland express the gene constitutively. In such plants, ashortened primary shoot terminates early in clusters offlowers and all secondary shoots produce flowers fromthe rosette. By contrast, the lfy mutant produces moreinflorescence branches than a wildtype plant but the‘‘flowers’’ consist of green, leaf-like organs. Moreover,constitutive expression of the flowering-time gene COleads to a rapid activation of LEAFY in wildtype plants.LEAFY appears to have a central role in the floweringprocess. It is probably the principal target of the mobileflowering-time mobile stimulus FT when it arrives inthe meristem. LEAFY , in turn, activates organ identitygenes such as APETALA1 (AP1).

The Arabidopsis flower is rather typical amongadvanced flowering plants, consisting of four distinctwhorls of floral organs (Figure 25.2).The outermost

Stamens

Carpels

Petals

Sepals

FIGURE 25.2 (A) The Arabidopsis flower consists of fourdistinct whorls. The outermost whorl (whorl 1) consistsof four sepals, which are green and leaf-like. The nextwhorl (whorl 2) consists of four yellow petals. The thirdwhorl contains six stamens, and the innermost whorl(whorl 4) contains two fused carpels at the base of thepistil.

whorl (whorl 1) consists of four sepals, which are greenand leaf-like. The next whorl (whorl 2) consists of fouryellow petals. The third whorl (whorl 3) contains sixstamens, or male reproductive organs, and the inner-most whorl (whorl 4) contains two fused carpels at thebase of the female reproductive structure, the pistil.Mutations in combination with studies of temporal andspatial expression patterns have identified five genesthat are involved in the determination of organ iden-tity: APETALA1 (AP1), APETALA2 (AP2), APETALA3(AP3), PISTILATA (PI), and AGAMOUS (AG). Notethat AP1 and AP2 have both been previously identi-fied as floral identity genes as well. Mutations in theorgan-identity genes generally result in the modifica-tion, displacement, or total absence of floral organs. Inaddition, mutations in any one of these genes generallyinfluence the development of two adjacent floral organs.

The influence of organ-identity genes on the devel-opment of the Arabidopsis flower can best be understoodby viewing the floral meristem as three overlappingdevelopmental fields or fields of gene activity; desig-nated A, B, and C. Field A includes the sepals and petals(whorls 1 and 2), field B includes the petals and stamens(whorls 2 and 3), and field C includes the stamens andcentral carpels (whorls 3 and 4). This view is referredto as the ABC model for floral organ specification, inwhich a particular gene or pair of genes is associatedwith each developmental field, but controls the identityof two adjacent whorls of organs (Figure 25.3). Accord-ing to this model, expression of AP1 alone specifiessepals; AP1 in combination with AP3 and PI specifiespetals; AP3 and PI in combination with AG specify sta-mens; and AG alone specifies the carpels. Expression ofAP2 is apparently required throughout the meristem, inpart to suppress, in combination with a sixth, unknowngene (X ), the expression of AG in those whorls that aredestined to become sepals and petals.

25.2 Temperature Can Alter the Flowering Response to Photoperiod 437

Stamens CarpelsPetalsSepals

Genes APETALA1 AGAMOUS

APETALA2

APETALA3/PISTILLATA

Developmentalfields

Whorls

A

1 2 3 4

B

C

FIGURE 25.3 The ABC model for floral organ specifi-cation in Arabidopsis. (B) The floral meristem is visu-alized as being controlled by three developmental fields,identified as A, B, and C. Field A is involved in the speci-fication of whorls 1 and 2. Field B is involved in the speci-fication of whorls 2 and 3. Field C is involved in thespecification of whorls 3 and 4. Field A alone specifiessepals, fields A and B together specify petals, fields Band C together specify stamens, and field C alone speci-fies carpels. Each field is associated with a specific geneor gene pair. APETALA1 is expressed only in field A,APETALA3 and PISTILLATA are expressed in field B,and AGAMOUS is expressed only in field C. AGAMOUSalso represses (T-bar) the expression of APETALA1 infield C. APETALA2 is expressed throughout the meri-stem and, in conjunction with an unknown gene (X),represses the expression of AGAMOUS in field A. (AfterJ. D. Bewley et al., 2000.)

The ABC model can be used to either predictor interpret what will happen to organ developmentin loss-of-function mutants for each of these genes(Figure 25.4). For example, in the ap3 (or pi) mutant,developmental field B would be inoperative and the

3. Stamens

4. Carpels AP3

AP3

AP1

Se Pe St Ca

Se

Loss of B function

Loss of C function

Pe CaCa

Se SePePe

AP1

AP1

AG

AG

Wildtype

GenotypeObservedPhenotype

Whorls Interpretation

ap3(apetala3)

ag(agamous)

2. Petals

1. Sepals

3. Carpeloid stamens

4. Carpels

2. Sepaloid petals

1. Sepals

3. Petaloid stamens

4. Sepaloid carpels

2. Petals

1. Sepals

FIGURE 25.4 The ABC model helps tointerpret floral-identity in loss-of-functionmutants. In wildtype plants, all three devel-opmental fields are active and produce thenormal complement of sepals (Se), petals (Pe),stamens (St), and carpels (Ca). The apetala3mutant represents a loss of B function. Thisleaves AP1 to be expressed alone in whorl2, replacing the normal petals with sepaloidstructures. Also, AG is expressed alone inwhorl 3, producing carpeloid structures inplace of normal stamens. The agamous mutantrepresents a loss of C function, leaving AP1to be expressed in all four whorls. The resultis petaloid structures in whorl 3 and sepaloidstructures in whorl 4.

meristem would be unable to form either petals orstamens. Not only is this the case, but the whorls thatwould normally be filled with petals and stamens arefilled instead with sepal-like and carpel-like structures.Similarly, in the ag mutant, stamens and carpels are notformed and all four whorls are filled with sepals or petals.No doubt this model will continue to be refined as newgenes are discovered and the various genes and theirproducts are subjected to further molecular analysis.

Although it is becoming more evident which genesare active in controlling which aspects of floral initiationand development, their biochemical and physiologi-cal function is not yet clear. However, most of thegene products that have been analyzed thus far sharecertain characteristics, called motifs, which are typi-cal of transcription factors. Transcription factors areDNA-binding proteins that enable RNA polymerasesto recognize promoters and begin transcription of thegene in eukaryotes. It thus appears that the principalfunction of most flowering-time, floral-identity, andorgan-identity genes is to regulate other genes that maythen direct synthesis of the components that actuallymake up the individual organs.

25.2 TEMPERATURE CAN ALTERTHE FLOWERING RESPONSETO PHOTOPERIOD

There are many examples of interactions between tem-perature and photoperiod, particularly with respect toflowering behavior (see Salisbury, 1963, for an exten-sive listing). In most cases the interaction results inrelatively subtle changes in the length of the critical

438 Chapter 25 / Flowering and Fruit Development

photoperiod or a tendency toward daylength neutral-ity or an inability to flower altogether at high or lowtemperature extremes. There are other plants, however,for which flowering is either quantitatively or qualita-tively dependent on exposure to low temperature. Thisphenomenon is known as vernalization. Vernalizationis a means of preventing precocious reproductive devel-opment late in the growing season, ensuring insteadthat seed production does not begin until the beginningof the next growing season so that the seed will havesufficient time to reach maturity.

Vernalization refers specifically to the promotionof flowering by a period of low-temperature and shouldnot be confused with other miscellaneous effects oflow-temperature on plant development. The term itselfis a translation of the Russian yarovizatsya; both wordscombining the root for spring (Russian, yarov; Latin,ver) with a suffix meaning ‘‘to make’’ or ‘‘become.’’Coined by the Russian T. D. Lysenko in the 1920s, ver-nalization reflects the ability of a cold treatment to makea winter cereal mimic the behavior of a spring cerealwith respect to its flowering behavior. The responsehad actually been observed many years earlier by agri-culturalists, but didn’t receive critical attention of thescientific community until J. G. Gassner showed in 1918that the cold requirement of winter cereals could besatisfied during seed germination. For his part, Lysenkoreceived considerable notoriety for his conviction thatthe effect was an inheritable conversion of the winterstrain to a spring strain. His position—a form of thethoroughly discredited Lamarkian doctrine of inheri-tance of acquired characteristics—was adopted as Sovietdogma in biology and remained so until the 1950s.The adoption of Lysenko’s views as official dogmahad a significant impact on Soviet biology and placedagriculture in the USSR at a severe disadvantage fordecades.1

25.2.1 VERNALIZATION OCCURS MOSTCOMMONLY IN WINTERANNUALS AND BIENNIALS

Typical winter annuals are the so-called ‘‘winter’’ cere-als (wheat, barley, rye). ‘‘Spring’’ cereals are normallydaylength insensitive. They are planted in the springand come to flower and produce grain before the end ofthe growing season. Winter strains, however, if plantedin the spring would normally fail to flower or pro-duce mature grain within the span of a normal growing

1The story of vernalization is a classic example of what canhappen when science becomes enmeshed in politicalideology. For the interested student, this unfortunate episodein the history of science has been artfully documented byD. Joravsky in his book The Lysenko Affair (Chicago:University of Chicago Press, 1970).

season. Winter cereals are instead planted in the fall.They germinate and over-winter as small seedlings,resume growth in the spring, and are harvested usuallyabout midsummer. The over-wintering cold treatment,or vernalization, renders the plants sensitive to longdays.

One of the most thorough studies of vernaliza-tion and photoperiodism was carried out on the Petkuscultivar of rye (Secale cereale) by F. G. Gregory andO. N. Purvis, beginning in the 1930s. There are twostrains of Petkus rye: a spring strain and a winter strain.The spring strain of Petkus rye is a facultative long-dayplant. Under short days, floral initiation does not occuruntil after about 22 leaves have been produced, typi-cally requiring a season of about 4.5 months. Underthe appropriate long-day regime, however, flowering inthe spring strain is initiated after as few as seven leaveshave been produced, requiring only about two months.When sown in the spring, the winter strain is insensi-tive to photoperiod. The winter strain flowers equallyslowly—requiring four to five months—regardless ofdaylength.

If seeds of the winter strain are sown in the fall,however, the germinated seedlings are subjected toan over wintering low-temperature treatment. Whenthey resume growth in the spring, winter strain plantsrespond as long-day plants in the same way as the springstrain. The effect of the over wintering cold treatmentcan also be achieved by vernalizing the seed, that is, byholding the germinated seed near 1◦C for several weeks.Note that the low-temperature treatment, at least in thecase of winter annuals, does not alone promote earlyflower initiation. Rather, the effect of vernalization is torender the seedling sensitive to photoperiod.

Another example of vernalization is seen in bien-nial plants. Biennials are monocarpic plants that normallyflower (and die) in the second season, again follow-ing an over-wintering cold treatment. Typical bien-nials include many varieties of sugar- and table-beet(Beta vulgaris), cabbages and related plants (Brassicasp.), carrots (Daucus carota) and other members ofthe family Umbellifereae, foxglove (Digitalis purpurea),and some strains of black henbane (Hyoscyamus niger).Biennials share with the winter annuals the propertythat subjecting the growing plant to a cold treat-ment stimulates a subsequent photoperiodic floweringresponse.

Biennials typically grow as a rosette, charac-terized by shortened internodes, in the first season(Figure 25.5). Over winter, the leaves die back butthe crown, including the apical meristem, remainsprotected. New growth the following spring is char-acterized by extensive stem elongation, called bolting,followed by flowering. The cold requirement inbiennials is qualitative (i.e., absolute). In the absence ofa cold treatment many biennials can be maintained in

25.2 Temperature Can Alter the Flowering Response to Photoperiod 439

FIGURE 25.5 Vernalization and stem elongation in cab-bage. Left: Cabbage plants were vernalized for six weeksat 5◦C before being returned to the greenhouse. Center:Plants were sprayed weekly with a solution containing 5× 10−4 M gibberellic acid. Right: Control plants grownat normal greenhouse temperature remain in a rosettehabit. Except for the vernalization treatment, all plantswere maintained in the greenhouse under a long-day(16-hour) photoperiod.

the nonflowering rosette habit indefinitely. As a rule,vernalized plants, whether winter annuals or biennials,tend to respond as long-day flowering plants, althoughsome biennials are daylength-indifferent followingvernalization. One exception to the rule is the perennialChrysanthemum morifolium, a SDP. Some varieties ofChrysanthemum require vernalization before respondingas a quantitative SDP. As a perennial, Chrysanthemumnormally requires vernalization on an annual basis.Many other plants such as pea (Pisum sativum) andspinach (Spinacea oleracea) can be induced to flowerearlier with a cold treatment but it is not an absoluterequirement.

25.2.2 THE EFFECTIVE TEMPERATUREFOR VERNALIZATION ISVARIABLE

The range of temperatures effective in vernalizationvaries widely depending on the species and duration ofexposure. In Petkus rye, the effective range is −5◦Cto +15◦C, with a broad optimum between +1◦C and+7◦C. Within these limits, vernalization is proportionalto the duration of treatment. Flowering advances sharplyafter as little as one to two weeks’ treatment at 1◦C to2◦C and is maximally effective after about seven weeksat that temperature (Figure 25.6). Within the effectiverange, the temperature optimum is generally higherfor shorter treatment periods. Presumably, a longerexposure to lower temperatures within the effective

Duration of cold treatment (days)

2010 30 504040

60

40

20

100

80

Day

s to

flo

wer

ing

120

FIGURE 25.6 Vernalization in Petkus rye (Secale cereale).Seeds were germinated in moist sand at 1◦C for thetime indicated. Cold treatments were scheduled so thatall seeds were returned to the greenhouse at the sametime. The number of days to flowering progressivelydecreased with increasing length of the cold treat-ment. (From Purvis, O. N., F. G. Gregory. 1937. Annalsof Botany, N.S. 1:569–591. Copyright, The Annals ofBotany Company.)

range is required because the metabolic reactions leadingto the vernalized state progress more slowly.

Like flowering, the vernalized state is more or lesspermanent in most species, giving rise to the con-cept of an induced state. For example, vernalizedHyoscyamus, a LDP, can be held under short daysfor up to 10 months before losing the capacity torespond to long-day treatment. On the other hand,all cold-requiring plants that have been studied arecapable of being devernalized—vernalization can bereversed if followed immediately by a high-temperaturetreatment. Flowering in vernalized winter wheat, forexample, can be fully nullified if the seedlings are heldnear 30◦C for three to five days. For most plants, then,there is a ‘‘neutral’’ temperature where neither vernal-ization nor devernalization occurs. For Petkus rye theneutral temperature is about 15◦C. Vernalized seeds ofPetkus rye can also be devernalized by drying them forseveral weeks or by maintaining the seeds under anaer-obic conditions for a period of time following the coldtreatment.

440 Chapter 25 / Flowering and Fruit Development

25.2.3 THE VERNALIZATIONTREATMENT IS PERCEIVEDBY THE SHOOT APEX

A vernalization treatment is effective only on activelygrowing plants. Cold treatment of dry seeds will notsuffice. Thus winter cereals may be vernalized as soonas the embryo has imbibed water and the germinationprocess has been initiated. Other plants, in particularthe biennials, must reach a certain minimum size beforethey can be vernalized. Hyoscyamus (black henbane), forexample, is not sensitive before 10 days of age and doesnot reach maximum sensitivity until 30 days of age. Ineither case, the cold treatment appears to be effectiveonly in the meristematic zones of the shoot apex. Thiscan be shown by localized cooling treatments or ver-nalization of moistened embryos. Early studies showedthat even the cultured apex of isolated rye embryoswas susceptible to vernalization. Thus the induced stateestablished in a relatively few meristematic cells can bemaintained throughout the development of the plant.Most biennials, however, cannot be induced as seeds.In these plants it is the over-wintering stem apex thatperceives the stimulus, although there are some reportssuggesting that leaves and even isolated roots may besusceptible in some cases.

25.2.4 THE VERNALIZED STATEIS TRANSMISSIBLE

Experiments with isolated embryos have shown that ver-nalization treatments are effective only when the embryois supplied with carbohydrate and oxygen is present,indicating that it is an energy-dependent metabolic pro-cess. Still, the nature of the induced state has eludedresearchers for many years. To the extent that themeristem itself is the site of perception, any necessityfor a transmissible hormone appears to be ruled out.A cold-induced, permanent change in the physiologicalor genetic state of the meristematic cells (referred toas ‘‘mitotic memory’’) would be self-propagating, thatis, it could be passed on to daughter cells through celldivision. There is some support for this argument. Inplants such as Petkus rye and Chrysanthemum, only tissueproduced in a direct cell line from the induced meristemis vernalized. If the cold treatment is localized to a singleapex, it will flower, but all the buds that did not receivethe cold treatment will remain vegetative.

In other experiments, especially with Hyoscyamus,transmission of the vernalized state across a graft unionhas been demonstrated. A list of successful experimentshas been tabulated by A. Lang in his 1965 review (seeFurther Reading). These experiments are comparableto the transmission of florigen’’ across a graft union(Chapter 24) and result in flowering in nonvernal-ized receptor plants. If a vernalized Hyoscyamus plant is

grafted to a nonvernalized plant, both will flower underlong days. Transmission requires a successful (i.e., liv-ing) graft union and appears to be coordinated with theflow of photoassimilate between the donor and receptor.

Experiments such as those described above ledG. Melchers to propose the existence of a transmissi-ble vernalization stimulus called vernalin. Like florigen,vernalin has resisted all attempts at isolation and remainsa hypothetical substance. Unfortunately, the vernalinstory is to some extent clouded by interpretation. Thegrafting experiments all require vernalization followedby long days. They do not clearly distinguish betweenthe transmission of ‘‘vernalin’’ and the possibility thatthe nonvernalized partner is responding instead to thefloral stimulus itself (e.g., FT), which would be trans-mitted from the vernalized donor under long days.

Adding to the complexity of vernalization is theapparent involvement of gibberellins in the response tolow temperature (see Figure 25.5). This was dramaticallydemonstrated by A. Lang in 1957 when he showed thatrepeated application of 10 μg of GA3 to the apex wouldstimulate flowering in nonvernalized biennial strains ofHyoscyamus and several other biennials maintained undershort days. No such promotion occurred in Xanthiumand other short-day plants treated with gibberellin undernoninductive long days. Subsequently it has been shownthat gibberellin levels tend to increase in response tolow-temperature treatments in several cold-requiringspecies.

The role of gibberellins is not clear, although innoninduced plants very high concentrations of the gib-berellin precursor ent-kaurenoic acid accumulate in theshoot apex. This suggests that the cold treatment isrequired to complete the synthesis of gibberellins inthese plants.

25.2.5 GIBBERELLIN ANDVERNALIZATION OPERATETHROUGH INDEPENDENTGENETIC PATHWAYS

Results such as those described in the previous sectionhave raised the question: Are vernalin and gibberellinequivalent? The answer, on both physiological andgenetic grounds, is no. It is true that gibberellin appearsto substitute for the cold requirement of some vernal-izable plants and for the long-day requirement in somelong-day plants, or, in the case of vernalization, both.But virtually every situation in which gibberellin has suc-cessfully substituted for low temperature or long daysin promoting flowering involves bolting, or the rapidelongation of stems from the rosette vegetative state.Far less success has been achieved with gibberellinsin caulescent long-day plants—those whose stems arealready elongated in the vegetative state. Moreover,the developmental pattern in responsive plants differs

25.2 Temperature Can Alter the Flowering Response to Photoperiod 441

significantly depending on whether stem elongation isstimulated by low temperature or gibberellin treatment.Following low-temperature treatment, flower buds areevident at the time stem elongation begins. Followinggibberellin treatment, on the other hand, the stem firstelongates to produce a vegetative shoot. Flower buds donot appear until later. These results suggest independentpathways for vernalization and gibberellins.

Recent genetic studies of flowering have confirmedthat gibberellin and vernalization operate via separategenetic pathways (see Figure 25.1). When a triplemutant was constructed containing mutant alleles(co-2, fca-1, ga1-3) that impair each of the long-day,autonomous, and GA-dependent pathways, the mutantplants failed to flower under either long days or shortdays in controlled environment rooms. After 90 to 100rosette leaves had been produced without flowering, theplants were then transferred to a long-day greenhouse.After six months, the majority of the mutant plantshad died without ever flowering. However, if the triplemutant seedlings were first vernalized at 5◦C for 7weeks, all the plants flowered after approximately 50leaves had been produced. The most straightforwardinterpretation of these results would be that (1) thetriple mutant has an absolute requirement for vernaliza-tion and (2) vernalization promotes flowering throughyet another genetic pathway that is separate from theGA-dependent, long-day, and autonomous pathways.

Both the autonomous pathway and vernalizationreduce the expression of the gene FLOWERING LOCUSC (FLC). The product of this gene is a transcriptionfactor that represses flowering. However, when thewildtype FLC gene is absent, control by the autonomouspathway is also eliminated but the effect of vernalizationis not. Thus it appears that the autonomous pathwayacts solely through controlling FLC expression, butvernalization is able to promote flowering through twopathways: either through suppressing FLC expression orthrough some yet-to-be-discovered FLC-independentmechanism.

25.2.6 THREEE GENES DETERMINETHE VERNALIZATIONREQUIREMENT IN CEREALS

This brings us to the question of whether vernalin, likeflorigen, is a hormone. Or a better question to ask mightbe: what is the molecular basis for vernalization? It hasactually been known for more than 30 years, primarilythrough plant breeding experiments, that three genes(VERNALIZATION 1, 2, and 3, or VRN1, VRN2, andVRN3) have a major role in determining the vernaliza-tion requirements in cereal grains. These genes havenow been isolated and characterized.

The VRN1 gene encodes a transcription factor andis induced by a vernalization treatment. Furthermore,

there is a quantitative relationship between the amountof VRN1 expressed in a vernalized plant and the amountby which flowering time is reduced under long days.VRN2 represses flowering under long days by block-ing the expression of the floral stimulus FT . On theother hand, VRN2 is itself repressed by VRN1. Vari-eties of winter cereals that lack a functional copy ofVRN2 respond normally to long days without requiringa prior cold treatment. VRN3 is the cereal equivalent(called an ortholog) of FLOWERING LOCUS T (FT) inArabidopsis.

A model to illustrate how these three genes inter-act to control flowering in winter wheat and barleyis presented in Figure 25.7. Prior to receiving a coldtreatment, the winter cereals are unable to respond tolong days because FT (or VRN3) expression is repressedby the presence of VRN2. When the seeds are sownin late summer or early autumn, the shoot apex devel-ops vegetatively until winter, when VRN1 expressionis promoted. In the spring, growth is renewed; thelow-temperature-induced expression of VRN1 remainshigh; and VRN1 suppresses any further expression of

Arabidopsis

Lowtemperature

Flowering

Long days (PPDI)

FT(VRN3)

Lowtemperature

CO

VRN1VRN2

FLC VRN2 VRN1

VRN1

Winter cereals

FIGURE 25.7 A model comparing the regulation of flow-ering by vernalization in winter cereals and Arabidop-sis. Long days (mediated by PHOTOPERIOD 1 (PPD1)in cereals) are sensed by the CONSTANS gene (CO)which activates FLOWERING LOCUS T (FT) expres-sion. VRN2 prevents floral induction before winter byrepressing FT expression. A low-temperature vernaliza-tion treatment induces expression of VRN1, whichrepresses VRN2 and allows expression of FT under longdays. How the low temperature induces VRN1 expres-sion isn’t known. In cereals, VRN1 also acts as a floralmeristem identity gene. In Arabidopsis, FT expressionis repressed by FLOWERING LOCUS C (FLC). Flow-ering proceeds under long days following vernalizationbecause the low temperature represses FLC expression.(Based on Trevaskis et al. 2007. Trends in Plant Science.)

442 Chapter 25 / Flowering and Fruit Development

VRN2. FT is then allowed to be expressed and the plantnow responds to long days by initiating floral primordia.VRN2 is referred to as a floral integrator gene because,as shown in the model, its central role is to integrate thelow temperature and photoperiod responses.

Flowering in the dicot Arabidopsis is also acceleratedby vernalization and the molecular mechanism is verysimilar to that in the cereals. The principal differenceis that in Arabidopsis the gene FLOWERING LOCUSC (FLC), not VRN2, is most directly responsible forsuppressing FT expression and maintaining the vegeta-tive state of the apex. The repression of FLC, in turn,appears to be regulated by low temperature through acomplex involving VRN1 and VRN2 (Figure 25.7).

25.3 FRUIT SET ANDDEVELOPMENT ISREGULATED BY HORMONES

The fruit is the final stage in the growth of the repro-ductive organ. Botanically a fruit is a mature or ripenedovary wall and its contents, although in some plantsother floral parts may become involved. There is a widediversity of fruits, depending on how the ovary develops.In its simplest form, such as peas or beans, the fruit con-sists of the seed or seeds enclosed within an enlarged butdry ovary (the pod). Such fruits are classified as dehis-cent fruits—dehiscent because at maturity the ovary wallbreaks open to free the seeds. The fruit of Arabidopsis is adry dehiscent fruit. Maize (Zea mays) is a non-dehiscentdry fruit consisting of a single seed with its seed coatsfused with the dry ovary wall (a structure called the peri-carp). Tomato is an example of a fleshy fruit (actually aberry) with an enlarged, fleshy inner fruit wall. In somespecies, a structure other than the ovary wall developsas the fruit. These are called pseudocarpic fruits. Astrawberry is one example. A strawberry ‘‘fruit’’ actu-ally consists of a number of individual one-seeded fruits(called achenes) borne on the surface of an enlarged,fleshy receptacle. In many cases, it is clear that the fruitundergoes considerable cell division and cell enlarge-ment as well as significant qualitative changes. Thesechanges are due largely to changes in hormone content.

Fruit development, maturation, and ripeninghave been widely studied because of their biologicalsignificance—fruits protect the developing seed andserve as a vehicle for dispersal of the mature seed—aswell as their practical importance as a significantcomponent in human nutrition. The development,maturation, and ripening of fleshy fruits have receivedthe bulk of the attention over the years because ofproblems associated with transportation, storage, andother aspects of post-harvest physiology.

25.3.1 THE DEVELOPMENT OF FLESHYFRUITS CAN BE DIVIDED INTOFIVE PHASES

Tomato (Lycopersicon esculentum) has become a popularmodel in which to study fleshy fruit development, in partbecause there are numerous mutants available and theplant is easily transformed. With tomato as a model, thelife history of a fruit can be divided into five more-or-lessdistinct phases. Phase I involves the development of theovary in preparation for fertilization and seed develop-ment and ends with the decision to either abort furtherdevelopment or to proceed with further cell divisionand cell enlargement in the ovary walls. This decisionto proceed with ovary development is generally referredto as fruit set. In phase II, or the initial phase of fruitdevelopment, growth of the nascent fruit is due primar-ily to cell division in the ovary walls. The cells thusbecome small and dense, with very small vacuoles. Dur-ing phase III, cell division effectively ceases and furthergrowth of the fruit is mostly by cell enlargement. Oncethe fruit has reached its final size, it enters phase IV,or a period of ripening. In a fleshy fruit like tomato,ripening involves the development of color and flavorconstituents (e.g., carotenoids, sugars, and acids) and asoftening of the tissue that render the fruit attractive toanimals. Tissue softening is due primarily to increasedactivity of enzymes such as polygalacturonase (PG).PG degrades the pectic substances that are found in themiddle lamella and which are responsible for cell-to-celladhesion. Finally, in phase V, senescence sets in and thefruit begins the decay process.

As might be expected, all of the plant hormonesare active at various stages during fruit development(Figure 25.8). During seed development and first andsecond phases of fruit development, auxins, cytokinins,and gibberellins are all present and active. Cytokininlevel peaks during phase II, the period of most active celldivision. Auxin level peaks in early phase III, coincidingwith the initiation of cell enlargement, and then declinesas the fruit reaches mature size. A second surge in auxinlevel occurs in the early stages of ripening, along with theappearance of significant levels of ethylene. The role ofgibberellins is not well understood, but they are probablyinvolved with cytokinins in initiating cell division andwith auxin in maintaining cell enlargement. Tomato is aclimacteric fruit and the burst in respiration is related tothe appearance of ethylene and the qualitative changesin the fruit that represent ripening (see Chapter 21 fora discussion of ethylene).

25.3.2 FRUIT SET IS TRIGGEREDBY AUXIN

Normally, successful pollination and fertilization of theovule by sperm nuclei are required for fruit set to occur.

25.3 Fruit Set and Development is Regulated by Hormones 443

AUXIN

RR

PHASE I II III IV

Fruitset

GA

CK

GA

AUXIN

ETHYLENE

MG BR

FIGURE 25.8 Hormonal changes during thefirst four phases of fruit development in atomato (Lysopericon esculentum). The graphsfor auxin, gibberellin (GA), cytokinins (CK),and ethylene show the approximate pointin development when hormone concentra-tions peak. Mature green (MG). Breaker(BR) refers to the stage when the first signsof color appear. Breaker marks the begin-ning of the ripening phase. The red ripestage (RR) marks the end of ripening andthe beginning of senescence.

In the absence of fertilization, flowers fail to produceseeds and, in most cases, the floral parts senesce withoutforming fruits. The commitment to fruit set thereforeappears to rely on positive growth signals generatedprior to or immediately following fertilization. Thereare some cases, however, in which unfertilized flow-ers go on to produce normal, but seedless, fruits. Thephenomenon of fruit development in the absence ofpollination and fertilization is known as virgin fruit-ing or parthenocarpy (from parthenos, meaning virgin).Parthenocarpy may occur naturally due to lack of polli-nation or pollination that does not result in fertilization,or from pollination followed by embryo abortion. Com-mon examples of naturally parthenocarpic fruits arebananas, navel oranges, most varieties of figs, pineapple,and some seedless grapes. Studies involving either nat-ural or induced parthenocarpy have had a major role inhelping to understand the initial phases of normal fruitdevelopment, especially fruit set.

In the mid-1930s, it was found that pollen andextracts of pollen were both a rich source of auxin andcould stimulate fruit set in unpollenated plants of thefamily Solanaceae (e.g., tomato, peppers). Auxin inducesparthenocarpy in a small number of plants in otherfamilies as well, particularly in the Cucurbitae (cucum-ber, pumpkin), and Citrus. Parthenocarpy is a biologicalcuriosity—of what value is a seedless fruit—but it hassignificant economic implications as well. In California,for example, synthetic auxins are used to stimulate fruitset in early tomato crops, when cool night temperatureswould otherwise tend to reduce fruit set. Many otherfruits such as citrus and watermelons are more salablein the seedless form. Gibberellins, either alone or incombination, may also induce parthenocarpy in speciessuch as pear and citrus.

Other than pollen, at least one source of auxin forfruit initiation is the seed. This was demonstrated ina classic study of strawberry (Fragaria ananassa) con-ducted by J. P. Nitsch in the 1950s. Since the fruit ofa strawberry is the floral receptacle and the achenes

are borne on the surface, it is a simple task to removethe seeds without damaging the underlying receptacle.Nitsch found that removal of the seeds prevented fur-ther development of the fruit, but supplying the fruitwith auxin restored normal development. Furthermore,if some seeds were removed and others left in place,the fruit would develop only in those regions wherethe seeds were undisturbed (Figure 25.9). Other studiesconfirmed that strawberry achenes were a good sourceof auxin.

Molecular approaches have followed up on the clas-sical physiological studies and confirmed that auxin fromthe developing ovule has a central role in fruit set andearly fruit development. In general, these approacheshave taken advantage of artificially induced partheno-carpic fruit development, either by (1) increasing thesynthesis of auxin in the ovule or ovary, or (2) by manip-ulating auxin regulatory proteins such as AUX/IAA orauxin response factors (ARF). (Refer to Chapter 18 forthe role of AUX/IAA and auxin response factors inauxin-regulated responses.)

FIGURE 25.9 Nitsch’s experiment showing the effectof seed (achenes) removal on the development of straw-berry fruit. All but two rows of seeds were removed fromthe receptacle during early development of the fruit.Subsequent fruit development was limited to the regionwhere the seeds were left in place.

444 Chapter 25 / Flowering and Fruit Development

One approach, for example, involves the Agrobac-terium tumifaciens tumor-inducing genes iaaM and iaaH.These are the genes that encode IAA biosynthesis andcause the overproduction of IAA when Agrobacteriuminvades host cells. IaaM and iaaH can be used toconstruct a transgene by linking them to a promoterthat restricts their expression specifically to the ovule.When this transgene construct is inserted into egg-plant, tobacco, tomato, strawberry, and raspberry plants,the constitutive production of IAA induces partheno-carpic fruit development. Another gene that inducesparthenocarpy is the rolB gene, isolated from anotherAgrobacterium species (Agrobacterium rhizogenes). TherolB gene, again linked to a promoter specific for theovary and young fruit, induced parthenocarpy whenit was inserted into tomato. The mechanism of rolBaction is not known, but it appears to increase the tissuesensitivity to auxin.

In Arabidopsis, loss-of-function mutants for the auxinresponse factor gene ARF8 induce parthenocarpic fruitdevelopment. ARF8 is a transcription factor that isexpressed primarily in the ovule and surrounding tis-sues. Further experiments indicated that ARF8 acts as anegative regulator for fruit set, i.e., it inhibits fruit set,probably in combination with one or more AUX/IAAproteins. Expression of the ARF8 gene is also switchedoff after fertilization.

Most of the data for the role of auxin in fruit set isconsistent with the following model. Prior to pollina-tion and fertilization, further development of the ovaryis blocked by the presence of AUX/IAA and/or ARFrepressor proteins such as ARF8. Pollination (and fertil-ization of the ovules) induces an increase in the level ofauxin, thus leading to the auxin-mediated degradationof the AUX/IAA proteins via the 26S-proteasome path-way. This frees up the auxin-response factors to regulateauxin-responsive genes that are necessary for fruit setand subsequent fruit development.

25.3.3 RIPENING IS TRIGGERED BYETHYLENE IN CLIMACTERICFRUITS

In many, but not all, fleshy fruits the metabolic andvisual changes that occur during the ripening process areaccompanied by a significant burst in respiratory activityor CO2 evolution, called the climacteric (Figure 25.10).Examples of climacteric fruits include tomato, cucurbits(cucumber and related fruits), banana, apple, peaches,and plums. Nonclimacteric fleshy fruits, which do notshow the CO2 burst, include strawberry, grape, cit-rus, and all nonfleshy, or dry fruits such as Arabidopsisor maize. The ripening process in climacteric fruitshas attracted a lot of research over the years becauseof its economic importance and because just prior tothe respiratory burst there is a significant increase in

MG

FruitSet

FirstColor

C2H4

CO2

III IVIIBR

FIGURE 25.10 The pattern of CO2 and ethylene (C2H4)production in tomato (Lycopersicon esculentum), a climac-teric fruit. CO2 evolution is high during the cell divisionphase and steadily declines during the cell expansion andmaturation phase. It then rises sharply at the breakerstage which identifies the beginning of the ripeningphase. The CO2 ‘‘burst’’ is immediately preceded by asignificant increase in the production of ethylene. (AfterMcGlasson, W. D. 1978. In: H.O. Hultin, M. Milner(eds.), Postharvest Biology and Biotechnology. Westport,CT: Food and Nutrition Press.)

the production of ethylene (Figure 25.10). Moreover,ethylene synthesis is also auto-catalytic. Once ethyleneproduction begins in one fruit, its production is stim-ulated in surrounding fruit—hence the old axiom thatone rotten apple spoils the barrel. The role of ethylenein fruit ripening has assumed significant commercialimportance. For example, tomatoes, bananas, and otherclimacteric fruits that have to be shipped any distance arecommonly picked at the mature green stage and thenripened at their destination by gassing with ethylene(Box 25.1).

The rate limiting steps in the biosynthesis of ethy-lene are catalyzed by the enzymes ACC synthase (ACS)and ACC oxidase (ACO) (see Chapter 21). In tomatothere are two ACS genes that are expressed in the fruitand appear to be responsible for triggering ripening.Both genes, LeACS1A and LeACS4, are under develop-mental control and are induced at the onset of ripening.2Furthermore, the induction of both genes is impairedby mutations at either the ripening inhibitor (rin) or thenonripening (nor) locus. In other words, fruits of tomatoplants carrying the rin and nor mutations do not produceethylene, do not exhibit a climacteric CO2 burst, anddo not ripen. The expression of the gene LeACS4 isalso controlled by ethylene itself and thus appears to beresponsible for regulating the autocatalytic productionof ethylene by a positive feedback system.

A large number of ethylene signaling componentshave been identified in both Arabidopsis and tomato,

2In gene designations such as LeACS, the Le identifies genesisolated specifically from tomato, Lycopersicon esculentum.

Summary 445

RR

BOX 25.1ETHYLENE:IT’S A GAS!

Ethylene has been unwittingly used by humans for cen-turies to stimulate fruit ripening. In ancient China,harvested fruit was commonly ripened in rooms whereincense was being burned and in California in the earlytwentieth century kerosene stoves were used to ripenlemons. These practices worked because ethylene is acommon combustion product. For the same reason,ethylene can still occasionally cause ethylene-mediatedgrowth problems for modern-day greenhouse opera-tions that heat with natural gas.

Once the role of ethylene in ripening becameknown, however, attention turned to methods for con-trolling fruit ripening by preventing or delaying theeffects of ethylene production in climacteric fruits.Apples, for example, are harvested in the fall but aregenerally available in the markets throughout the yearbecause of controlled atmospheric storage. Controlledatmospheric storage employs a combination of low tem-perature, low ambient oxygen, and high ambient CO2.Oxygen levels are generally reduced to 1 percent to3 percent from the normal 21 percent. CO2 levels maybe increased up to 8 percent from the normal 0.035 per-cent. These conditions lower both ethylene productionand the respiration rate of the fruit, and thus modu-late, if not prevent, the ethylene-stimulated respiratoryclimacteric.

including some ethylene receptors that are present onlyin the fruit and are strongly induced during the ripen-ing process. The challenge now is to understand howthese many components interact to form a coherentsignal transduction chain that regulates a multitude offruit-ripening genes.

SUMMARY

Flower initiation and development involves the actionof at least three sets of genes. These genes regulateflowering-time (including the circadian clock), floral-meristem identity, and floral-organ identity.Flowering-time mutants may flower either later than orearlier than the wildtype. Flowering time in Arabidopsisis under the control of at least four different geneticpathways. The long-day pathway genes constitutethe photoperiodic control system. The autonomouspathway monitors the state of vegetative developmentand initiates flowering in response to endogenoussignals. The GA pathway involves a single gene,mediated by gibberellin. The fourth pathway involvesa set of genes that normally repress flowering. Mutantsin the repressor pathway flower early.

Mutations in floral-identity genes disrupt thetransition to a floral meristem, while organ-identitygenes control the initiation of sepals, petals, stamens,and carpels. Some floral-identity genes and organ-identity genes overlap in time of expression and func-tion. The ABC model for floral organ specificationviews the meristem as three overlapping developmen-tal fields and identifies the genes and combinationof genes that specify specific floral organs. Most ofthe flowering-time, floral-identity, and organ-identity

genes encode products that have some characteristicsof transcription factors.

Vernalization is the promotion of flowering bya period of low temperature. In the case of winterannuals, such as cereals, vernalization changes the pho-toperiodic behavior from daylength indifference toa quantitative long-day response. Biennials typicallygrow as a rosette until vernalized. The flowering stemthen bolts (elongates) and responds as a long-day plant.A temperature of approximately 0◦C to 5◦C, applied tothe actively growing apex of the plant for several weeks,is required for vernalization to be effective. Tempera-ture in vernalizable plants is perceived in the stem apexand is transmissible. The use of flowering-time mutanthas shown that vernalization operates independently ofthe long-day, autonomous, and gibberellin-dependentgenetic pathways.

Fruits are classified as dry or fleshy, dependingon the extent of ovary development following fer-tilization. The development of fleshy fruits such astomato and apple can be divided into five phases.Phases II through IV represent the progression fromfruit set to the fully ripe fruit. Auxin is responsiblefor fruit set, while auxins, cytokinins, and gibberellinsare involved at various times until the fruit reachesmaturity. Fruit development is normally initiated bya release of auxin associated with pollination and/orfertilization of the ovule. Parthenocarpy is the devel-opment of seedless fruits in the absence of pollinationor fertilization. Parthenocarpy can be induced in somefruits by the application of auxin. Phase IV, or ripen-ing, is characterized by the development of color andflavor constituents and softening of the fruit due to abreakdown in the pectic substances that are involve incell-to-cell adhesion. Ripening in some fleshy fruits is

446 Chapter 25 / Flowering and Fruit Development

accompanied by a marked increase in respiratory rateand CO2 release (the respiratory climacteric) triggeredby ethylene. Ethylene release is catalyzed at least in partby ACC synthase and ACC oxidase, two rate limitingenzymes that are developmentally controlled. Ethy-lene initiates a signal transduction chain that turns onripening genes.

CHAPTER REVIEW

1. How are events in the leaf connected with the con-version of a vegetative meristem to a floral meris-tem?

2. Distinguish between flowering-time genes,floral-identity genes, and organ-identity genes.

3. What would be the expected observed phe-notype of an Arabidopsis flower that carries aloss-of-function mutation for the APETALA1gene?

4. Most plants have several pathways thatcontrol flowering-time. What might bethe advantage(s) of multiple pathways?

5. How does the concept of induction apply to ver-nalization?

6. Gibberellin often appears to substitutefor vernalization. What is the evidencethat vernalization and gibberellin operatethrough independent genetic pathways?

7. Compare the roles of FLOWERING LOCUS Cand VERNALIZATION 2 in controlling flowering.

8. What is ‘‘fruit set’’? What do you under-stand the meaning of ‘‘decision’’ to beas it is used with respect to fruit set?

9. What is the ‘‘climacteric’’?10. How does controlled atmospheric storage

enhance the storage life of climacteric fruit?

FURTHER READING

Adams-Phillips, L., B. Cornelius, J. Giovannoni. 2004. Signaltransduction systems regulating fruit ripening. Trends inPlant Science 9:331–338.

Bewley, J. D. et al. 2000. Reproductive development. In:Buchanan, B., W. Gruissem, R. Jones (eds.), Biochemistryand Molecular Biology of Plants. Rockville, MD: AmericanSociety of Plant Physiologists. (An excellent review ofreproductive development.)

Coen, E. S., E. M. Meryerowitz. 1991. The war of the whorls:Genetic interactions controlling flower development.Nature 353:31–37.

Dennis, E. S., W. J. Peacock. 2007. Epigenetic regulation offlowering. Current Opinion in Plant Biology 10:520–527.

Gillaspy, G., H. Ben-David, W. Gruissem. 1993. Fruits: Adevelopmental perspective. The Plant Cell 5:1439–1451.

Giovannoni, J. 2004. Genetic regulation of fruit developmentand ripening. Plant Cell 16:S170–S180.

Lang, A. 1965. Physiology of flower initiation. In:W. Ruhland (ed.), Handbuch der Pflanzenphysiolgie(Encyclopedia of Plant Physiology). Berlin: Springer-Verlag,XV (1) 1380–1536.

Pandolfini, T., B. Molesini, A. Spena. 2007. Molecular dis-section of the role of auxin in fruit iniitation. Trendsin Plant Science 12:327–329.

Salisbury, F. B. 1963. The Flowering Process. Oxford: Perga-mon Press.

Trevaskis, B., M. N. Hemming, E. S. Dennis, W. J. Pea-cock. 2007. The molecular basis of vernalization-inducedflowering in cereals. Trends in Plant Science 12:352–357.

Trevaskis, B., M. N. Hemming, W. J. Peacock, E. S. Dennis.2006. HvVRN2 responds to daylength, whereas HvVRN1is regulated by vernalization and developmental status.Plant Physiolgy 140:1397–1405.

Yanofsky, M. F. 1995. Floral meristems to floral organs:Genes controlling early events in Arabidopsis develop-ment. Annual Review of Plant Physiology and Plan Molecu-lar Biology 46:167–188. (Currently titled Annual Reviewof Plant Biology.)

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26Temperature: Plant Development

and Distribution

Plants are chemical machines and one universal char-acteristic of chemical machines is their sensitivity totemperature. Temperature, along with light and water,is one of the most critical factors in the physical envi-ronment of plants. This is especially so because plants,unlike homeothermic animals, are not able to maintaintheir tissues at a constant temperature. Environmentaltemperature therefore exerts a profound influence oncellular metabolism and, as a result, plant growth andtheir geographic distribution.

All of the chemical machinery of nature—everyindividual enzymatic reaction, every metabolic function,every physiological process—has temperature limitsabove and below which it cannot function and anoptimum temperature range where it proceeds at amaximum rate. Temperature also affects the integrityof cell structure (especially the structure and proper-ties of membranes), limits the distribution of species inspace and time, and influences the direction of specificdevelopmental events.

Temperature as an environmental stress and flow-ering regulator has been discussed in earlier chapters. Inthis chapter we will introduce some of the other ways inwhich temperature is known to influence plant growth,development, and distribution. Specific topics include:

• the role of temperature in perennial plants, in par-ticular its role in bud and seed dormancy, and

• some examples of how temperature influences thegeographic distribution of plants.

26.1 TEMPERATURE IN THEPLANT ENVIRONMENT

Of all the planets, the thermal environment on earth isparticularly fit to give rise to and sustain life. This isbecause life functions in an aqueous medium and therange of temperatures encountered over most of theearth’s surface generally ensures that sufficient wateris maintained in the liquid state. The temperatureat which biological processes can occur is generallylimited by the freezing point of water on the lowside and the irreversible denaturation of proteins onthe high side. Between these two extremes, a plot ofgrowth versus temperature for individual organismsassumes the shape of an asymmetric bell curve, similarto that for individual enzyme reactions or multipleenzyme-catalyzed metabolic sequences (Figure 26.1).In fact, the temperature curve for growth of an

447

448 Chapter 26 / Temperature: Plant Development and Distribution

Photosynthesis

Grossphotosynthesis

Netphotosynthesis

Respiration

Growth

Temperature

Tmax

Topt

Tmin

B.

A.

FIGURE 26.1 Temperature and plant growth. (A) Aschematic illustration of the three cardinal temperaturesfor plant growth. Typically, the pattern of the growthcurve reflects the pattern of temperature effects on netphotosynthesis. (B) Net photosynthesis is the differencebetween gross carbon uptake by photosynthesis and car-bon evolution by respiration.

organism effectively represents a composite of thetemperature curves for photosynthesis, respiration, andother critical metabolic processes. Growth curves thusexhibit, just as do individual metabolic and enzymereactions, the three cardinal temperatures (minimum,optimum, and maximum). Just as the actual values ofcardinal temperatures vary between different metabolicprocesses, the actual values of cardinal temperaturesfor growth curves will vary from species to species.Assuming other factors are not limiting, these cardinaltemperatures generally define the temperature rangeover which growth is possible. It is close to the extremesof this range that plants experience temeprature stressas described in earlier chapters.

Green plants probably first evolved in the tropicalregions, not so much because of warmer temperatures(although that may have been a factor), but because thetemperatures there were relatively stable. With time,plants gradually migrated into the temperate and polar

regions as they adapted to wider variations in temper-ature on a daily and seasonal basis. Green plants arenow found in regions as extreme as the Antarctic conti-nent and the northern tundra, where temperatures overmuch of the year are near or below freezing, and in thewarmest places on earth such as Death Valley (Califor-nia), where summer temperatures commonly approachor even exceed 50◦C.

Plants and related organisms may be broadly clas-sified according to their ability to withstand tempera-ture. Those that grow optimally at lower temperatures(between 0◦C and 10◦C) are called psychrophiles. Thepsychrophiles include primarily algae, fungi, and bac-teria. Higher plants generally fall into the category ofmesophiles, whose optimum temperatures lie roughlybetween 10◦C and 30◦C. Thermophiles will growunhindered at temperatures between 30◦C and 65◦C,although there are reports of cyanobacteria growing attemperatures as high as 85◦C. These temperature rangesapply to hydrated, actively growing organisms. Dehy-drated organisms and organs, such as resurrection plants(Selaginella lepidophylla) and dry seeds with moisture con-tents as low as 5 percent, are able to withstand a muchbroader range of temperatures for extended periods oftime.

Plants in nature are subjected to a complex mosaicof fluctuating air and soil temperature regimes such thatit is very difficult to study the effects of temperature in anatural setting. Air temperature, for example, fluctuateswidely, and often rapidly, depending on the time ofday, cloud cover, season, and other factors. Soil is amajor heat sink as it absorbs and stores solar energyduring the day. At night, some of this heat is radiatedback into the atmosphere, which both cools the soiland warms the surface. Soil temperature also varies withthe soil structure, organic content, and other physicalcharacteristics as well as slope and aspect (the directionit faces with respect to the sun).

Both air and soil temperatures have an impact onplant growth. Air temperature influences leaf tempera-ture and therefore the rates of photosynthesis, respira-tion, and other metabolic reactions. On the other hand,soil temperature influences germination, root develop-ment, and nutrient uptake. For example, maize seeds willnot germinate below about 10◦C and the time requiredfor germination of winter wheat increases linearly with adecrease in soil temperature below 25◦C. Several inves-tigators have shown that the uptake of nutrients suchas calcium, boron, nitrogen, and phosphorous increaseswith increasing temperature. In cold soils, soybean rootsspread out closer to the soil surface, while in warmersoils the roots grow more deeply. The stem apex (or,crown) of many grasses is situated at or near the soilsurface, which means that leaf development in grassesis probably influenced more by soil temperature thanby air temperature. The study of soil temperature and

26.2 Bud Dormancy 449

its influence on growth and development is of morethan passing interest because soil temperature can be asignificant factor in determining agricultural yield.

While the temperature in tropical climes is relativelystable, plants growing in temperate regions and closerto the poles are subject to more or less predictable vari-ations in temperature on a daily and seasonal basis. It isperhaps not too surprising that plants have evolved waysto incorporate this information in their developmen-tal and survival strategies. Plants use this informationto ensure dormancy of buds, tubers, and seeds, and tomodify their flowering behavior, all of which appear tobe keys to survival over periods unfavorable to normalgrowth and development.

26.2 BUD DORMANCY

One aspect of development that is strongly influencedby temperature is dormancy. Dormancy is a term that isapplied to tissues such as buds, seeds, tubers, and cormsthat fail to grow even though they are provided withadequate moisture and oxygen at an appropriate temper-ature. Dormancy is a difficult process to study becauseit is a progressive process, often occurring in degrees.Buds that are just entering dormancy, for example,may be stimulated to renew growth rather easily. Onthe other hand, buds that have developed full dormancymay require prolonged or severe treatment to break dor-mancy and renew growth. Dormancy in some organsmay be enforced by other organs in the plant or byexternal factors. As a result, dormancy terminology hasbeen quite inconsistent and can be confusing. Termssuch as quiescence, rest, true dormancy, and imposed orenforced dormancy have been used by different authori-ties to describe various states or conditions. However, itis most convenient to group dormancy mechanisms intoone of three types: paradormancy, in which the inhibi-tion of growth arises from another part of the plant (e.g.,apical dominance); ecodormancy, in which growth isimposed by limitations in the environment (e.g., lack ofwater); and endodormancy, in which the dormancy isan inherent property of the dormant structure itself (seeLang, 1987).

Bud dormancy is an example of endodormancy.A bud is a shortened, very compact terminal or axillaryshoot axis in which the internodes have failed to elongateand leaves or floral parts have not enlarged. The wholeis enclosed in a set of modified leaves, called bud scales(Figure 26.2). Bud scales serve a protective function,both insulating the bud and preventing desiccation.Many popular fall flowering bulbs are also large buds(Box 26.1).

The growth of deciduous perennial plants typical oftemperate regions, where warm summers alternate withcold winters, is cyclical; such plants normally undergo

FIGURE 26.2 An axillary bud of sweet bay (Laurus noblis).Note the prominent bud scales that enclose and protectthe bud.

a cessation of active growth on an annual basis. Bothvegetative and floral buds formed in the latter part of thesummer or early fall will survive over the winter in a dor-mant condition and do not normally renew their growthuntil more favorable temperature conditions return inthe spring. However, it is not the low temperature itselfthat inhibits bud growth. Dormancy mechanisms arewell under way before the cold temperatures arrive and,similar to vernalization, a period of low temperature isrequired before bud regrowth can occur in the spring.Just as vernalization prevents precocious flowering inthe fall, bud dormancy prevents precocious bud growth,ensuring that the meristems enclosed within the bud areable to survive the adverse conditions of winter.

In order to ensure survival of the plant, dormancymechanisms must be in place before the arrival of unfa-vorable conditions. This means that the plant must beable to anticipate climatic change. Mechanisms must alsobe in place to ensure that the buds do not break dor-mancy until such time that environmental conditions areappropriate to sustain normal growth and development.Premature breaking of buds during an unseasonablywarm period in the winter, for example, could have seri-ous consequences for the survival of the plant. In short,dormancy is a precisely regulated requirement for theperennial habit, cued by factors in the environment andmaintained, and ultimately broken, by specific metabolicchanges in the organism.

Dormancy studies have focused on three principalquestions. (1) What are the environmental signals thatstimulate the onset of dormancy and how are they

450 Chapter 26 / Temperature: Plant Development and Distribution

BOX 26.1BULBS AND CORMS

Terminal and axillary buds borne on the aerialportion of plants are not the only buds that aresubject to dormancy mechanisms. A large numberof plants have fleshy underground storage organscapable of carrying the plants through seasonal coldperiods. Popularly referred to as bulbs, they are moreaccurately defined as bulbs, tubers, or corms. The truebulb (e.g., lily, hyacinth, tulip, daffodil, and onion)

B.A.

FIGURE 26.3 Examples of fall bulbs and corms. (A) Tulip; a true bulb. The bud isenclosed by layers of clasping, fleshy storage leaves, similar to an onion. (B) Crocus;a corm. The buds are borne on a flattened stem. Both go dormant in late summer toearly fall. Dormancy is broken by planting the bulbs or corms in the fall and subject-ing them to cool winter ground temperatures.

is a large bud, consisting of a small, conical stemwith numerous leaves surrounding one or more cen-tral meristems (Figure 26.3). The leaves are modifiedfor food storage. In an onion, for example, these leavesare the portion of the bulb that we eat. Corms, rep-resented by plants such as crocus and gladiolus, aresolid shortened stems with numerous buds systemati-cally arranged under a protective covering of paper-thinleaves.

The most popular spring-flowering bulbs andcorms, including ornamental onion (Allium sps.) aregenerally referred to as ‘‘fall bulbs’’ because they areplanted in the fall after entering a period of dormancyand require an extended cold period before the budsbreak dormancy and flower in the spring.

perceived? (2) What metabolic changes are responsiblefor the reduced activity? (3) What signals the startup ofrenewed growth at the appropriate time?

26.2.1 BUD DORMANCY IS INDUCEDBY PHOTOPERIOD

Plants anticipate seasonal change by monitoring pho-toperiod. The onset of dormancy in buds is a typicalshort-day response, coincident with leaf fall, decreased

cambial activity, and increased capacity to withstandlow temperature, or cold hardiness In temperate woodyspecies, the short days and decreasing temperatures oflate summer and autumn induce the leaf primordia toform bud scales instead of leaves. The formation of scalesis followed by the induction of cold hardiness and thecessation of cell division in the meristem. Once growthhas ceased and meristem has entered dormancy, themeristem becomes insensitive to any growth-promotingsignals.

26.3 Seed Dormancy 451

Like flowering, the short-day photoperiod signalthat initiates the onset of dormancy is perceived inthe leaves. It should not be surprising, then, that thesame players that detect photoperiod for the controlof flowering are also involved in controlling dormancy.For example, when the genes CONSTANS (CO) andFLOWERING LOCUS T (FT) were over-expressed intransgenic poplar trees, bud growth did not stop fol-lowing exposure to short days. On the other hand,down-regulation of FT triggers the onset of dormancy.Thus it appears that the combination of CO and FTrepresents a universal photoperiodic signal module. It isnot known, however, which gene(s) or gene product(s)FT interacts with to trigger bud dormancy.

Relatively little is also known about the physio-logical state of dormant buds except that during theirformation the bud primordia accumulate storage mate-rials such as starches, fats, and proteins. Dormant budsare further characterized by low respiratory activity, asignificant loss of water, and the inability to grow evenif temperature, oxygen, and water supply are adequate.There have been reports that the endogenous gibberellinlevels decline at the onset of dormancy and that somebuds may be released from dormancy if treated withgibberellin or cytokinins, but, otherwise, little is knownabout the hormonal status of dormant buds.

26.2.2 A PERIOD OF LOWTEMPERATURE IS REQUIREDTO BREAK BUD DORMANCY

Although induction of bud dormancy is coincident withdecreasing temperature and short days, the principalrole of temperature appears to be in breaking dor-mancy. Most dormant buds have a chilling requirementthat must be met before the cells are capable of renewedcell division and enlargement. Studies on the chillingrequirement for breaking dormancy have concentratedon commercial fruit species and deciduous ornamen-tals. This is because in the northern hemisphere, thechilling requirement largely determines the southerlylimits of cultivation for these plants. The process isespecially critical in fruit trees since the flower bud thatbears fruit are initiated in the previous summer. Thebud then over-winters and, having satisfied its chillingrequirement, floral development continues the follow-ing spring. Apple trees, for example, will not bear fruitwithout a cold winter.

Temperatures near or just above freezing appear tobe most effective at breaking dormancy. The amountof chilling required varies with species, cultivar, andeven location of the buds on the trees. Species such asapple (Malus pumila), pear (Pyrus communis), and cherry(Prunus sps.) require approximately 7 to 9 weeks ofexposure to temperatures below 7◦C in order to over-come dormancy. Others may require up to 22 weeks

(American plum, Prunus americana) or as few as fourto six weeks (apricot, Prunus armeniaca). Persimmons(Diospyros kaki ) require only four days of low tem-perature and so can be grown successfully much furthersouth than other fruit trees. The temperature in temper-ate regions often varies widely throughout the winter,but this generally poses no problem for dormant tissues.In most cases, buds and other dormant tissues are ableto sum the periods of cold and will not renew growthuntil the appropriate amount of cold treatment has beenaccumulated.

There is wide variation in the chilling requirementof different species and ecotypes (or, genetic races) ofmaple. More than 12 weeks of chilling are required tobreak dormancy in sugar maple (Acer saccharum) col-lected in southern Canada, while those collected fromthe warmer regions near the southern limits of its dis-tribution required only a few weeks of low temperature.Similar results were obtained for seedlings of red maple(Acer rubrum).

Very little is known about the molecular mech-anisms involved in breaking bud dormancy. What isknown comes largely from the study of various speciesof poplar (Populus sps.), birch (Betula papyrifera), andapple where recent efforts have been directed towardcharacterizing the expression of dormancy-responsivegenes. In one species of poplar (P. tricocarpa), forexample, a gene homologous with FLOWRING LOCUSC (FLC-like gene or PtFLC) has been implicated inbud dormancy. FLC is the floral suppressor that isdown-regulated in Arabidopsis during a vernalizationtreatment (Chapter 25). PtFLC is expressed in the shootapices of poplars grown under long days but, like FLC inArabidopsis, PtFLC declines in dormant buds during thelow temperature period. By analogy with vernalization,the down-regulation of PtFLC could be a key compo-nent in the cold-mediate release of bud dormancy inpoplar. In another study, it was found that expressionof the gene KNAP2 increased during the onset of dor-mancy in apple buds, but was down-regulated during thebreaking of dormancy. KNAP2 is a KNOTTED-likehomeobox protein; a group of master control proteinsthat have a fundamental role in development of plantsand animals alike (see Chapter 20).

26.3 SEED DORMANCY

26.3.1 NUMEROUS FACTORSINFLUENCE SEED DORMANCY

Seeds are in many respects similar to buds—they consistof a small embryonic axis (along with some storage tis-sues) enclosed by a series of membranes, collectivelycalled the seed coat. The seed coat serves a pro-tective function much as bud scales do. Its presence

452 Chapter 26 / Temperature: Plant Development and Distribution

often suppresses germination by restricting the uptakeof water and exchange of oxygen, it mechanically lim-its the expansion of the embryo and, in some cases,contains inhibitors that prevent growth of the embryo.These limitations can be removed and the germinationof many seeds accelerated by mechanically disrupting orremoving the seed coat, a process called scarification.In the laboratory, scarification may be accomplishedwith files or sandpaper. In nature, abrasion by sand,microbial action, or passage of the seed through animalgut will accomplish the same end. Seed coats can be verytough. Uniformity and rate of germination of morningglory (Pharbitis nil), cotton, and some tropical legumeseeds, for example, can be improved by soaking the dryseed in concentrated sulfuric acid for up to an hour.Scarification by passage through animal gut no doubtoccurs as a result of the acidic conditions in the gut.

As with buds, dormancy in seeds refers to the situa-tion wherein the embryo fails to grow because of physi-ological or environmental limitations. These limitationscommonly include the inability of water or oxygen topenetrate the seed coat. Seeds of some plants, par-ticularly in the family Leguminoseae, have specializedstructures that control seed moisture content. E. Hydedescribed a structure in seeds of lupine (Lupinus arboreus)that functions as a hygroscopically operated check-valveand that limits imbibition of water by the seed. Becausewater cannot pass through the unscarified seed coat,the only possible route of entry is through a smallpore, called the hilum (Figure 26.4). When the water

Counterpalisade

Parenchyma

Tracheid cluster

Palisadeepidermis

CuticleHilumfissure

FIGURE 26.4 A cross-section through a portion of theseed of Lupinus arboreus (tree lupine) showing the hilum,a hygroscopic valve that regulates water loss from theseed. The counter palisade is a group of thick-walledcells lying on the outer surface of the cuticle. Whenthe seed moisture content is higher than the moisturelevel in the ambient air, the hilum fissure is open andwater escapes from the tracheid cluster. When the waterambient moisture level is higher than inside the seed,the counter palisade cells swell and close off the hilumfissure in order to prevent the seed from rehydrating.(From Hyde, E. O. C. 1954. Annals of Botany 18:241.With permission of The Annals of Botany Company.)

content of the seed is higher than ambient, the hilumis open to permit the exit of water and allow the seedto dry. But when the moisture content outside the seedis higher than inside, cells surrounding the hilum swell,thus closing off the pore and preventing the uptake ofwater. In addition, as the seed dries out the permeabilityof the seed coat to water also decreases and the dormancyof the seed increases. Other seeds have pores that areblocked with a plug, called the strophiolar plug, whichmust be mechanically dislodged before water and oxygencan enter.

There is a considerable body of evidence to suggestthat seed coats also interfere with gas exchange, oxygenuptake in particular. As noted above, removal of the seedcoat often leads to a significant increase in respiratoryconsumption of oxygen. Measurements of the oxygenpermeability of seed coats have been made and thereis general agreement that permeability is very low inthose seeds tested. However, it is not always clear thatlimited oxygen permeability is the primary cause ofdormancy. The complexity of the situation and problemsof interpretation are well illustrated by studies of thegenus Xanthium, or cocklebur.

A cocklebur contains two seeds: an upper, dormantseed and a lower, nondormant seed. Dormancy of theupper seed can be overcome either by removing the seedcoat or by subjecting the intact seed to high oxygen ten-sion. The inference is that seed coat permeability in thedormant seed limits the supply of oxygen to the embryoand thus prevents germination. However, several otherobservations have cast doubt on this conclusion. Thereare, for example, no measurable differences betweenthe dormant and nondormant seed with respect to thepermeability of the seed coat to oxygen. Moreover, therate of oxygen diffusion through the seed coats is morethan sufficient to support measured rates of oxygen con-sumption by the embryos inside. Clearly, dormancy ofthe upper seed in Xanthium cannot be due to limitedpermeability of the seed coat to oxygen. Why then,do the upper, dormant seeds require a higher oxygenlevel to elicit germination? It appears that the seedcoat is a barrier, not to the uptake of oxygen but tothe removal of an inhibitor from the embryo. Aqueousextracts of Xanthium seeds have revealed the presence oftwo unidentified inhibitors, based on tests of the extractsin a wheat coleoptile elongation assay. The same twoinhibitors are found in diffusate collected from isolatedembryos placed on a moist medium, but not in diffusatefrom seeds surrounded by an intact seed coat. Thus ger-mination in the dormant seed appears to be prevented bythe presence of these inhibitors and the seed coat servesas a barrier that prevents those inhibitors from beingleached out. The oxygen requirement can be explainedby the observation that high oxygen tension reduces thequantity of an extractable inhibitor, presumably by someoxidative degradation.

26.3 Seed Dormancy 453

Even the role of inhibitors in seed dormancy isnot clear. Along with hormones such as auxins andgibberellins, a large number of inhibitors have beenidentified in seeds, fruits, and other dispersal units.These include hormones (ABA), unsaturated lactones(coumarin), phenolic compounds (ferulic acid), variousamino acids, and cyanogenic compounds (i.e., com-pounds that release cyanide) characteristic of apple andother seeds in the family Rosaceae (see Chapter 27).The simple presence of an inhibitor does not, how-ever, prove its role in dormancy. The inhibitors couldbe localized in tissues not directly involved in growthof the embryo or otherwise sequestered so as to pre-clude any role in preventing germination. Evidence insupport of a role for inhibitors is generally limited toleaching experiments such as that described above forXanthium. In some cases, dormancy can then be restoredby exposing the leached seed to the putative inhibitor.In order to clearly establish whether an inhibitor has anactive role in regulating germination, it is necessary toestablish whether inhibitor levels in the seed correlatewith the onset and termination of dormancy. In spiteof the voluminous literature relating inhibitors to dor-mancy, there is very little critical support for a directrole. For the present, evidence for the imposition andmaintenance of dormancy by inhibitors remains largelycircumstantial.

26.3.2 TEMPERATURE HAS ASIGNIFICANT IMPACTON SEED DORMANCY

Temperature has a significant impact on termination ofdormancy in many seeds. In fully imbibed seeds, bothalternating and low (chilling) temperatures are knownto terminate dormancy. Many seeds, even though fullyhydrated, will not germinate when maintained underconstant temperature. They require instead a diurnalcycle of fluctuating temperature. The required temper-ature differential between the high and low temperatureis often not great, ranging from a few degrees to perhaps5◦C or 10◦C, depending on the species. Germination ofbroad-leaved dock (Rumex obtusifolia) seeds, for example,exceeds 90 percent when the temperature differential isabout 10◦C and when the high temperature is given for16 hours each day.

The reaction to alternating temperature is complexand poorly understood. In Rumex, alternating treatmentsare effective only when the high temperature is greaterthan 15◦C. Also, when the high temperature is givenfor only 8 hours each day, a differential of only 5◦C isrequired to induce 90 percent germination. Although insome cases the effect of alternating temperature appearsto be localized in the embryo itself, there are manywell-documented cases where the effect of alternatingtemperature is mechanical. It is, in effect, a form of

scarification, releasing the seed from some kind of seedcoat–imposed dormancy.

It has long been known that freshly shed seedsof many herbaceous and woody species have dormantembryos that can be induced to growth only by aprolonged low-temperature treatment. These includemaples (Acer sps.), hazel (Corylus), and many generain the family Rosaceae (pear, Pyrus; apple, Malus;hawthorne, Crateagus). Normally, following therequired period of low temperature, the seeds will notgerminate until temperatures are more favorable forembryo emergence and seedling development. In mostcases this requirement ensures that the seed shed in latesummer or fall will not germinate until spring.

The exposure to low temperature that satisfiesthis germination requirement is known as eitherpre-chilling, or stratification. The latter term has itsorigin in the horticultural practice of layering seedsin moist sand or peat moss and exposing them to lowtemperature for several weeks or months to inducegermination. It is important that the pre-chillingrequirement for release of seed dormancy not beconfused with vernalization, which is a cold treatmentto an already germinated seedling, as discussed earlierin the previous chapter. As with breaking of buddormancy, temperatures near freezing but below 10◦Care most effective for terminating seed dormancy. Theoptimum for most species is near 5◦C. In a populationof seeds, the effectiveness is also a function of the lengthof the cold treatment (Figure 26.5).

It is presumed that seeds undergo some metabolicchanges during the period of low temperature, gener-ally referred to as after-ripening, but the exact natureof these changes is unclear. There is some evidencefor redistribution of carbon and nitrogen from theendosperm to the embryo, a decline in the inhibitorcontent, and a rise in gibberellin and cytokinin content.

Ger

min

atio

n, %

Length of cold treatment (days)

200 40 80600

75

50

25

100

FIGURE 26.5 Breaking dormancy in apple seeds with lowtemperature. Moist seeds were held at 4◦C for the timeindicated. (Redrawn from Luckwell, L. C. 1952. Journalof Horticultural Science 27:53.)

454 Chapter 26 / Temperature: Plant Development and Distribution

Gibberellin treatments will substitute at least partiallyfor the cold requirement in many seeds, just as they doin other cold-requiring systems.

26.4 THERMOPERIODISMIS A RESPONSE TOALTERNATINGTEMPERATURE

Growers have long recognized the beneficial effect onplant growth of lowering greenhouse temperatures dur-ing the night. This effect has been particularly welldocumented by the work of F. Went and his colleaguesin the 1940s. Went found that tomatoes (Lycopersicumesculentum) grown at constant temperatures of 26◦C and18◦C grew poorly and (at 26◦C) failed to produce fruit.Plants maintained under alternating conditions of 26◦Cduring the day and 18◦C at night grew vigorously andproduced a maximum number of fruit. In order to beeffective, the day–night differential had to be synchro-nized with the light–dark cycle. If the temperature cyclewas inverted, with the high temperature falling dur-ing the dark period, growth was even poorer than at aconstant 26◦C. To describe this phenomenon, F. Wentcoined the term thermoperiodism.

It is now recognized that many, but certainly notall, plants perform better under regimes with a similartemperature differential. For those that do, the effectis primarily on vegetative development, in contrast tophotoperiodism where the influence is primarily on flo-ral production. In some plants, such as potato (Solanum)and tobacco (Nicotiana), low night temperature leadsto a decline in shoot–root ratio, due to preferentialroot growth. Stems of Begonia also respond to ther-moperiodism. It has been reported that stem elongationwas inhibited by low daytime temperatures alternatingwith high nighttime temperatures (14◦C/24◦C) whencompared with constant daytime/nighttime tempera-tures (19◦C/19◦C). On the other hand, stem elongationwas promoted by alternating high daytime temperatureswith low nighttime temperatures (24◦C/19◦C).

Another example of the effect of temperature dif-ferentials is illustrated by floral movements in membersof the Liliaceae, such as tulip (Tulipa) and Crocus. Flow-ers in these plants normally open during the day andclose at night, but these movements are only slightlyaffected by light. Instead, the perianth segments ortepals1 respond to changing temperature. This is aform of thermonasty, involving a differential growthresponse of cells on the inner (or adaxial) and outer

1Tepal is the collective term for sepals and petals when thetwo share a common morphology and are indistinguishableone from the other.

(or abaxial) surfaces of the tepals. The optimum tem-perature for growth differs by approximately 10◦Cbetween the two surfaces (Figure 26.6). The openingof the flower in response to an increase in tempera-ture (Figure 26.6A) corresponds to a sharp but transientincrease in the growth rate of cells on the inner surface(Figure 26.6B). Conversely, closure following a dropin temperature (Figure 26.6C) appears to be causedby a similar change in the growth rate of cells on theouter surface (Figure 26.6D). Other investigators havereported opening following a rise of as little as 0.2◦Cfor Crocus and 1◦C to 2◦C for tulip. There are lowerlimits—Crocus, for example, will not open at tempera-tures below 8◦C. Thus if the spring days are very cold,the flowers may not open at all. Thermonasty is not lim-ited to flower parts; it has also been demonstrated for thestem angle of Phryma leptostachya, a perennial Asian herb.

26.5 TEMPERATUREINFLUENCES PLANTDISTRIBUTION

Temperature is thought to be one of the most impor-tant factors limiting the worldwide distribution of plants.Distribution limits often reflect temperature character-istics of major metabolic processes, especially photosyn-thesis. The temperature range compatible with growthof higher plants lies generally between 0◦C and 45◦C,although there are some plants that exceed either ofthese limits and within those limits temperature compat-ibility is very much species dependent. Various cultivarsof wheat (Triticum vulgare), for example, will grow attemperatures from near zero to over 40◦C, although thetemperature optimum for growth falls in the range of20◦C to 25◦C.

As a general rule, temperatures that are optimumfor growth reflect the geographical region in which thespecies originated. Thus, plants native to warm regionseither require or perform better at higher temperaturesthan those that originated in cooler areas of the world.The optimum for maize (Zea mays), a plant of tropicalorigin, is in the range 30◦C to 35◦C and it will not growbelow 12◦C to 15◦C. Garden-cress (Lepidium sativum),a temperate herb, will grow at temperatures as low as2◦C but its maximum temperature for growth is 28◦C.

The effects of temperature on physiology andmetabolism in turn influence plant distribution,called biogeography. At times, temperature-relatedmetabolic effects not only limit distribution, but havesignificant economic implications as well. Cotton(Gossypium), for example, is a southern crop in partbecause cool night temperatures in northern latitudesadversely affect fiber cell wall thickening, and thenorthern limits for maize production are very muchlimited by its inability to grow at lower temperatures.

26.5 Temperature Influences Plant Distribution 455

A.

B.

0 1 2 3 4

C.

D.

Ang

leLe

ngth

(cm

)

Hours0 1 2 3 4

Hours

Inner

110

100

90

80

70

1.49

1.47

1.45

1.43

110

100

90

80

70

1.56

1.54

1.52

Inner

Outer

Outer

FIGURE 26.6 The effect of 10◦C temperature shifts on flower opening and differentialgrowth in Tulipa flowers. (A, B) The effect of raising temperature from 7◦C to 17◦C.(C, D) The effect of lowering temperature from 20◦C to 10◦C. The temperature shiftis indicated by the arrows. A and C show floral opening and closure, respectively. Band D show the growth of cells on the inner and outer surfaces of the tepals. (FromWood, W. M. 1953. Journal of Experimental Botany 4:65–77. Reprinted by permissionof The Company of Biologists, Ltd.)

However, advances in agronomy and plant breedinghave moved the limits for maize steadily northwardover the past several decades.

Although extensive studies of temperature effectshave been conducted using controlled environmentfacilities, it is difficult to carry out field studies withany degree of precision. This is because the leaves androots of plants are commonly subject to a mosaic offluctuating temperature regimes. Leaf temperature, forexample, depends not only on daily and seasonal vari-ations in atmospheric temperature, but such factors ascloud cover, wind speed, their position in the canopy,and so forth. Root temperature depends on depth inthe soil, soil moisture content, soil structure, and otherphysical parameters of the soil. Thus, in a natural envi-ronment, individual leaves and roots may be respondingto distinctly different microclimates, each with its ownunique temperature regime.

In one study, a variety of species native to eitherthe cool coastal regions of northern California or thehot, dry desert of Death Valley were transplanted intoexperimental gardens in both locations. At the DeathValley site summer air temperatures commonly reach50◦C, a temperature that is lethal for many organisms.By contrast, average daily maximum temperatures at thecoastal site were less than 20◦C. Plants at both sites were

irrigated and fertilized so that water supply and nutri-ents were not limiting factors and their performancewith respect to growth and survival was assessed on aregular basis. On the basis of their growth responses, theplants could be grouped into three main categories: (1)those that were unable to survive the summer months;(2) those that survived but grew slowly during the sum-mer; and (3) those that grew most rapidly during thesummer months (Table 26.1).

Virtually all of the species that are native to thecool coastal climate were unable to survive the highdesert temperatures. Of the plants tested, only Tide-stromia oblongifolia, a deciduous C4 perennial nativeto Death Valley, was able to thrive in the summerdesert heat. Strikingly, T. oblongifolia was unable tosurvive the cool coastal temperatures. At the otherextreme, Atriplex glabriuscula, a C3 annual native tothe coastal region, thrived in the coastal garden butdied in the desert in spite of ample irrigation. Twoclones of the C4 species Atriplex lentiformis were alsotested—one native to the coastal regions of southernCalifornia and one that occurs naturally in Death Val-ley. In terms of biomass production, the desert cloneoutperformed the coastal clone in the Death Valleygarden; their relative performance was reversed in thecoastal garden. The relative success of A. glabriuscula and

456 Chapter 26 / Temperature: Plant Development and Distribution

TABLE 26.1 Growth responses of selectedAtriplex and Tidestromia species planted in hotdesert and cool coastal climates.

Summer Death Valley CoastalGrowth Garden Garden

1. No survival A glabriuscula T. oblongifolia2. Slow summer A. lentiformis

growth (Coastal clone)A. lentiformis(Desert clone)

3. Rapid summer T. oblongifolia A. glabriusculagrowth

A. lentiformis(Coastal clone)A. lentiformis(Desert clone)

T. oblongifolia in the two environments appeared tocorrelate with their capacity to assemble a compe-tent photosynthetic apparatus. For example, the relativegrowth rates of the two species under laboratory con-trolled conditions compared favorably with the responseof photosynthetic rate to temperature under the dameconditions. Both the maximum relative growth rate andmaximum rate of photosynthesis occurred at approx-imately 25◦C for A. glabriuscula and at approximately40◦C to 45◦C for T. oblongifolia.

Other species are more flexible with regard totemperature. Several species, including A. hymenelytra,Nerium oleander, and the creosote bush (Larrea divari-cata), were able to survive in both the desert and coolcoastal habitats, although their growth rate was not asgreat as either A. glabriuscula or T. oblongifolia. Mostof their growth was in fact accomplished during thespring or fall when temperatures were less extreme.In all three cases, growth at low or high tempera-ture under controlled conditions caused an appropriateshift (by as much as 15◦C) in the optimum temper-ature for photosynthesis. More importantly, however,there was no significant change in the maximum rateof photosynthesis, only the temperature at which themaximum rate occurred. Thus some plants exhibit asignificant degree of phenotypic plasticity with respectto photosynthesis and temperature, which enables themto survive a wider range of climatic conditions. Plantsrestricted to one climate or another apparently do notexhibit the same degree of plasticity in their metabolicreactions.

On a worldwide basis, temperature is the mostimportant factor affecting the relative distribution of C3and C4 grasses. On a smaller scale, this is illustrated bythe distribution of plants along an elevational gradient,

Ocean 300 C

330 C

130 C

150 C

AD

1.5 km

2 km

Fog and cloud

Trees

B

C

FIGURE 26.7 Adiabatic lapse and air temperature on thewindward and leeward sides of a mountain. Unsaturatedair rises from point A to point B at the adiabatic lapse rateof 1◦C/100 m. At point B the air becomes saturated withwater vapor and cools more slowly at a wet lapse rate of0.4◦C/100 m. The dry lapse rate applies as the air warmson its descent to D and is no longer saturated. Thiswarming trend is responsible for the Chinook winds thatblow out of the Rocky Mountains or the Foehn winds ofalpine Europe. (From Rosenberg, N. J. et al. 1983. Micro-climate: The Biological Environment. New York: Wiley.With permission.)

such as up a mountainside where temperature decreaseswith increasing altitude. This decrease in temperaturewith elevation is called the adiabatic lapse. As air rises,it expands and cools. The term adiabatic refers to thefact that cooling occurs without an exchange of heat.In the case of an elevational gradient, cooling as theair rises is entirely due to expansion of the air massas the pressure decreases. The heat content and, conse-quently, the temperature of a unit volume of air mass aretherefore lower. It is because of adiabatic lapse that thetemperature gradient remains stable and the less-densecooler air does not descend from high in the mountainsto displace the warmer air in the valleys. The adiabaticlapse rate for dry air is constant at about 1◦C/100 melevation (Figure 26.7). The lapse rate for moist air(wet lapse rate) is more variable and lower than the drylapse rate because condensation of water vapor releasesheat.

P. W. Rundel has studied the distribution of C3and C4 grasses along an elevational gradient in Hawaii.Rundel found a sharp transition in the distributionof the two photosynthetic types at about 1400 m. C4grasses were predominant at warmer, drier elevationsbelow 1400 m, while in the cooler, moist environmentabove 1400 m the C3 grasses were predominant. Themidpoint of the transition zone is the elevation wherethe maximum daily temperature for the warmest monthof the year is approximately 21◦C. Similar distributionsof C4 and C3 grasses have been reported in other

Further Reading 457

elevational studies carried out in Africa and Costa Rica,and in latitudinal gradients in North America. In thelatter case, the transition temperatures are slightly lower,but the principle is still valid.

SUMMARY

All living organisms can be broadly classified accord-ing to their ability to withstand temperature. Psy-chrophiles grow optimally at temperatures of 0◦C to10◦C; mesophiles, 10◦C to 30◦C; and thermophiles,30◦C to 65◦C. Most higher plants are mesophiles,although plants will generally survive temperaturesbetween 0◦C and 45◦C. Temperature limits generallyreflect the freezing point of water on the low side anddenaturation of protein on the high side.

Plants also use temperature as a cue in their devel-opmental and survival strategies. Decreasing tempera-ture in the autumn in concert with photoperiod inducesdormancy in buds, characterized by low respiratory rateand an inability to grow even if temperature, oxygen,and water supply are adequate. Most dormant budshave a chilling requirement that must be met beforedormancy can be broken and growth renewed. Dor-mancy is also a property of many seeds, a situationin which the seed fails to germinate because of envi-ronmental and physiological limitations. Seed coatsmay interfere with water uptake or oxygen uptake, ormay contain inhibitors that must be broken down orleached out before germination can proceed. Manyseeds require alternating temperatures or a period oflow temperature (pre-chilling or stratification) to breakdormancy.

Some plants, such as tomato, grow poorly at con-stant temperature, but require alternating day–nighttemperatures (thermoperiodism) for optimum growth.In others, such as tulip and Crocus, changing tem-perature regulates the opening and closure of floralpetals.

Temperature is a principal factor in the distribu-tion of plants, or biogeography. Survival in extremeenvironments appears to be due to intrinsic differencesbetween species in the temperature dependence of theirgrowth responses and photosynthetic metabolism. Ina study of elevational gradients up a mountainside, asharp transition was found between C4 species (in thewarmer, drier, lower elevations) and C3 species (in thecooler, moister, higher elevations). It is clear that tem-perature stability of principal metabolic pathways is asignificant determinant in plant distribution.

CHAPTER REVIEW

1. In what ways does temperature influence physio-logical processes? Does temperature interact withother environmental factors? If so, which ones?

2. How does temperature influence the geographicaldistribution of plants? What modifications mightyou expect to find in plants adapted to high-temperature habitats? To plants in arctic or alpinehabitats?

3. Review the distinction between vernalization andstratification.

4. Dormant buds and seeds normally require an ex-tended treatment of some sort in order to breakdormancy. What is the survival value of such arequirement?

5. In what way(s) do the daily movements of floralpetals, such as tulip and Crocus, differ from themovements of bean leaves?

FURTHER READING

Bewley, J. D., M. Black. 1994. Seeds: Physiology of Developmentand Germination. New York: Plenum.

Bjorkman, O., 1980. The response of photosynthesis to tem-perature. In: J. Grace, E. D. Ford, P. G. Jarvis (eds).Plants and Their Atmospheric Environment. Oxford: Black-well Scientific Publications, pp. 273–301.

Dickinson, R. E. 1987. The Geophysiology of Amazonia. Vegeta-tion and Climate Interactions. New York: Wiley.

Fitter, A. H., R. K. M. Hay. 1987. Environmental Physiology ofPlants. 2nd ed. London: Academic Press.

Gash, J. H. C., C. A. Nobre, J. M. Roberts, R. L. Victoria.1996. Amazonian Deforestation and Climate. Chichester:Wiley.

Gibson, A. C. 1996. Structure-function Relations of WarmDesert Plants. Berlin: Springer Verlag.

Jacobs, M. 1988. The Tropical Rain Forest: A First Encounter.Berlin: Springer-Verlag.

Lang. G. A. 1987. Dormancy: A new universal terminology.HortScience 22:817–820.

Rhode, A., R. P. Bhalerao. 2007. Plant dormancy in theperennial context. Trends in Plant Science 12:217–223.

Rosenberg, N. J., B. L. Blad, S. B. Verma. 1983. Microclimate:The Biological Environment. 2nd ed. New York: Wiley.

Rundel, P. W. 1980. The ecological distribution of C4and C3 grasses in the Hawaiian Islands. Oecologia45:354–359.

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respiration

amino acids

Alkaloids

TerpenoidsSterols

Cyanogenicglycosides

GlucosinolatesCardiac

glycosides

Saponins

Lignin

Phenols

Flavonoids Tannins

acetyl − CoA

malonyl − CoA

fatty acidslipids

protein

CO2+ H2O

photosynthesis

sugars

27Secondary Metabolites

The sum of all of the chemical reactions that take placein an organism is called metabolism. Some aspectsof metabolism, such as the metabolism of carbon andnitrogen assimilation and energy conversions, have beenaddressed in earlier chapters. Most of that carbon, nitro-gen, and energy ends up in molecules that are commonto all cells and are required for the proper functioning ofcells and organisms. These molecules, e.g., lipids, pro-teins, nucleic acids, and carbohydrates, are called primarymetabolites (see Appendix). Unlike animals, however,most plants divert a significant proportion of assim-ilated carbon and energy to the synthesis of organicmolecules that may have no obvious role in normalcell function. These molecules are known as secondarymetabolites.

In this chapter, we examine some of the broaderaspects of secondary metabolites. The focus will be thebiosynthesis, physiology, and ecological roles of fourmajor classes of secondary metabolites:

• terpenes, including hormones, pigments, essentialoils, steroids, and rubber,

• phenolic compounds, including coumarins, flavo-noids, lignin, and tannins,

• glycosides, including saponins, cardiac glycosides,cyanogenic glycosides, and glucosinolates, and

• alkaloids.

Some secondary metabolites are also involved indefense against invading pathogens, a subject that isalso addressed in this chapter.

27.1 SECONDARY METABOLITES:A.K.A NATURAL PRODUCTS

The distinction between primary and secondarymetabolites is not always easily made. At the biosyn-thetic level, primary and secondary metabolites sharemany of the same intermediates and are derived fromthe same core metabolic pathways (Figure 27.1). Inthe strictest sense, however, secondary metabolitesare not part of the essential molecular structure orfunction of the cell. Secondary metabolites generally,but not always, occur in relatively low quantities andtheir production may be widespread or restricted toparticular families, genera, or even species. Also knownas natural products, these novel phytochemicals wereof little interest to biologists because of their apparentlack of biological significance. They were known,however, to have significant economic and medicinalvalue and were thus of more than a passing interest tonatural products chemists. Natural products have founduse in antiquity as folk remedies, soaps, and essences.They include drugs and other medicinal products,

459

460 Chapter 27 / Secondary Metabolites

FIGURE 27.1 A schematic to illustratebiosynthetic relationships between prin-cipal primary and secondary metabolites(circled).

respiration

amino acids

Alkaloids

TerpenoidsSterols

Cyanogenicglycosides

GlucosinolatesCardiac

glycosides

Saponins

Lignin

Phenols

Flavonoids Tannins

acetyl − CoA

malonyl − CoA

fatty acidslipids

protein

CO2+ H2O

photosynthesis

sugars

dyestuffs, feedstocks for chemical industries (gums,resins, rubber), and a variety of substances used to flavorfood and drink. In recent years, however, it has becomeincreasingly evident that many natural products dohave significant ecological functions, such as protectionagainst microbial or insect attack.

27.2 TERPENES

27.2.1 THE TERPENES AREA CHEMICALLY ANDFUNCTIONALLY DIVERSEGROUP OF MOLECULES

With nearly 15,000 structures known, terpenoids areprobably the largest and most diverse class of organiccompounds found in plants. As discussed earlier inChapter 19, the unifying feature of terpenes is thatthey are generally lipophilic polymers based on the sim-ple 5-carbon unit 2-methyl-1,3-butadiene, or isoprene,which is derived via either the mevalonic acid pathwayor the MEP pathway (Chapter 19, Section 19.3).

Terpenes can be grouped into several classes, basedon the number of carbon atoms (Figure 27.2). Thislarge chemical diversity arises from the number of basicunits in the chain and the various ways in which theyare assembled. Formation of cyclic structures, addi-tion of oxygen-containing functions, and conjugationwith sugars or other molecules all add to the possiblecomplexity. The name terpenoid derives from the factthat the first compounds in the group were isolatedfrom turpentine (Ger. terpentin), an essential oil (chieflypinene) distilled from the resins of several coniferoustrees.

The terpene family includes hormones (gibberellinsand abscisic acid): the carotenoid pigments (carotene andxanthophyll); sterols (e.g., ergosterol, sitosterol, choles-terol) and sterol derivatives (e.g., cardiac glycosides);latex (the basis for natural rubber); and many of theessential oils that give plants their distinctive odors andflavors. Cytokinin hormones and chlorophyll, althoughnot terpenes per se, do contain terpenoid side chains. Itis apparent from this list that many terpenes have signifi-cant commercial value as well as important physiologicalroles. Many terpenes and terpene derivatives may beconsidered primary metabolites. The hormones abscisicacid and gibberellin, the carotenoid and chlorophyllpigments, and sterols (steroid alcohols) all play signifi-cant roles in plant growth and development. The vastmajority of terpenes, however, are secondary metabo-lites, many of which appear to act as toxins or feedingdeterrents to herbivorous insects.

27.2.2 TERPENES ARE CONSTITUENTSOF ESSENTIAL OILS

Many plants, such as citrus, mint, Eucalyptus, and vari-ous herbs (sage, thyme, etc.), produce complex mixturesof alcohols, aldehydes, ketones, and terpenoids, knowngenerally as essential oils (essence, as in perfume).Essential oils are responsible for the characteristic odorsand flavors of these plants but they are also knownto have insect-repellant properties. The terpenes andterpene derivatives found in essential oils are predom-inantly hemi-, mono-, and sesquiterpenes, which canbe moderately to highly volatile. Several examples areshown in Figure 27.3. In most plants, the essential oilsare synthesized in special glandular trichomes (hairs) onthe leaf surface (Figure 27.4), although the essential oils

CH3 C CH CH2

CH3 C CH (CH2)2 C CH CH2OH

COOH

CH3 CH3

OH

HOCH2

CH2OH

HO

Numberof

Carbons Class Example

40 Tetraterpenoid -carotene(a carotenoid)

30 Triterpenoid

30 Triterpenoid

20 Diterpenoid

15 Sesquiterpenoid

10 Cyclic monoterpenoid

10 Monoterpenoid

5 Hemiterpenoid Tigilic acid

Geraniol

Farnesol (widespread)

Phytol (chlorophyll)

Squalene(a steroid precursor)

Stigmasterol(a sterol)

Menthol (peppermint oil)

FIGURE 27.2 Terpenoids are classified according to the number of carbon atoms inthe basic skeleton.

461

462 Chapter 27 / Secondary Metabolites

HC

Geranial(Ctenium aromaticum)

(lemon grass)

CH3

CH3

CH3

CH3

CH3 CH3H3C

H3C

H3C

H3C

H3C

CO

O

H

H

O

CHCH2CH2OHH3C

H3CCHCH2C

HOCH2

iso-Amyl Alcohol(Mentha, Eucalyptus)

iso-Valeraldehyde(Eucalyptus)

1:8 Cineole(Artemesia)

Farnesol(widespread)

FIGURE 27.3 Representative terpenes that commonlyoccur in essential oils include: hemiterpenes (isoamylalcohol, isovaleraldehyde), monoterpenes (geranial, cine-ole), and the sesquiterpene, farnesol.

of citrus are produced by glands in the peel. Theresins of certain conifers, for example, also accumu-late mixtures of terpenes, including the monoterpenes,α- and β-pinene, and myrcene (Figure 27.5).

27.2.3 STEROIDS AND STEROLS ARETETRACYCLIC TRITERPENOIDS

Steroids and sterols are synthesized from the acyclictriterpene squalene, although they generally are modi-fied and have fewer than 30 carbon atoms. Steroids withan alcohol group, which is the case with practically allplant steroids, are known as sterols. The most abundantsterols in higher plants are stigmasterol and sitosterol(Figure 27.6), which often make up more than 70 percentof the total sterols. However, plants also contain a largenumber of the more than 150 other sterols known tooccur in nature. Plant sterols include cholesterol which,although widespread in occurrence, is present in only

Extracellularcavity

Cuticle

Secretory cells

Stalk cell

Epidermis

Mesophyll

FIGURE 27.4 A schematic diagram of an epidermal glan-dular hair in cross-section. Essential oils are produced inthe secretory cells and accumulated in a cavity that formsbetween the secretory cells and the overlying cuticle.Glandular hairs are found on the leaf surface, where it isthought they might serve to deter feeding by herbivores.

trace quantities. The extremely low level of cholesterolallows plant oils to be marketed as ‘‘cholesterol-free.’’

Sterols are constituents of plant membranes, whichis perhaps their most important known function inplants. Because sterols are planar molecules, their pack-ing properties are different from phospholipids thatmake up the bulk of the membrane bilayer. Sterols packmore tightly than phospholipids and therefore tend toincrease the viscosity and enhance the stability of mem-branes. Otherwise, little is known about the function ofsterols in plants. Unlike the steroid hormones in ani-mals, there is no known hormonal role for sterols inplant development. Some sterols may have a protectivefunction, such as the phytoecdysones, which have astructure similar to the insect molting hormones. Wheningested by insect herbivores, phytoecdysones disruptthe insect’s molting cycle. Other sterols are present asglycosides, which give rise to a number of interest-ing and economically significant secondary products.Steroid glycosides are discussed below.

27.2.4 POLYTERPENES INCLUDETHE CAROTENOID PIGMENTSAND NATURAL RUBBER

Larger terpenes include the tetraterpenes (40-carbon)and the polyterpenes. The principal tetraterpenes arethe carotenoid family of pigments (Chapter 6). The onlyimportant isoprene derivatives with a greater molecu-lar mass than the tetraterpenes are rubber and gutta.Rubber is a polymer consisting of up to 15,000 isopen-tenyl units. The polymer may be linear, as shown inFigure 27.7, or cross-linked into more complex config-urations. The only difference between rubber and guttais the configuration of the double bonds. In rubber the

27.3 Glycosides 463

CH3

α-Pinene

CH3

CH3

OH

H3C

Menthol

CH3

CH2

H3C

Myrcene

CH2

FIGURE 27.5 Pinene, myrcene, and men-thol are monoterpenes. Pinene andmyrcene are found in the resins of someconifers. Menthol is the principal con-stituent of the essential oil of peppermint(Mentha piperta). Pinene also has insecti-cidal properties.

double bonds are all cis configurations, while in guttathe double bonds are all trans.

In the plant, rubber occurs as small particles sus-pended in a milky-white emulsion called latex. Latexproduction is widespread in plants, with estimates rang-ing from a few hundred to several thousand species thatproduce latex in some form. The principal commer-cial source is Hevea brasiliensis, a rubber tree native tothe Amazon rainforest. Others include the ornamen-tal rubber tree (Ficus elastica), milkweed (Asclepias), andthe Russian dandelion (Taraxacum Kok-saghyz). Latexcontains about 30 to 40 percent rubber and 50 per-cent water. The balance is a complex mixture of resins,terpenes, proteins, and sugars. In most plants, latex is

CH3

CH3

CH3

CH3

CH3H3C

HO

Stigmasterol

CH3

CH3

CH3

CH3

CH3H3C

HO

SitosterolFIGURE 27.6 Stigmasterol and sitosterol differ only inthe presence or absence of a double bond (highlighted).These are the two most abundant plant sterols.

CC

H

CH2

CH3

CH3

CC

H

CH2

CH3

CH2

CC

H

CH3

CH3

CH2n

FIGURE 27.7 Rubber is a linear polymer of isoprene unitswhere the value of n may range from a few hundred toseveral thousand.

produced in the phloem, accumulating in a series oflong, interconnected vessels called laticifers.

The best-known source of gutta is a desert shrub,Parthenium argentatum, which grows in northern Mexicoand southwestern United States. Parthenium (commonlyknown as guayule) may contain as much as 20 percentlatex by weight, which is stored not in laticifers but inthe vacuoles of stem and root cells. Guayule was at onetime a significant commercial source of gutta for use inrubber products. However, while a single rubber tree, ifproperly tapped, can continue to produce for up to 30years, guayule plants must be harvested (and, of course,replanted) annually.

Finally, there is a connection between terpenesand air pollution. Many of the essential oils, especiallyhemiterpenes, monoterpenes, and sesquiterpenes, arehighly volatile and are given off in large quantities byplants, particularly during warm weather. Known gen-erally as volatile organic carbon (VOC), these naturalemissions from plants contribute to the formation ofhaze and cloud, and are involved in the formation oftoxic tropospheric ozone.

27.3 GLYCOSIDES

Some of the more interesting, if not important,derivatives synthesized by plants are glycosides. Mostglycosides are thought to function as deterrents toherbivores. The term glycoside (Gr. glykys, sweet)refers to the bond formed (called a glycosidic bond)when a sugar molecule condenses with anothermolecule containing a hydroxyl group. Sugars may formglycosidic bonds with other sugars, such as when linked

464 Chapter 27 / Secondary Metabolites

together to form polysaccharides, or with hydroxylgroups on noncarbohydrate molecules, such as steroidsor amino acids. The sugar most commonly found inglycosides is glucose, although specific glycosides oftencontain rare sugars.

Three particularly interesting glycosides are thesaponins, the cardiac glycosides (cardenolides), and thecyanogenic glycosides. A fourth family, the glucosino-lates, although technically not glycosides, are a similarstructure and so are included here.

27.3.1 SAPONINS ARE TERPENEGLYCOSIDES WITH DETERGENTPROPERTIES

Saponins may take the form of (1) steroid glycosides,(2) steroid-alkaloid glycosides, or (3) triterpene glyco-sides (Figure 27.8). Saponins may also occur as aglycones(e.g., the terpene without the sugar), which are knownas sapogenins. In much the same way as soap, whichis the sodium salt of a fatty acid, the combination ofa relatively hydrophobic triterpene with a hydrophilic

sugar gives saponins the properties of a surfactant ordetergent. When agitated in water, saponins form a sta-ble soapy foam. The name saponin is in fact derived fromSaponaria (soapwort), which at one time was employedas a soap substitute.

The principal role of saponins appears to be asa preformed defense against attack by fungi. Evidenceindicates that saponins form complexes with sterols con-taining an unsubstituted 3-β-hydroxyl group. When thesaponins react with sterols in the membranes of invadingfungal hyphae, the result is a loss of membrane integrity.In a classic example of one-upmanship, however, manypathogenic fungi have developed strategies, such as thedevelopment of detoxifying enzymes, for circumvent-ing this defense mechanism. Oat (Avena), for example,produces a triterpenoid saponin, avenacin A-1, whichis localized in the root epidermal cells and effectivelyprotects against an invasion by a fungal pathogen (Gaeu-mannomyces graminis var. tritici) that infects the roots ofboth wheat (Triticum) and barley (Hordeum). However,another strain of G. graminis (var. avenae) produces anenzyme, avenacinase, that detoxifies avenacin A-1 and

FIGURE 27.8 Saponins are triterpenoids or steroidscontaining one or more sugar units. Medigenicacid glucoside is a triterpenoid saponin from alfalfa(Medicago sativa). Disogenin glycoside is a steroidalsaponin isolated from clover (Meliotus spp.). Theaddition of a hydrophilic sugar group to a normallyhydrophobic terpenoid gives saponins surfactantproperties similar to soap.

O

CH3

H3C

H3C

CH3

CH3

CH3

CH3 COOH

COOH

Glucose

HO

O

CH3

CH3

CH3

Glucose

Rhamnose

Rhamnose

O

O

Medigenic acid glucoside

Disogenin-glycoside

27.3 Glycosides 465

allows the pathogen to invade oats as well as wheatand rye.

The effect of saponins on eukaryotic membranesis highly nonspecific and it is not clear how plantsprotect their own membranes against the deleteriouseffects of their own saponins. One possibility is that thesaponins are stored in the form of a biologically inac-tive molecule, called a bisdesmosidic saponin, whichhas two sugar chains rather than one. When underattack, the inactive form may be converted to the activemonodesmosidic form by hydrolytic removal of thesecond sugar chain. Alternatively, biologically active,monodesmosidic saponins may be sequestered in vac-uoles or organelles whose membranes contain a highproportion of sterols with a protected 3-β-hydroxylposition.

The effect of saponins on animals is somewhat vari-able. While not significantly toxic to mammals, saponinsdo have a bitter, acrid taste and will cause severe gas-tric irritation if ingested. Saponins will hemolyze redblood cells, however, if injected into the bloodstream.This action is presumably because of their detergentproperties and their ability to disrupt membranes gen-erally. On the other hand, saponins are highly toxic tofish and have been used as fish poisons. Saponins havealso been implicated in reports of livestock poisoning.Alfalfa saponins, for example, can cause digestive prob-lems and bloating in cattle. At the same time, there arereports that saponins contained in alfalfa sprouts willlower serum cholesterol levels. Commercially, saponinsfrom the bark of Quillaja saponaria have been used assurfactants in photographic film, in shampoos, liquiddetergents, toothpastes, and beverages (as emulsifiers).The saponin glyscyrrhizin from licorice (Glyscyrrhizaglabra) has been used in medicines and as a sweetenerand flavor-enhancer in foods and cigarettes.

27.3.2 CARDIAC GLYCOSIDES AREHIGHLY TOXIC STEROIDGLYCOSIDES

The cardiac glycosides (or, cardenolides) are struc-turally similar to the steroid saponins and have similardetergent properties. They are distinguished from othersteroid glycosides by the presence of a lactone ring(attached at C17) and the rare sugars (found almostexclusively in this group of steroids) that form the gly-coside (Figure 27.9). Like the saponins, cardenolidesoccur naturally as either the glycoside or the aglycone(or genic).

The cardenolides have a wide distribution; they havebeen recorded in more than 200 species representing55 genera and 12 families and are a principal agentin accidental poisonings of humans. Perhaps the bestknown is digitalis, a mixture of cardenolides extractedfrom the seeds, leaves, and roots of purple foxglove,

Digitalis purpurea or Grecian foxglove, D. lanata. Thetwo principal cardenolides in digitalis are digitoxin andits close analog digoxin. Digitalis is also the source of asaponin, digitonin.

Since the late eighteenth century, digitalis has beenused for its therapeutic value in treating heart conditionssuch as atherosclerosis. Because they disrupt the heartmuscle Na+/K+-ATPase pumps (hence the appellationcardiac), cardenolides are highly toxic to vertebrates. Theextreme toxicity of cardenolides has long been exploitedby African hunters, who coated their arrows and spearswith cardenolide-rich extract from plants. In therapeuticuse, however, carefully regulated doses can both slowand strengthen the heartbeat. Unfortunately, the lethaland therapeutic doses are very close, so the therapy mustbe carefully monitored.

Other common sources of cardenolides are themilkweeds, Asclepias and Calotropis. These two speciesare known as ‘‘milkweeds’’ because they produce amilky-white, cardenolide-rich latex. The milkweeds areparticularly interesting because they are the principalhost for ovipositing monarch butterflies. The emerg-ing larvae feed on the milkweed leaves and sequesterthe cardenolides without ill effect. The cardenolidesare retained through metamorphosis and are presentin the adult monarchs. When birds, such as blue jays,

H3C C

OH

H

C

OH

H

C

OH

H

C

H

C

H O

O

H

CH

O

CO

OH

CH3 CH2

C

CH3

Sugars

Digitoxin

DigitoxoseFIGURE 27.9 Digitoxin, a cardiac glycoside. The sugarcomponent of digitoxin consists of 1 molecule of glucoseand 1 molecule of acetyl-digitoxose. The structure ofdigitoxose, one of the rare sugars found in cardiac glyco-sides, is shown below.

466 Chapter 27 / Secondary Metabolites

attempt to feed on monarchs, the accumulated carede-nolides induce an emetic reaction that forces the bird tovomit. The bird then wisely avoids attempting to feedon monarch larvae for some time.

27.3.3 CYANOGENIC GLYCOSIDESARE A NATURAL SOURCE OFHYDROGEN CYANIDE

It might seem odd that plants synthesize chemicals capa-ble of releasing deadly hydrogen cyanide or prussic acid(HCN), but more than 60 different cyanogenic com-pounds of plant origin have been described from morethan a dozen plant families. Predominant among theseare the cyanogenic glycosides. A common cyanogenicglycoside is amygdalin (Figure 27.10), which occursin many representatives of the family Rosaceae. It isfound in the seeds of apples and pears and in the bark,leaves, and seed of the stone fruits (apricot, peaches,plums, cherries). Most cyanogenic glycosides appear tobe derived from one of four amino acids (phenylala-nine, tyrosine, valine, and isoleucine) or from nicotinicacid. Intact cyanogenic glycosides are not themselvestoxic, but when the plant is damaged by a herbivore,the glycoside undergoes an enzymatic breakdown andcyanide is released. Cyanide, a noncompetitive inhibitorof cytochrome oxidase, is acutely toxic.

The enzymatic breakdown of cyanogenic glycosidesis a two-step process (Figure 27.10). First, the sugars arereleased by the enzyme β-glycosidase. The resultinghydroxynitrile is moderately unstable and will slowlydecompose, liberating HCN in the process. Normally,however, decomposition is accelerated by a secondenzyme, hydroxynitrile lyase. Enzymatic release ofcyanide does not normally occur in intact plants becausethe enzymes and the substrate are spatially separated.In some cases, separation is maintained within thecell, but in others, the enzymes are in one cell andthe cyanogenic glycosides in another. In Sorghum, forexample, the cyanogenic glycoside dhurrin is synthe-sized and stored in epidermal cells, while the glycosidase

and lyase enzymes are found in the mesophyll cells. Onlywhen the tissue is crushed and the contents of the twocells are mixed will cyanogenesis occur.

There is some evidence that the presence ofcyanogenic glycosides deters feeding by insects andother herbivores, although most animals have theability to detoxify small quantities of cyanide. Clearlythe effectiveness of cyanogenic glycosides as a deterrentdepends on many factors, such as the amount present,the rate of release of cyanide, and the ability of theanimal to detoxify. The level of cyanogenic glycosidesin plants is highly variable, influenced by both geneticcontrol and environmental stress. The latter is aconcern when using Sorghum for livestock forage.Dhurrin accumulates rapidly and can cause livestockpoisoning when Sorghum plants are stressed by droughtor frost.

Many common food plants naturally containcyanogenic glycosides in concentration sufficiently lowthat they are not normally a health hazard. Theseinclude soy and other beans (Fabaceae); apples, apricots,peaches, plums, and other fruits in the family Rosaceae;and flax seed (Linum), which is a popular healthfood. One food source that contains large amountsof cyanogenic glycosides is cassava, a potato-like rootof the tropical plant Manihot esculenta. Cassava, alsoknown as manioc or, in North America, tapioca, is amajor source of starch for millions of people in tropicalcountries. However, poisoning is avoided by carefulpreparation of the plant. This includes grinding theroot and expressing the fluids, or boiling the root inseveral changes of water.

27.3.4 GLUCOSINOLATES ARE SULFUR-CONTAINING PRECURSORSTO MUSTARD OILS

Glucosinolates are found primarily in the mustardfamily (Brassicaceae) and related families in the orderCapparales. They are precursors to the mustard oils,an economically important class of flavor constituents

FIGURE 27.10 Amygdalin is a cyanogenic glycosidefound in large quantities in seeds of common fruitsin the family Rosaceae. Hydrolysis of amygdalin is atwo-step process, resulting in the release of highlytoxic hydrocyanic acid (HCN).

C O

CH

GluO Glu

CN

C

C

H

CH

OH

CN

+ 2 Glucose

+ HCN

Cβ-Glycosidase

MandelonitrileAmydalin

Benzaldehyde Hydrocyanic acid

27.4 Phenylpropanoids 467

that gives the pungent taste to condiments such asmustards and horseradish as well as the distinctive flavorof cabbages, broccoli, and cauliflower.

All glucosinolates are thioglucosides (thio, sulfur)with the general structure shown in Figure 27.11A.The sugar is always glucose. The diversity encoun-tered in structure and properties is due to the Rgroup, which may range from a simple methyl groupto large linear or branched chains containing aromaticor heterocyclic structures. The biological activity ofglucosinolates depends primarily on their hydrolysis tomustard oils (Figure 27.11B). Hydrolysis of glucosi-nolates is catalyzed by an enzyme called myrosinase(a thioglucosidase). The hydrolysis product is unstableand immediately undergoes a rearrangement to form athiocyanate or isothiocyanate. Like the cyanogenic gly-cosides, glucosinolates are spatially separated from thehydrolytic enzymes so that the mustard oils are normallyformed only when the cells are disrupted, allowing theenzyme and substrate to come together. As with otherdefense compounds, some herbivores are deterred or

repelled by the presence of glucosinolates in a plant,while others have adapted to use the glucosinolatesor mustard oils as attractants to stimulate feeding andovipositing.

Glucosinolates, or rather their absence, have hada significant impact on the oilseed industry. Rape seed(principally Brassica napus) is a good source of veg-etable oil, but its high content of glucosinolate togetherwith high erucic acid (a 22-carbon fatty acid) gives theoil undesirable taste and poor storage properties. Newstrains have been bred with low glucosinolates and erucicacid. These strains, called canola in order to distin-guish them from normal rape, are now an economicallyimportant oil source.

27.4 PHENYLPROPANOIDS

Aromatic amino acids may be directed toward eitherprimary or secondary metabolism. Also known as phe-nolics, or polyphenols, phenylpropanoids are a large

(thioglucosidase)

spontaneousrearrangement

glucose

N O SO3

CHCH2CCH2

S

–

N O

N

SO3

CHCH2CCH2

CHCH2CH2 C S

S–

–

iso-thiocyanate

+ SO42–

SCHCH2CH2 C N

thiocyanate

+ SO42–

+ S + SO4CHCH2CH2 C N

nitrile

2–

Sinigrin

glucose

N O SO3

CRS

–

A.

B.

FIGURE 27.11 (A) All glucosinolates are thioglu-cosides with the same basic structure. In athioglucoside the sugar is linked to the rest ofthe molecule via a sulfur atom. Variation is intro-duced by the composition of the R group. (B)Enzymatic removal of glucose from a glucosi-nolate creates an unstable product that sponta-neously rearranges to form the pungent mustardoils in the thiocyanate, isothiocyanate, or nitrileforms.

468 Chapter 27 / Secondary Metabolites

OH

FIGURE 27.12 Phenylpropanoids are derivatives of thesimple hydroxylated aromatic ring, phenol.

family of secondary metabolites derived from the aro-matic amino acids. Phenylpropanoids are a chemicallydiverse family of compounds ranging from simple phe-nolic acids to very large and complex polymers suchas tannins and lignin. Also included are the flavonoidpigments that were described earlier in Chapter 6.The basic structure is phenol, a hydroxylated aromaticring (Figure 27.12). As with other secondary products,many phenolics appear to be involved in plant/herbivoreinteractions. Some (e.g., lignin) are important structuralcomponents, while others appear to be simply metabolicend-products with no obvious function.

27.4.1 SHIKIMIC ACID IS A KEYINTERMEDIATE IN THESYNTHESIS OF BOTH AROMATICAMINO ACIDS ANDPHENYLPROPANOIDS

The biosynthesis of most phenylpropanoids beginswith the aromatic amino acids phenylalanine, tyrosine,and tryptophan. These aromatic amino acids are,in turn, synthesized from phosphoenolpyruvate anderythrose-4-phosphate by a sequence of reactionsknown as the shikimic acid pathway (Figure 27.13).The shikimic acid pathway is common to bacteria,fungi, and plants, but is not found in animals.Phenylalanine and tryptophan are consequently amongthe 10 amino acids considered essential for animals(including humans) and represent the principal sourceof all aromatic molecules in animals. Tyrosine is notclassified as essential because animals can synthesize itby hydroxylation of phenylalanine.

Synthesis of the aromatic amino acids begins withthe condensation of one molecule of erythrose-4-P,from the pentose-phosphate respiratory pathway, withone molecule of phosphoenolpyruvate (PEP) from gly-colysis. The resulting 7-carbon sugar is then cyclizedand reduced to form shikimate. Shikimate is then con-verted to chorismate, a critical branch point that leadseither to phenylalanine and tyrosine or to tryptophan.The shikimic acid pathway is an excellent example offeedback regulation, where the concentrations of prod-uct molecules regulate the flow of carbon throughkey enzymes. The first key enzyme in the pathwayis the aldolase that catalyzes the initial condensationerythrose-4-P with PEP. Aldolase is inhibited by all

three end-products: tyrosine, phenylalanine, and trypto-phan. In a similar manner, the conversion of chorismateto prephenate is inhibited by both phenylalanine andtyrosine, while the conversion of chorismate to anthrani-late is inhibited by tryptophan.

Another step of interest in the pathway isthe second step in the conversion of shikimate to3-enolpyruvylshikimate-5-P. The enzyme 3-enolpy-ruvylshikimate-5-phosphate synthase (EPSPS) isinhibited by the herbicide glyphosate (marketedcommercially as RoundUp™). Plants treated withglyphosate will die because they are unable to synthesizethe aromatic amino acids and their derivatives,especially protein. Thus glyphosate-treated plantseffectively die of protein starvation.

27.4.2 THE SIMPLEST PHENOLICMOLECULES ARE ESSENTIALLYDEAMINATED VERSIONS OF THECORRESPONDING AMINO ACIDS

Synthesis of most secondary phenolic products beginswith the deamination of phenylalanine to cinnamic acid(Figures 27.14, 27.15). The enzyme that catalyzes thisreaction, phenylalanine ammonia lyase (PAL), is akey enzyme because it effectively controls the diversionof carbon from primary metabolism, such as proteinsynthesis, into the production of phenylpropanoids.An alternate route, the deamination of tyrosine top-OH-cinnamic acid, appears to be limited largely, ifnot exclusively, to grasses.

Cinnamic acid is readily converted top-OH-cinnamic acid by the addition of a hydroxylgroup (Figure 27.15). The sequential addition ofhydroxyl and methoxy groups gives rise to caffeic acidand ferulic acid, respectively. None of these four simplephenols appear to accumulate to any extent. Theirprincipal function appears to be as precursors to morecomplex derivatives such as coumarins, lignin, tannins,flavonoids, and isoflavonoids.

27.4.3 COUMARINS AND COUMARINDERIVATIVES FUNCTION ASANTICOAGULANTS

The coumarins (Figure 27.16) are a widespread fam-ily of lactones called benzopyranones. More than 1,500examples are known, from more than 800 species ofplants. The biosynthesis of coumarins is not well under-stood because the putative enzymes and their geneshave yet to be isolated. It is most likely, however, thatthe two simplest forms, coumarin and 7-OH-coumarin(umbelliferone), are formed by an ortho-hydroxylationof cinnamic acid and p-coumaric acid, respectively, fol-lowed by ring closure (Figure 27.17).

SHIKIMATE

CHORISMATE

TYROSINE

TRYPTOPHAN

PHENYLALANINE

NH3+

+NH3+

HO

H2C CH COO_

NH3+

H2C CH COO_

OHOHHO

COO_

CH2

HO

OC COO

_

COO_

CH2 CH

NH3+

NH

COO_

Erythrose-4-phospahate + Phosphoenolpyruvate

NADPH

NADP+

ATP

EPSPS

ADP

Glyphosphate

Pi

32

Anthranilate

Arogenate

1

FIGURE 27.13 The shikimic acid pathway for biosynthesis of aromatic amino acidsin plants. Initial precursors are erythrose-4-P from the pentose-phosphate path-way and phosphoenolpyruvate from glycolysis. Enzymes indicated as 1, 2, and 3 aresubject to feedback inhibition and thus are important regulatory enzymes in the path-way. Those enzymes are (1) 3-deoxyarabino-heptulosonate-7-phosphate synthase;(2) anthranilate synthase; (3) chorismate mutase. The enzyme EPSP synthase(EPSPS), which catalyzes the second of three reactions in the conversion of shikimateto chorismate, is inhibited by the herbicide glyphosate. Glyphosphate thus preventsthe synthesis of the amino acids tyrosine and phenylalanine.

469

470 Chapter 27 / Secondary Metabolites

P

P

4-OH-

P

Pentose- pathway Glycolysis

Erythrose-4- -enol Pyruvate

Shikimic acidpathway

Phenylalanine

Tryptophan

Gallic acid

Tyrosine

Cinnamic acid(ρ-Coumaric acid)

Cinnamic acid

Coumarin

Flavonoids

ρ-Coumaric acid

Ferulic acid

Lignin

Condensedtannins

PALCS

Alkaloids

Indole-3-acetic acid(IAA)

Hydrolyzabletannins

Protocatechuic acid

FIGURE 27.14 The shikimic acid pathway plays a central role in the synthesis ofnumerous primary and secondary metabolites. PAL = phenylalanine ammonia lyase.CHS = chalcone synthase.

As a family, coumarins are noted for their rolesas antimicrobial agents, feeding deterrents, and germi-nation inhibitors. The simplest example, coumarin, isthe product that gives new-mown hay its characteristicpleasantly sweet odor. Coumarin is also a constituent ofBergamot oil, an essential oil that is used to flavor pipetobacco, tea, and other products. While coumarin itselfis only mildly toxic, many of its derivatives can be highlytoxic. One derivative, dicoumarol (Figure 27.16), istypically found in moldy hay or silage containing sweetclover (Meliotus spp.). Dicoumarol causes fatal hemor-rhaging in cattle by inhibiting vitamin K, an essentialcofactor in blood clotting. The discovery of dicoumarolin the 1940s led to the development of a syntheticcoumarin derivative, Warfarin™, widely used as a rodentpoison. Scopoletin (Figure 27.16), the most prevalentcoumarin in higher plants, is often present in seed coats.It is suspected of being a germination inhibitor thatmust be leached out of the seed coat before germinationcan proceed.

The most toxic coumarin derivatives are synthe-sized not by plants, but by fungi. The fungus Aspergillusflavus commonly infects foodstuffs such as livestockfeed, peanuts, and maize (the latter especially fol-

lowing damage by the European corn borer). Theinvading Aspergillus produces a group of mycotoxinscalled aflatoxins (from A. fla. + toxin), believed to bethe most potent and carcinogenic of natural toxins(Figure 27.16). Deaths have been recorded from inges-tion of maize contaminated with as little as 6 to 15 partsper billion (or, 6 to 15 μg kg−1) of aflatoxin. Aflatox-ins have multiple effects: they are mutagenic; they bindto DNA and prevent RNA transcription; they competewith hormones for binding sites, impair the immune sys-tem, and damage the liver and kidney. Peanut productsare a common source of aflatoxin poisoning in humansand all raw, shelled peanuts in the United States must beinspected for their aflatoxin content. Aflatoxins in cattlefeed is another particular problem. When cattle ingestthe toxin, they convert it to another equally toxic formand secrete it into the milk.

27.4.4 LIGNIN IS A MAJOR STRUCTURALCOMPONENT OF SECONDARYCELL WALLS

Lignin is a highly branched polymer of threesimple phenolic alcohols known as monolignols

27.4 Phenylpropanoids 471

CHH2C COO–

NH3

Phenylalanine

+

PAL

CHHC COO–

Cinnamicacid

CHH2C COO–

NH3

Tyrosine

+

CH

OH

HC COO–

OH OH

OH

ρ-Coumaricacid

CHHC

HO

COO–

Caffeicacid

CHHC COO–

Ferulicacid

H3CO

FIGURE 27.15 Phenolic building blocks. Deamination of phenylalanine followed byhydroxylation to form p-coumaric acid are the first two steps in phenylpropanoidbiosynthesis. In grasses, p-coumaric acid may be formed directly by deaminationof tyrosine. Phenylalanine ammonia lyase (PAL) is a critically regulated enzyme thatcontrols the diversion of phenylalanine from protein biosynthesis to phenylpropanoidbiosynthesis.

(Figure 27.18). Gymnosperm lignin is comprisedmainly of coniferyl alcohol subunits while angiospermlignin is a mixture of coniferyl and sinapyl alcoholsubunits. The biosynthesis of lignin is not wellunderstood, but it is believed that the alcohols areoxidized to free radicals by the ubiquitous plant enzyme,peroxidase. The free radicals then react spontaneouslyand randomly to form polymeric lignin.

Lignin is found in cell walls, especially the sec-ondary walls of tracheary elements in the xylem. In spiteof its abundance (second only to cellulose), the struc-ture of lignin is not well understood. Lignin is a verylarge polymer; it is insoluble in water and most organicsolvents, and impossible to extract without consider-able degradation. Moreover, the three basic monomersmay link together in a variety of ways to form a multi-branched, three-dimensional structure. The complexityis so great that, like snowflakes, each lignin ‘‘molecule’’may be unique. For the same reason that monolignolsare able to form extensive linkages with each other,the lignin polymer is able to form numerous cross-linkswith other cell wall polymers. The result is a high degreeof mechanical strength and rigidity in lignified woodystems. Lignin is in fact what makes wood, wood!

Although the principal function of lignin is struc-tural, it has also been implicated as a defensive chemical.Lignin itself is not readily digested by herbivores and,because it is covalently linked to cellulose and cell wall

xyloglucans, its presence decreases the digestibility ofthese polymers as well. Also, when fungal pathogensenter host cells, they do so by enzymatically degradingthe host cell wall. Several studies have shown that ligninsand other phenolic derivatives accumulate at the site offungal penetration, presumably slowing the rate of cellwall degradation.

27.4.5 FLAVONOIDS AND STILBENESHAVE PARALLEL BIOSYNTHETICPATHWAYS

As noted earlier in Chapter 6, flavonoids include theanthocyanin pigments that serve to attract insect polli-nators and the isoflavonoids that function as phytoalex-ins, or antibacterial and antifungal agents. Flavonoidshave a role in symbiont recognition between rhizo-bia and host roots. It has also been demonstrated thatexposure of plants to UV radiation increases the con-tent of flavonoids, suggesting that flavonoids such askaempferol may offer a measure of protection by screen-ing out harmful UV-B radiation.

Flavonoids represent a very large class of pheno-lic derivatives (more than 4,500 representatives areknown), with a variety of functions. Biochemically, how-ever, all flavonoids share a common structure consistingof three rings, labeled A, B, and C (Figure 27.19).

472 Chapter 27 / Secondary Metabolites

O

Coumarin

O

O

Umbelliferone

OHO

O

Scopoletin

OHO

H3CO

O

O O

O

O

OCH3

CH2

OH OH

O

Dicoumarol

O O

Aflatoxin B1

FIGURE 27.16 Coumarins. More than 300 coumarinshave been reported from more than 70 plant fami-lies. Coumarin and 7-hydroxycoumarin (umbelliferone)are derived from cinnamic acid and p-coumaric acid,respectively. Scopoletin is probably the most widespreadcoumarin in plants. Dicoumarol is a powerful anticoag-ulant found in moldy hay. Dicoumarol derivatives andsynthetic coumarins are commonly used to thin bloodin cardiac patients. Aflatoxin B1, a mycotoxin, is amongthe most carcinogenic and toxic of naturally occurringcompounds.

Both ring B and the 3-carbon bridge that makes upring C are derived from the shikimic acid pathway viaphenylalanine and p-coumaric acid. The six carbonsthat make up the A ring are derived from malonicacid, in the form of a malonyl-coenzyme A complex,malonyl-CoA. (Malonyl-CoA is also a principal inter-mediate in fatty acid synthesis.) The key enzyme ischalcone synthase (CHS), which catalyzes the first

Cinnamic acid

Coumarin

COOH

HO -Coumaric acid

COOH

OO O7-Hydroxy coumarin

OHO

FIGURE 27.17 Biosynthesis of coumarins. Coumarin and7-hydroxycoumarin (umbelliferone) are formed fromcinnamic acid and p-coumaric acid, respectively. Biosyn-thesis involves a trans/cis-isomerization, or rotationaround the C-C double bond (arrows) in the side chain,followed by ring closure.

committed step in flavonoid biosynthesis. CHS cat-alyzes the stepwise condensation of three molecules ofmalonyl-CoA with one molecule of p-coumaryl-CoAto form 4,2′,4′,6′-tetrahydroxychalcone, or, naringeninchalcone. A chalcone is a C6—C3—C6 pattern whereinring C is not yet closed. The enzyme chalcone isomerasethen catalyzes the closure of ring C to form narin-genin. Naringenin is the precursor to flavonols suchas kaempferol and quercitin as well as the anthocyaninpigments.

Stilbenes are a group of C6—C2—C6 compoundsthat have a chemical defense role, primarily againstfungal invasions of heartwood (the distinctively coloredcore of the tree). Stilbenes are one of several phenyl-propanoid derivatives that are continually infused intothe heartwood of trees after the cells have been lignified.A defensive role is suggested by the localized depositionof stilbenes in regions of sapwood as well, when the sap-wood is attacked by insects of pathogenic fungi. The roleof stilbenes in this case appears to be one of preventingthe spread of the invading insect or pathogen.

Stilbene biosynthesis, like the flavonoids, involvesthe sequential condensation of malonyl-CoA units witheither cinnamoyl-CoA or p-coumaroyl-CoA, exceptthat the enzyme catalyzing the reaction is stilbenesynthase.

27.4.6 TANNINS DENATURE PROTEINSAND ADD AN ASTRINGENTTASTE TO FOODS

The name tannin is derived from the historic practiceof using plant extracts to ‘‘tan’’ animal hides, that is,to convert hides to leather. Such extracts contain amixture of chemically complex phenol derivatives thatbind to, and thus denature, proteins. For years, the mostcommon test for tannins was the capacity to precipitate

OO

Coumaryl alcohol Coniferyl alcohol Sinapyl alcohol

OH

OH

CH

CH

CHCH3

CH2

O

OH

OH

CH

CH

CH3

CH2

OH

OH

CH

CH

CH2

FIGURE 27.18 Monolignols are the principal lignin monomers. Extensivecross-linkages most commonly form between the ring alcohol group and thedouble-bonded carbon atoms.

OHO

OH

B

A C

O

O

Naringenin(a flavonone)

FlavonolsAnthocyanins

OHHO

OH HO

R1

R2

CoAS

B

A

OOH

OH

OH

R1

+

R2

Naringeninchalcone

Stilbenes

Stilbene synthaseChalcone synthase

Cinnamoyl CoA(R1, R2 = H)

(3) Malonyl - CoA

Coumaroyl CoA( R1 = OH, R2 = H)

C

O

O-

CH2 S - Co A

C

O

FIGURE 27.19 The flavonoid ring structure and stilbenes are both synthesized bysequential condensations of 3 molecules of malonyl-coenzyme A (CoAS) with eithercinnamoyl-coenzyme A or p-coumaroyl-coenzyme A. Malonyl-coenzyme A is inturn formed by the carboxylation of acetyl-CoA, the first step in the formation oflong-chain fatty acids.

473

474 Chapter 27 / Secondary Metabolites

COOH

OHGallic acid

HO OH

FIGURE 27.20 Gallic acid is the basic structural unit ofhydrolyzable tannins.

gelatin (a protein). Two categories of tannins are nowrecognized: condensed tannins and hydrolyzable tan-nins. Condensed tannins are polymers of flavonoid unitslinked by strong carbon-carbon bonds. These bondsare not subject to hydrolysis but can be oxidized bystrong acid to release anthocyanidins. The basic struc-tural unit of hydrolyzable tannins is a sugar, usuallyglucose, with its hydroxyl groups esterified to gallic acid(Figure 27.20). Gallic acid residues are in turn joined toform an extensively cross-linked polymer.

Like other phenolics, the biological role of tanninsis not clear. Tannins do appear to deter feeding by manyanimals when tannin-free alternatives are available.This effect could be related to the astringency—a sharp,somewhat unpleasant sensation in the mouth—forwhich tannins are noted. The astringent property oftannins is a component in the flavor of many fruitsas well as drinks such as coffee, tea, and red wine.As well, tannins tend to suppress the efficiency offeed utilization, growth rate, and survivorship. Theconventional interpretation has been that tanninsreduce digestibility of dietary protein, presumably bybinding with protein in the gut. Other studies, however,have cast doubt on this interpretation, suggesting thattannins may be toxic in other, yet unknown, ways.

27.5 SECONDARY METABOLITESARE ACTIVE AGAINSTINSECTS AND DISEASE

27.5.1 SOME TERPENES ANDISOFLAVONES HAVEINSECTICIDAL ANDANTI-MICROBIAL ACTIVITY

One of the better-known and commonly used naturalinsect toxins is pyrethrin, a monoterpene ester that isproduced in the ovaries of flowers of Chrysanthemum(or, Pyrethrum) cineraiifolium (Figure 27.21). C. cinerai-ifolium is widely cultivated in Montenegro, Japan, andeastern Africa, where the unexpanded flower heads aredried and powdered. Pyrethrin is a neurotoxin thatinterferes with sodium channels in the insect nerve

CH3

C CH

COO

H3C

H3C

H3C

Pyrethrin I

R

FIGURE 27.21 Pyrethrin, a monoterpenoid ester, is acommonly used organic insecticide. Pyrethrin is isolatedfrom thedried, unexpanded flowerheads of Chrysanthe-mum cineraiifolium (Asteraceae).

membrane. Pyrethrins are popular organic insecticidesbecause they have a relatively low mammalian toxic-ity. Natural pyrethrins are also readily inactivated byoxidation and so do not persist in the environment.

Many other terpenes, while not insecticides per se,do help plants to repel invasions by insects. Many plantsrespond to insect attack by producing additional quan-tities of toxic or repellant metabolites. Unfortunately,in some cases chemical deterrents can turn against theplant. As insects evolve resistance, they are able to usethe same chemical as a host-recognition cue to helpthem locate hosts they can feed on without ill effect.

At least one group of flavones, the isoflavones,has become known for its anti-microbial activities(Figure 27.22). Isoflavones are one of several classesof chemicals of differing chemical structures, knownas phytoalexins, which help to limit the spread ofbacterial and fungal infections in plants. Phytoalexinsare generally absent or present in very low concentra-tions, but are rapidly synthesized following invasion bybacterial and fungal pathogens.

The details of phytoalexin metabolism are not yetclear. Apparently a variety of small polysaccharides,glycoproteins and proteins of fungal or bacterial origin,serve as elicitors (L. elicere, to entice) that stimulatethe plant to begin synthesis of phytoalexins. Studieswith soybean cells infected with the fungus Phytophthoraindicate that the fungal elicitors trigger transcription

O

OH

HO

A C

B

O

2

3

FIGURE 27.22 Structure of the isoflavone formononetin,isolated from red clover (Trifolium pratense). In a flavone,the B ring is attached to the 2 position of the C ring. Inan isoflavone, the B ring is attached to the 3 position ofthe C ring.

27.5 Secondary Metabolites are Active Against Insects and Disease 475

of mRNA for enzymes involved in the synthesis ofisoflavones. The production of phytoalexins appears tobe a common defense mechanism. Isoflavones are thepredominant phytoalexin in the family Leguminoseae,but other families such as Solanaceae, appear to useterpene derivatives.

27.5.2 RECOGNIZING POTENTIALPATHOGENS

As discussed earlier in Chapter 13, plants challengedby insects or potentially pathogenic organisms alsorespond with a hypersensitive reaction, characterizedby changes in the composition and properties of the cellwall and other factors, including necrotic lesions at thesite of the invasion, that limit the spread of the invadingpathogen. Because the hypersensitive reaction is a formof developmental response, we must assume that a signaldetection and transduction chain is involved.

Attempts to explain the susceptibility of plants toinfection have shown that disease has an underlyinggenetic basis. In fact both pathogens and potential hostplants carry genes that determine whether or not diseasewill occur and disease will occur only when those genesare compatible. Compatibility can be explained by thegene-for-gene model, which predicts that resistancewill occur only when the pathogenic microorganismcarries a dominant allele at the avirulence (Avr) locusand host plants carry a complementary dominant alleleat the resistance (R) locus (Figure 27.23). Any othercombination of dominant and recessive alleles leads toa successful infection. A matching pair of dominantpathogen avirulence genes and dominant plant resis-tance genes initiates a hypersensitive reaction in theplant.

Although a number of avirulence genes have beenisolated from both bacteria and fungi, the specific

incompatible compatible

Pat

hoge

n ge

noty

pe

Host plant genotype

no disease disease

R

Avr/R Avr/r

Avr

avr

r

compatible compatible

disease disease

avr/R avr/r

FIGURE 27.23 The gene-for-gene model predicts thatincompatibility between a pathogen and host requirescomplementary dominant avirulence and resistancegenes in the pathogen and host, respectively. All othergenotypes are compatible and lead to invasion of the hostby the pathogen or, i.e., disease.

function of their products is not known. One pos-sibility is that avirulence genes encode enzymes forthe production of elicitors and resistance genes encodereceptors that recognize elicitors. A variety of elicitorshave been identified, most of them extracellular micro-bial products commonly associated with cell walls ofbacteria and fungi. For example, fungal elicitors includeβ-glucans, chitosan (a chitin subunit),1 and arachi-donic acid (an unsaturated lipid). Other elicitors includevarious polysaccharides, glycoproteins, and small pep-tides. Even pectic fragments, resulting from initialdegradation of the plant cell wall pectins, or mechan-ical damage are capable of eliciting a hypersensitivereaction.

Recognition of elicitors by the plant cell likelytakes place at the plasma membrane. It is expected thatsome form of signal transduction pathway is requiredto relay this information to the nucleus in order toeffect gene transcription. A variety of common signal-ing agents have been suggested, including changes inpH, and ion fluxes (especially potassium and calcium).For example, a transient uptake of Ca2+ (and effluxof K+) was observed when cultured cells were chal-lenged with a fungal elicitor. Moreover, expression ofdefense response genes can be regulated by regulat-ing intracellular Ca2+ levels. Thus, defense responsescan be activated by stimulating Ca2+ uptake with Ca2+ionophores or inhibited by blocking Ca2+ channels.Other early events in elicitor-treated cells include pro-tein phosphorylation and the production of active oxy-gen species (O−

2 and H2O2), known as the oxidativeburst. The precise role of these various signals and howthey interact in the signal cascade is unknown. It isa topic that is under active investigation in numerouslaboratories.

27.5.3 SALICYLIC ACID, A SHIKIMICACID DERIVATIVE, TRIGGERSSYSTEMIC ACQUIREDRESISTANCE

In many cases, the hypersensitive reaction leads to ageneral immunity against infection known as systemicacquired resistance or SAR (Chapter 13). Most of theevidence suggests that the signal for SAR is salicylicacid (SA) or its volatile methylated derivative methyl-salicylate. The role of SA in SAR is indicated by severallines of evidence. For example, SAR is characterized bya rise in the levels of SA along with the activation ofdefense-related genes and the appearance of their prod-ucts, pathogenesis-related (PR) proteins. In addition, SA

1The principal carbohydrate in most fungal cell walls is chitinrather than cellulose. Chitin is a polymer of N-acetylgluco-samine that forms microfibrils similar to cellulose.

476 Chapter 27 / Secondary Metabolites

is synthesized from chorismate via isochorismate, a reac-tion catalyzed by the enzyme, but Arabidopsis mutantsthat failed to accumulate SA after SAR induction wereshown to be deficient of this enzyme. Accumulation ofSA in infected plants carrying mutations of the isocho-rismate synthase gene is no more than 5 to 10 percentof wildtype plants and systemic resistance is severelycompromised.

The molecular genetics and biochemistry of SARis slowly yielding to investigation. A central player isthe regulatory protein NPR1 (NON EXPRESSOROF PATHOGENESIS-RELATED GENES1). NPR1is expressed at low levels throughout the plant, but itsmessenger RNA levels rise some two- to threefold fol-lowing pathogen infection or salicylic acid treatment.In addition, several Arabidopsis mutants that were notresponsive to salicylic acid were found to have muta-tions in the NPR1 gene. NPR1 protein interacts with agroup of transcription factors known as the TGA fam-ily of transcription factors; an interaction that requiressalicylic acid and stimulates the transcription of PRgenes.

Of particular interest is the recent discovery thatchanges in the redox status of the cell play an impor-tant role in the NPR1/TGA interaction. A changein the cellular redox potential, from an initial burstof reactive oxygen species to a more reducing envi-ronment, is induced by salicylic acid and appears tofacilitate the transcription of PR genes by triggering thetranslocation of NPR1 from the cytosol into the nucleuswhere it is free to interact with the TGA transcriptionfactors.

27.6 JASMONATES ARE LINKEDTO UBIQUITIN-RELATEDPROTEIN DEGRADATION

As previously shown in Chapter 13, a plant’s resistanceto insects and disease is also mediated by the fatty acidderivative jasmonic acid and its methyl ester (methyl-jasmonate), collectively referred to as jasmonates (SeeFigure 13.16). Jasmonates were first recognized for theirability to promote senescence of detached barley leafsegments, but a role in disease resistance was suggestedwhen phytoalexin biosynthesis in cell cultures was linkedto jasmonic acid content. It is now known that jasmonicacid accumulates in wounded plants and in plants treatedwith many elicitors. Jasmonic acid has also been linkedto the activation of a number of genes encoding proteinswith antifungal properties.

Jasmonic acid is synthesized by the oxidation andrearrangement of linolenic acid, an unsaturated fatty acid(see Appendix). Plant membranes are a rich source oflinolenic acid as a constituent fatty acid of phospholipids.It is thought that elicitors first bind with an unknown

receptor in the plasma membrane. The elicitor-receptorcomplex activates a membrane-bound phospholipasethat releases linolenic acid. The formation of jasmonicacid then involves several steps in which the linolenicacid undergoes successive oxidizations and cyclization.

Central to jasmonate signaling is a family oftranscriptional repressors (JAZ) and an F-box protein(COI1) that is structurally and functionally related tothe TIR1 auxin receptor (Chapter 18). Because of thissimilarity, the current model for jasmonate signalingbears a close resemblance to that of auxin-mediateddegradation of AUX/IAA transcriptional repressorspreviously described in Chapter 18. According to thismodel, jasmonates promote binding of JAZ repressorproteins to the E3 ubiquitin ligase (SCFCOI1) andtheir subsequent degradation via the ubiquitin-26Sproteasome system, thus allowing for the activation ofjasmonate-responsive genes.

27.7 ALKALOIDS

27.7.1 ALKALOIDS ARE A LARGE FAMILYOF CHEMICALLY UNRELATEDMOLECULES

As a group, alkaloids share three principal characters:they are soluble in water, they possess at least onenitrogen atom, and they exhibit high biological activity.Often the nitrogen will accept a proton, which gives ita slightly basic, or alkaline, character in solution (hencethe name alkaloid). Alkaloids are for the most part het-erocyclic, although a few aliphatic (noncyclic) nitrogencompounds, such as mescaline and colchicine, are con-sidered alkaloids. Altogether, some 12,000 alkaloids havebeen found to occur in approximately 20 percent of thespecies of flowering plants, mostly herbaceous dicots.

27.7.2 ALKALOIDS ARE NOTEDPRIMARILY FOR THEIRPHARMACOLOGICALPROPERTIES AND MEDICALAPPLICATIONS

The word ‘‘alkaloid’’ is virtually synonymous with theword ‘‘drug’’; as recently as 1985, 10 of the 12 com-mercially most important plant-derived drugs werealkaloids. Alkaloids generate varying degrees of phys-iological and psychological response in humans, mostoften by interfering with neurotransmitters. In largedoses, most alkaloids are highly toxic, but in smallerdoses they may have therapeutic value.

From prehistory to the present, alkaloids oralkaloid-rich extracts have been used for a variety ofpharmacological purposes, such as muscle relaxants,tranquilizers, antitussives, pain killers, poisons, and

27.7 Alkaloids 477

mind-altering drugs. One of the oldest known is opium,an exudate obtained from the immature seed capsuleof the opium poppy (Papaver somniferon). The use ofopium mixed with wine to induce sleep and relieve painwas noted on Sumerian tablets dating back to 2500 BC.The species name somniferon was chosen by Linnaeusbecause of its sleep-inducing properties. Opium isa latex gum containing a mixture of more than 20different alkaloids, including morphine, codeine, andpapaverine (Figure 27.24). The opium poppy has beentraditionally cultivated in the Golden Crescent of theeastern Mediterranean (presently Iran, Afghanistan,and Pakistan) and in parts of southeast Asia. Thegenus Papaver contains 9 other species, many of themcommon garden ornamentals, but none of whichproduce alkaloids of medical interest. In 399 BC, theGreek philosopher Socrates was executed by consumingan extract of hemlock (Conium spp.) containing thealkaloid coniine. In the Western Hemisphere, theAztecs and other native cultures used the peyote cactus(Lophophora williamsii), containing mescaline and alarge number of other alkaloids, for its hallucinogenicproperties.

Alkaloids are generally classified on the basis ofthe predominant ring system present in the molecule(Figure 27.25). In spite of the extensive variation instructure, however, alkaloids are generated from a lim-ited number of simple precursors. Most alkaloids aresynthesized from a few common amino acids (tyrosine,tryptophan, ornithine or argenine, and lysine). Thetobacco alkaloid nicotine is synthesized from nicotinicacid and caffeine is a purine derivative.

Although a few alkaloids are found in several generaor even families, most species display their own unique,genetically determined pattern. As with other secondarymetabolites, individual alkaloids may be restricted toparticular organs, such as roots, leaves, or young fruit.

Codeine

H3CO

NCH3

HO

O

Heroin

OCOH3C

NCH3

O

Morphine

NCH3

HO

HO

O

OCOH3C

FIGURE 27.24 Codeine, morphine, and heroin are structurally related alkaloids.Codeine and morphine are naturally occurring alkaloids isolated from the seed cap-sules of the opium poppy (Papaver somniferon). Heroin is a semisynthetic alkaloidproduced by acetylation of morphine. Codeine is commonly used as a cough-suppressant and local anesthetic. Morphine is used primarily as an analgesic or painkiller. Heroin was originally synthesized by pharmaceutical chemists in the late nine-teenth century as part of an effort to find a more effective alternative to morphine.

Berberine, however, is found in several sources, includ-ing seeds of barberry (Berberis vulgaris). Barberry wasat one time a common horticultural hedge but is nowout of fashion due to the high incidence of poisoning inyoung children attracted to its bright-red berries.

The quinolizidine alkaloids, such as lupinine, arefrequently called lupine alkaloids because of their highabundance in the genus Lupinus. Although range ani-mals are deterred from eating lupines because of theirbitter taste, grazing on lupines is still a common causeof poisoning of grazing cattle. The highest concen-tration of alkaloids occurs in the seed, so livestocklosses are generally highest in the fall. Other alkaloidsthat cause poisoning of livestock include senecionine(Senecio, groundsel), lycotonine (Delphinium, larkspur),scopolamine (Datura stramonium, jimson weed), andatropine (also known as hyoscyamine) from blackhenbane (Hyoscyamus niger). Agricultural crops are notwithout some risk to human consumption. The familySolanaceae is noted for genera such as Datura, Hyoscya-mus, and Atropa belladonna (the deadly nightshades),all containing high amounts of toxic alkaloids. Ediblemembers of the Solanaceae include potato (Solanumtuberosum) and tomato (Lycopersicum esculentum), whichproduce the steroidal alkaloids α-solanine and tomatine,respectively. The solanine alkaloids are cholinesteraseinhibitors that interfere with nerve transmission. For-tunately, the alkaloid content of both potato tubers andripe tomato fruit is well below toxic levels, althoughgreen vines have higher levels and are potentially toxicif eaten. Potato tubers that have been exposed to strongsunlight and have begun to green may also synthesizetoxic levels of the α-solanine and should not be eaten.

The indole alkaloids are often referred to asterpenoid-alkaloids because, although the basic indolering structure is derived from tryptophan, the restof the molecule is derived from the mevalonic acid

O

O

H3CO

H3CO

H3C

OCH3

OCH3

OCOCH3

H3CO

H3C

CH3

CH3

OH

OH

CH3

HO

COOCH3

HO

N

N

N

N

N

N

N

N NCH3 O C

O

N

N

N

N

N

N

N

N

N

N

N

N

O

CH

CH CH2

OO

H2COH

CH3

CH3

CH3

H

N

N

N

N

Alkaloid Class Example Other Representatives

Quinoline

Isoquinoline

Indole

Pyrrolizidene

Quinolizidine

Tropane

Piperidine

Purine

quinine

papaverine

vindoline

senecionine

lupinine

atropine

nicotine

caffeine

morphinecodeineberberine

vinblastinereserpinestrychnine

retrorsine

cytisine

scopolaminecocaine

coniine

O

H

FIGURE 27.25 Alkaloids are a chemically diverse group of heterocyclic,nitrogen-containing compounds that are classified according to the predominanttype of ring structure.

478

Summary 479

pathway. Two well-known terpenoid-indole alkaloids,vinblastine and vincristine, are produced by the Mada-gascar periwinkle, Catharanthus roseus. In the 1950s itwas discovered that vinblastine and vincristine arrestcell division in metaphase by inhibiting microtubuleformation. They have since been used in the treatmentof Hodgkin’s lymphoma, leukemia, and other formsof cancer. Unfortunately, C. roseus produces a complexarray of indole alkaloids of which vinblastine and vin-cristine represent only a very small proportion (about0.05% of leaf dry weight). This makes their extractionand purification difficult and costly. Recent efforts haveled to the development of cell culture systems in order toproduce higher yields at reasonable cost. Unfortunately,because alkaloid production is so often tissue-specific,production in cell culture has not been highly success-ful. Other well-known alkaloids such as cocaine (fromleaves of the coca plant, Erythroxylum coca), codeine(from the opium poppy, Papaver somniferum), nicotine(tobacco), and caffeine (coffee beans and tea leaves) arewidely used as stimulants or sedatives.

27.7.3 LIKE MANY OTHER SECONDARYMETABOLITES, ALKALOIDSSERVE AS PREFORMEDCHEMICAL DEFENSEMOLECULES

Whether alkaloids have any specific function in plantshas been a subject of debate for many years. Someplants appear to invest a significant proportion of theirresources, especially nitrogen, in a diverse array of alka-loids and yet they have no obvious function in thephysiology of the plants themselves. Like other sec-ondary products, however, there are at least two generalarguments supporting a defensive role. Firstly, mostalkaloids have a bitter taste, which is considered a uni-versally repellant character for all animals, includinginsects. Secondly, all alkaloids are biologically activeand many are significantly toxic to insects and otheranimals. Nicotine, for example, is a potent insect poisonand one of the first insecticides used by humans.

Many alkaloids have antibiotic properties and mayhave a role in defense against microbial infection. Inter-estingly, it has been known for a long time that alkaloidconcentrations are very low when measured on a wholeplant basis, but, as noted earlier, may be very high inselected organs or tissues. Often the tissues that accumu-late alkaloids are those most vulnerable in terms of plantfitness (young influorescences or seed pods, for example)or peripheral tissues that would be first attacked by her-bivores. Thus the morphine content of whole youngpoppy capsules is less than 2 percent, but 25 percentor more in the latex that exudes when the capsule iswounded. Quinine accumulates in the outer bark of the

tree Cinchona officinalis and Rauwolfia alkaloids are con-centrated primarily in the root bark. Although alkaloidshave been studied extensively for their pharmacologicaland medicinal value, there is much yet to be learnedwith respect to their physiology and chemical ecology.

SUMMARY

Although the products of plant metabolism are oftendesignated as either primary or secondary metabolites,the distinction between the two is not easily made.Primary metabolites such as protein, lipid, carbohy-drate, and nucleic acids comprise the basic metabolicmachinery of all cells. Others, such as chlorophyll andlignin, are more restricted in occurrence but are equallyessential to the growth and development of the organ-ism. Secondary metabolites, on the other hand, may befound only in specific tissues or at particular stages ofdevelopment and have no obvious role in the develop-ment or survival of the organism.

Three metabolic pathways that give rise to bothprimary and secondary metabolites are the mevalonicacid and MEP pathways and the shikimic acid pathway.The mevalonic acid and MEP pathways give rise totwo 5-carbon compounds, isopentenyl pyrophosphateand dimethylallyl pyrophosphate, that form the basisfor the terpenoid family. Terpenoids include primarymetabolites such as phytol (a portion of the chlorophyllmolecule), membrane sterols, carotenoid pigments, andthe hormones gibberellin and abscisic acid. Secondaryproducts include a range of 5-carbon hemiterpenoidsthrough 40-carbon tetraterpenes and the polyterpenes,rubber and gutta.

The shikimic acid pathway produces thearomatic amino acids phenylalanine, tyrosine, andtryptophan—all primary metabolites required forprotein synthesis. Deamination of phenylalanine tocinnamic acid, catalyzed by the enzyme phenylalanineammonia lyase (PAL), effectively diverts carbon fromprimary metabolism into the synthesis of a wide rangeof secondary metabolites—coumarins, lignin, tannins,flavonoids, and isoflavonoids—based on simplephenolic acids.

Salicylic acid is involved in establishing the hyper-sensitive reaction to insect herbivory and pathogeninfection. The gene-for-gene model predicts that noinfection will occur when the pathogen and hostplant carry dominant avirulence and response genes,respectively. An early event in this sensing/signalingpathway is the activation of defense-related genes andsynthesis of their products, pathogenesis-related (PR)proteins. The plant immune response continues withthe development of a general immune response knownas systemic acquired resistance (SAR). SAR signalingincludes the protein NPR1, which responds to a change

480 Chapter 27 / Secondary Metabolites

in cellular redox potential by migrating into the nucleuswhere it interacts with transcription factors to stimulatethe transcription of PR genes.

Other interesting and useful secondary metabolitesinclude the saponins (terpene glycosides), cardiac gly-cosides, cyanogenic glycosides, and glucosinolates. Thealkaloids are a heterogeneous group of nitrogenouscompounds with significant pharmacological prop-erties. Although the physiological roles of secondarymetabolites are poorly understood, most are toxic tosome degree and appear to serve primarily in defenseagainst microbial infection and attack by herbivores.

CHAPTER REVIEW

1. Distinguish between primary and secondarymetabolism.

2. What are terpenes? How do they originate withinthe plant and what functions do they serve?

3. Explain why sterols may be consideredboth primary and secondary metabolites.

4. What do saponins, cardenolides, and amygdalinhave in common?

5. Why are amino acids such as phenylalanineand tryptophan essential in the human diet?

6. What are EPSPS and PAL? What key metabolicroles do they play?

7. Alkaloids are, for the most part, chemicallyunrelated. What is the basis for grouping themtogether?

8. How does the herbicide glyphosate kill plants?When plants are treated with glyphosate,

it often takes a week before any signs ofinjury are observed. Explain this delay.

9. In what ways do plants have some control overinsect herbivory?

10. How do plants respond to invasion by microbialpathogens?

FURTHER READING

Cordell, G. (ed.). 1997. The Alkaloids. San Diego: AcademicPress.

Durrant, W. E., X. Dong. 2004. Systemic acquired resistance.Annual Review of Phytopathology 42:185–209.

Fobert, P. R., C. Despres. 2005. Redox control of systemicacquired resistance. Current Opinion in Plant Biology8:373–382.

Grant, M., C. Lamb. 2006. Systemic immunity. Current Opin-ion in Plant Biology 9:414–420.

Harborne, J. B., F. A. Tomas-Barberan (eds.). 1991. EcologicalChemistry and Biochemistry of Plant Terpenoids. Oxford:Clarendon Press.

Klein, R. M. 1987. The Green World: An Introduction to Plantsand People. New York: Harper & Row.

O’Kennedy, R., R. D. Thornes (eds.). 1997. Coumarins: Biol-ogy, Application, and Mode of Action. Chichester: Wiley.

Rosenthal, G. A., M. R. Berenbaum. 1991. Herbivores: TheirInteractions with Secondary Metabolites. Vol. 1: The Chemi-cal Participants. 2nd ed. San Diego: Academic Press.

Staswick, P. E. 2008. JAZing up jasmonate signaling. Trendsin Plant Sciences 13:66–71.

Kinase

AppendixBuilding Blocks: Lipids, Proteins,

and Carbohydrates

I.1 LIPIDS

Lipids (Gr. lipos, fat) are a chemically diverse group ofmolecules defined primarily by their ability to dissolvereadily in organic solvents, but only sparingly in water.These include fats, oils, and the phospholipids andglycolipids that make up the cellular membranes. Sterols(steroid alcohols) and molecules containing long-chainhydrocarbons such as the pigments chlorophyll andcarotene, although chemically distinct from fats andoils, are also considered lipids on the basis of theirsolubility properties.

Fats and oils are composed of long hydrocarbonchains called fatty acids and the three-carbon alcoholglycerol (Figure I.1). Fatty acids are attached to theglycerol through an ester link between the carboxylgroup of the fatty acid and a hydroxyl group on the glyc-erol molecule. Because one fatty acid (an acyl group)is esterified to each of the three glycerol carbons, amolecule of fat or oil is known as triacylglycerol, or atriglyceride. Fatty acids vary according to the lengthof the hydrocarbon chain (i.e., the number of carbonatoms) as well as the number of carbon–carbon dou-ble bonds and the position of those double bonds in

the chain (Table I.1). Fatty acids that have no doublebonds are known as saturated fatty acids, referring tothe fact that all the carbon–carbon bonds are saturatedwith hydrogen atoms. Fatty acids with double bondsare considered unsaturated. Unsaturated fatty acids areidentified not only by the number of double bonds, butalso by their location in the molecule. For example,there are two forms of linolenic acid: α-linolenic acidhas double bonds at the 9, 12, and 15 carbons, whilein γ -linolenic the double bonds are at the 6, 9, and 12carbons. Most fatty acids contain an even number of car-bon atoms because their synthesis involves the successiveaddition of two-carbon acetate (CH3—COO−) units.

The difference between fats and oils is a matterof melting points. The melting points of fatty acidsincrease with chain length and the extent of saturation.Saturated fatty acids tend to be solid at room temper-ature and unsaturated fatty acids tend to be liquid atroom temperature. Thus fats, which are normally solidat room temperature, are triglycerides composed of pre-dominantly saturated fatty acids, while oils, normallyfluid at room temperature, are composed of predom-inantly unsaturated fatty acids. Plant cells, especiallythose in seeds, contain mostly oils that are often storedin lipid bodies called oleosomes. Lipids may, in fact,

481

482 Appendix / Building Blocks: Lipids, Proteins, and Carbohydrates

H2C OH

HC OH

H2C OH

C (CH2)n

O

HO CH3

O (CH2)n

(—3H2O)

O

CH2C

H2C

CH3

O (CH2)n

O

CHC CH3

O (CH2)n

O

C CH3

Triglyceride

Fatty Acid (�3)Glycerol

FIGURE I.1 A triglyceride is formed by ester linkagesbetween three long-chain fatty acids and glycerol. Thevalue of n is normally 14 to 18.

be the dominant form of stored carbon in smaller seedsbecause it is an efficient form in which to store energy.Lipids are less oxidized than carbohydrate and thereforewill yield more energy when oxidized during respiration.Lipid bodies are also commonly found in chloroplasts,which contain an extensive internal membrane system.These lipid bodies may represent a reservoir of lipidsfor membrane synthesis.

The principal class of lipids found in most cellularmembranes is the phospholipids. Phospholipids arediglycerides; meaning that only two fatty acids (ratherthan three) are esterified to the glycerol molecule.

The third position is occupied instead by a phosphategroup (Figure I.2). The simplest phospholipid isphosphatidic acid (PA), in which R = −OH. PA isfound only in small quantities in most membranes.In most membrane phospholipids, R is a small, polarmolecule such as choline, ethanolamine, serine, ormyo-inositol. Phospholipids containing these headgroups are known as phosphatidylcholine (PC),phosphatidylethanolamine (PE), phosphatidylserine(PS), and phosphatidylinositol (PI), respectively.

Further variation in phospholipids is introducedby the nature of the two fatty acids; in plant mem-branes, the most abundant are 16:0, 16:1, 18:0, 18:1,18:2, and 18:3. A high proportion of unsaturated fattyacids in the membrane lipids contributes to the fluid-ity of membranes. This is because the carbon–carbondouble bond introduces a ‘‘kink’’ in the fatty acid (seeFigure I.2) that prevents the resulting phospholipidsfrom packing as tightly as they do with only saturatedfatty acids. Phospholipids are often described as havinga charged, polar phosphate ‘‘head’’ and a long hydro-carbon ‘‘tail’’ represented by the two fatty acids. Thisgives the molecule a dual character in that the phosphatehead is hydrophilic (‘‘water-loving’’) and the fatty acid tailis hydrophobic (‘‘water-fearing’’). A molecule with bothhydrophilic and hydrophobic properties is known asamphipathic.

Some plant membranes contain large amounts ofglycolipids, lipids in which the head group containsone or more sugar residues. The internal membranes ofthe chloroplast are particularly distinctive in this regard.They contain only about 10 percent phospholipidand about 80 percent monogalactosyl diglyceride(MGDG) and digalactosyl diglyceride (DGDG).MGDG and DGDG are similar to phospholipidsexcept that the phosphate group is replaced with

TABLE I.1 Some Common Biological Fatty Acids.

Common Name Symbol1 Structure mp (◦C)2

Saturated fatty acidsLauric acid 12:0 CH3(CH2)10COOH 44Myristic acid 14:0 CH3(CH2)12COOH 52Palmitic acid 16:0 CH3(CH2)14COOH 63Stearic acid 18:0 CH3(CH2)16COOH 69Unsaturated fatty acidsPalmitoleic acid 16:1 CH3(CH2)5CH CH(CH2)7COOH −0.5Oleic acid 18:1 CH3(CH2)7CH CH(CH2)7COOH 13Linoleic acid 18:2 CH3(CH2)4(CH CHCH2)2(CH2)6COOH −9α-Linolenic acid 18:3 CH3CH2(CH CHCH2)3(CH2)6COOH −17

1The number before the colon indicates the number of carbon atoms in the chain. The number following the colon indicatesthe number of double bonds.2mp = melting point.

I.2 Proteins 483

C O O

O

H2CCH2

CH2

H2CCH2

H2CCH2

H2CCH2

H2CCH2

H2CCH2

H2CCH2

H2CCH2

H3C

C

O

H2CCH2

CH

H2CCH2

H2CCH2

H2CCH

CH2

CH2

CH2

CH2

HC

H2C

H2C

H2C

H3C

O

O

O O

CH2

P X

–

FIGURE I.2 A phospholipid is a triglyceride with aphosphate group in place of one fatty acid. WhenR = OH (hydrogen), the molecule is known as phos-phatidic acid. More commonly, R will represent anothersmall, polar molecule such as choline or ethanolamine.

one or two molecules, respectively, of the sugargalactose. The remaining 10 percent of the chloroplastmembrane lipid is accounted for by sulfolipid; in thiscase, a sulfur group is attached to the galactose inMGDG.

The principal sites of a fatty acid and lipid biosyn-thesis are the endoplasmic reticulum, the chloroplast,and the mitochondria. Chloroplasts appear to have theenzymatic machinery for synthesizing all the C16 andC18 fatty acids and the addition of galactosyl residues tothe diglycerides in order to make MGDG and DGDG.Chloroplasts do not, however, appear to make phospho-lipids; this is accomplished in the endoplasmic reticulum,which contains all the enzymes necessary for the synthe-sis of PC and PE. Mitochondria are apparently unableto synthesize either PC or PE, even though these arethe principal phospholipids of the mitochondrial mem-branes.

I.2 PROTEINS

Proteins are composed of amino acids. There are 20‘‘standard’’ amino acids, specified by the genetic code,that make up all proteins. All amino acids have thesame core structure, in which a single carbon atom(the α-carbon) carries both an amino group (—NH2)and a carboxylic acid group (—COOH) (Figure I.3).At physiological pH, both the amino group and thecarboxyl group of an amino acid are ionized. Aminoacids thus carry both a negative charge and a positivecharge at the same time. Such molecules are known aszwitterions.

The structures of the 20 individual amino acids varyaccording to the nature of the R group, or side chain.This side chain may consist of nonpolar groups whichgive the amino acid hydrophobic properties, or eitherneutral or charged polar groups, which give the aminoacid hydrophilic properties (Figure I.4).

A peptide bond forms when the amino group of oneamino acid forms a covalent link with the carboxyl groupof a second amino acid (Figure I.5). When two aminoacids are linked by a single peptide bond, the result-ing molecule is called a dipeptide. Three amino acidsconstitute a tripeptide, and so forth. A chain containinga large number of amino acids is called a polypep-tide. Most proteins are polypeptides containing in therange of 40 to 4,000 amino acids, which allows for anextremely large number of different protein molecules.For example, a simple dipeptide may have any one of20 choices for the first amino acid paired with anyone of the same 20 choices for the second amino acid.This allows for 202, or 400, possible unique dipep-tides. Similarly, for a protein containing 100 aminoacids, there are 20100, or 1.27 × 10130, possible uniquecombinations.

Because the chemical properties of the side chains ofindividual amino acids are each different, it is clear thatorganisms are able to synthesize an enormous numberof different proteins with an equally large range of bio-chemical properties. The variety of proteins is furtherenhanced with the addition of nonstandard amino acids,or amino acid derivatives. These amino acid deriva-tives are the result of specific modifications made to astandard amino acid after the peptide chain has been

CH COOH3N

R+ _

FIGURE I.3 All amino acids share the same basic struc-ture. Here the amino acid is represented in the zwitte-rionic form, which predominates at physiological pH.There are 20 different side chains (R) that make up the20 standard amino acids found in all proteins.

484 Appendix / Building Blocks: Lipids, Proteins, and Carbohydrates

CH

CH2

COOH3N+ _

O

C

NH2

Asparagine

CH

CH2

COOH3N+ _

_O

CO

CH2

Glutamic acid

CH COOH3N+ _

CH2

Phenylalanine

CH2 COOH3N+ _

CH

CH

CH3H3C

COOH3N+ _

Valine

1. Glycine, the simplest amino acid:

2. Nonpolar side chain:

3. Uncharged polar side chain:

4. Charged polar side chain:

5. Cyclic (aromatic) side chain:

FIGURE I.4 Examples of amino acids representingthe four principal groups of side chains. Glycine(R = hydrogen) is the simplest of the nonpolar sidechains. Phenylalanine and others with cyclic side chainsare known as aromatic amino acids.

synthesized. One example is hydroxyproline, a non-standard amino acid that is found in certain cell wallproteins.

Protein molecules have several levels of organiza-tion, as shown in Figure I.6. The sequence of amino acidsin a polypeptide is referred to as the primary structure.Primary structure is determined by the sequence ofnucleotides in the deoxyribonucleic acid (DNA) thatmakes up the gene for that protein. This sequence isfirst transcribed into a messenger ribonucleic acid(mRNA), which migrates from the nucleus into thecytoplasm where it attaches to particles called ribo-somes. Amino acids are sequentially assembled intopolypeptides on the ribosome according to the sequenceof nucleotides, or message, in the mRNA. When the

+H3N

H2O

CH CO H

HO_

N+ CH CO

O_

+ H

R1 R2

+H3N CH C

O

N CH C

H

O

O_

R1 R2

FIGURE I.5 A peptide bond is formed by the conden-sation of two amino acids with the elimination of a watermolecule. Note that the dipeptide formed in this examplehas both an amino end and a carboxyl end. Peptides ofany length, including proteins, always have an amino (N)terminus and a carboxyl (C) terminus.

completed polypeptide chain is released from the ribo-some, it folds spontaneously to form a three-dimensionalshape. The backbone of peptide bonds may form a heli-cal coil (an α-helix), or may fold such that segmentsof the chain lie side by side to form a pleated sheet,or it may form random coils. These specific spatialarrangements are referred to as secondary structureand all three may occur within a single peptide chain.The number and distribution of helices, pleated sheets,and random coils in the polypeptide determine thethree-dimensional configuration of the entire polypep-tide, called tertiary structure. Tertiary structure of theprotein is commonly referred to as conformation. Someprotein molecules are composed of a single polypep-tide, while others are composed of multiple polypeptidechains, called subunits. The arrangement of subunitsin the protein molecule is referred to as quaternarystructure. The three-dimensional structure of a pro-tein is stabilized by a variety of noncovalent bonds,including hydrogen bonds and electrostatic interactionsbetween ionized amino and carboxyl groups on theamino acid side chains. At the tertiary level, covalentdisulfide bonds (—S—S—) may form between nearbyamino acids with sulfur groups in their side chains.Alternatively, neighboring amino acids with nonpolarside groups may form strong hydrophobic interac-tions.

Subtle differences in the composition and structuregive each protein its unique characteristics and its abilityto discriminate between other molecules with which itinteracts. In spite of the large size and complex structureof most proteins, a change in even a single amino acidin the sequence may dramatically alter its biologicalproperties.

I.3 Carbohydrates 485

A.C C C

R

N

R

C C CN

N

R

C.

D.

B.

FIGURE I.6 The structural hierarchy in proteins.(A) Primary structure refers to the amino acid sequencein a peptide chain. (B) Secondary structure. The aminoacid chain spontaneously forms an α-helix, a β-sheet,or a random coil. All three configurations may occurwithin a single peptide chain. (C) Tertiary structure isthe three-dimensional shape, or conformation, of a com-plete, folded peptide chain. (D) Quaternary structurerefers to an assembly of multiple peptide chains.

I.3 CARBOHYDRATES

Carbohydrates contain carbon, hydrogen, and oxygenin the general proportions of 1:2:1. In other words,the hydrogen (H) and oxygen (O) are found in thesame proportions as they are found in water, hence‘‘carbon hydrate.’’ The composition of carbohydrates is(CH2O)n, where n ≥ 3. Smaller carbohydrate moleculesplay a central role in the energy metabolism of cells andare the principal source of carbon skeletons for almostall other organic molecules.

I.3.1 MONOSACCHARIDES

The simplest carbohydrates are known as monosac-charides (L. saccharum, sugar), because they cannot behydrolyzed to form simpler carbohydrates. The basic

CHO

HCOH

2COHH

C O

2COHH

2COHH

Glyceraldehyde DihydroxyacetoneFIGURE I.7 Glyceraldehyde and dihydroxyacetone are thetwo simplest carbohydrates. Glyceraldehyde is an aldosugar, or aldose. Dihydroxyacetone is a keto sugar, orketose.

features of monosaccharides are illustrated by the triose(three-carbon) sugars glyceraldehyde and dihydroxy-acetone (Figure I.7). Note that both molecules containa carbonyl oxygen (—C = O). In glyceraldehyde thecarbonyl oxygen forms an aldehyde group (—CHO),while in dihydroxyacetone it forms a ketone group(—C—CO—C—). Glyceraldehyde is therefore referredto as an aldo sugar, or aldose, and dihydroxyacetone isreferred to as a keto sugar, or ketose. Other commonmonosaccharides include the 4-carbon sugar, erythrose(a tetrose); the 5-carbon sugar, ribose (a pentose); the6-carbon sugar, glucose (a hexose); and the 7-carbonsugar, sedoheptulose (a heptose) (Figure I.8).

The four most common hexoses found in plantsare D-glucose, D-mannose, D-galactose, and D-fructose(Figure I.9). Mannose is similar to glucose except forthe orientation of the hydroxyl group on the secondcarbon. Similarly, galactose and glucose differ only inthe orientation of the hydroxyl group on carbon 4.These are examples of stereoisomers. Stereoisomerismarises because the four bonds of a carbon atom do notlie in a single plane as they are usually represented onpaper, but form a three-dimensional tetrahedron:

When the carbon atom is attached to four differentsubstituents, there are two possible configurations thatare mirror images of each other. Such carbons areknown as asymmetric carbons. Glucose, for example,

CH

HCOH

O

2

HCOH

COHH

CH

HCOH

O

2

HCOH

HCOH

COHH

CH

HCOH

HCOH

O

HCOH

2COHH

HCOH

HCOH

HCOH

C O

HOCHHOCH

2COHH

2COHH

D-Erythrose D-Ribose D-Glucose D-SedoheptuloseFIGURE I.8 Examples of common monosaccharides with4-, 5-, 6-, and 7-carbon atoms.

486 Appendix / Building Blocks: Lipids, Proteins, and Carbohydrates

CHO

HOCH

HCOH

HCOHHCOH

HOCH

2COHH

CHO

HCOH

HCOH

HOCH

HOCH

2COHH

CHO

HCOH

HCOH

HOCH

2COHH

HCOH

HCOH

C O

HOCH

2COHH

2COHH

D-Glucose D-Mannose D-Galactose D-FructoseFIGURE I.9 Example of four common hexoses found inplants. Note that the three aldo sugars, glucose, man-nose, and galactose, differ only with respect to the ori-entation of the hydroxyl groups on the second, third,fourth, and fifth asymmetric carbon atoms.

contains four asymmetric carbon atoms (carbons 2, 3, 4,5), which allows for up to 24 or16 possible sterioisomers.Mannose and galactose are just two of the 16 possiblesterioisomers of glucose. Each stereoisomer has differentchemical, physical, and biological properties.

Note that the names of the four sugars above containthe prefix D- (fr. L, dextra, right hand). This identifies thestructure of the D-enantiomer, which, in solution, rotatesplane polarized light to the right, or clockwise, fromthe point of view of the observer. By convention, theD-enantiomer is drawn with the hydroxyl group on thehighest-numbered asymmetric carbon placed to the rightof the carbon chain. If this hydroxyl is written to the left(and all other hydroxyl groups are similarly inverted), thedesignation is L- (fr. L, laevus, left). L-sugars rotate planepolarized light counterclockwise, or to the left. Mostnaturally occurring sugars are in the D configuration.One exception is L-galactose, a constituent of agar.

Although monosaccharides are often depicted aslinear molecules, in solution the aldehyde or ketonegroup normally reacts intramolecularly with a hydroxylgroup to form a cyclic molecule (Figure I.10). Notethat ring closure generates an additional asymmetriccenter at the former carbonyl carbon (C1 for glucoseand C2 for fructose). Depending on the position ofthe hydroxy group on this carbon, the configuration isknown as either α or β. These two forms are referredto as anomers, and the asymmetric carbon is referredto as the anomeric carbon. When glucose moleculesare linked together in a chain, the distinction betweenα and β is not trivial. For example, starch, a storagecarbohydrate, is a polymer of α-D-glucose units whilethe cellulose found in plant cell walls is a polymer ofβ-D-glucose units. The physical and chemical propertiesof starch and cellulose are very different!

The aldehyde or ketone group of monosaccharidesmay be reduced to an alcohol group (Figure I.11).Reduction of glucose, for example, yields sorbitol andmannose yields mannitol. The sugar alcohols (sorbitol

D-Fructose

D-Glucose

Pyran Furan

β-D-Glucopyranose

α-D-Fructofuranose

α-D-Glucopyranose

O

H

HOHOH

CH2OH

HOH C2HOH C2 CH2OH

H

HOH

OHHO

HHO

OH

HH

H

H

OH

OHC

OH

H

H

HO

HC

O

O

C C

C

CH2OHO

H

H

H

HOH

OHHO

HO

OHHH

H

H

OH

CH2OH

CO

C C

C

O O

CH2OH

FIGURE I.10 In solution, monosaccharides cyclize to forman internal lactone ring. The ring forms between thecarbonyl carbon and a nonterminal hydroxyl group. Amonosaccharide that forms a six-membered ring (fivecarbons and one oxygen) is called a pyranose, after pyran,the simplest molecule containing such a ring. Similarly,a monosaccharide that forms a five-membered ring (fourcarbons and one oxygen) is called a furanose, after furan.Glucose, a 6-carbon aldo sugar, forms a pyranose, whilefructose, a 6-carbon keto sugar, forms a furanose.

and mannitol) frequently serve as storage carbohydratein algae and other lower plants. Alternatively, the pri-mary alcohol group of an aldose may be oxidized toform a carbonyl, or acid, group. The acid form of glu-cose is glucuronic acid and of galactose, galacturonicacid. Glucuronic and galacturonic acids are importantcomponents of noncellulosic cell wall polymers.

Monosaccharides such as glyceraldehyde and dihy-droxyacetone, are known as reducing sugars because ofthe ease with which the free aldehyde group reduces mildoxidizing agents. The standard test is the reduction ofsilver ion (Ag+) to metallic silver using Tollens’ reagent.A silver mirror lining on the surface of the reaction vesselindicates the presence of a reducing sugar.

I.3.2 POLYSACCHARIDES

Individual monosaccharides may be linked together inchains of varying length by glycosidic bond between the

I.3 Carbohydrates 487

CHO CHO

HCOH

HCOH

HCOH

HCOH

HOCH

COOH

HCOH

HCOH

HOCH

HOCH

COOH

2COHH

HCOH

HCOH

HOCH

2COHH 2COHH

2COHH

HCOH

HCOH

HOCH

HOCH

D-Sorbitol D-Mannitol D-Glucuronicacid

D-Galacturonicacid

FIGURE I.11 Examples of alcohol and acid derivativesof some common hexose sugars. Sorbitol and mannitolare important carbon storage products, especially in thelower plants. Glucuronic acid and glalacturonic acid areimportant components of noncellulosic cell wall poly-mers.

anomeric hydroxyl group of one sugar and a hydroxylgroup of a second sugar (Figure I.12). Small carbohy-drate polymers are identified by the number of sugarresidues (e.g., 2 sugars = disaccharide, 3 = trisaccha-ride, etc.) while longer chains are referred to simply aspolysaccharides (also known as glycans). The glyco-sidic bond is effectively analogous to the peptide bondin proteins.

The principal disaccharide found in higher plantsis sucrose, formed by the condensation of α-D-glucosewith α-D-fructose (Figure I.12). Note that the bond isformed between the number-1 carbon of glucose and thenumber-2 carbon of fructose. Because the bond involvesthe number-1 and number-2 carbons of α-sugars, itis designated as an α(1 → 2) link. Also, because bothreducing groups are involved in forming the bond,sucrose is not a reducing sugar.

Polysaccharides usually consist of a single type ofsugar (i.e., glucose), although polysaccharides consistingof more than one sugar are known. The two most

Glucose Fructose

Sucrose

O

O

H

H HOHO

HO

HCH2OH

CH2OH

HOCH2O

H

HOH

OH

HH

H4 1 2 5

FIGURE I.12 Sucrose, the principal disaccharide in plants,is composed of a molecule each of glucose and fructose.A glycosidic bond is formed between carbon-1 of theglucose molecule and carbon-2 of fructose (α 1 → 2 link.)

Glucose GlucoseCellulose

n

CH2OHO

H

HOH

OH

HH

CH2OHO

H

HOH

OH

HH

H HO O

Glucose GlucoseAmylose

n

O

CH2OHO

H

HOH

OH

HH

H

O

CH2OHO

H

HOH

OH

HH

H

FIGURE I.13 The structures of cellulose and amylose.Both are polymers of glucose. In cellulose, carbon-1 ofthe first β-D-glucose residue is linked to carbon-4 of thenext β-D-glucose residue (a β(1 →4) glycosidic bond).Amylose is a linear form of starch in which α-D-glucoseresidues are linked by α(1 → 4) glycosidic bonds. Ineither case, the value of n may be up to several thousand.

common polysaccharides in higher plants are starchand cellulose. Starch is made up entirely of (1 → 4)-linked and (1 → 6)-branched α-D-glucose residues(Figure I.13). Cellulose, on the other hand, is a long,unbranched chain of (1 → 4)-linked β-D-glucoseresidues (Figure I.13). Because both starch and celluloseare made of glucose, they are commonly referred to asglucans. Starch forms a helically coiled structure usedprimarily for storage of carbon and energy. Celluloseis a linear molecule that may contain 3,000 or moreglucose residues and is noted for its high degree ofstructural strength. Cellulose, which may comprise upto 80 percent of the dry weight of a plant, is the mostabundant organic substance in the world and perhapsone of the most economically important. A polymersimilar to cellulose is chitin, the principal componentof the exoskeleton of some invertebrates (crustaceans,insects, spiders) and the cell walls of some fungi. Chitinis a β(1 → 4)-linked polymer of N-acetylglucosamine,which differs chemically from cellulose only in thatthe hydroxyl group on carbon-2 of the glucose unit isreplaced by an —NHCOCH3 group.

Another important class of polysaccharides is thefructans. Fructans are polymers with varying numbersof fructose molecules added to the fructose end ofsucrose and are particularly common in grasses. Fruc-tans built up of β(2 → 6) links are known as levans,while those with β(1 → 6) links are known as inulins.Inulin is an important storage carbohydrate in Jerusalemartichoke (Helianthus tuberosum).

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Index/Glossary

AABC model, floral organ specification, 436Abiotic stress A physical (e.g., light, temperature) or

chemical insult that the environment may imposeon a plant. 224–235, 238, 287, 361

Abscisic acid (ABA), stomatal response to water deficit,230–233

Abscisic acid (ABA) A plant hormone primarilyinvolved in regulating seed development andstomatal closure at times of water stress. 135, 170,176

biosynthesis, 355–358discovery, 356embryo maturation, 358inhibition of ion loading in xylem, 55perception and signal transduction, 359–362role in seed development, 284role in water stress, 230–233, 358–360seed germination, 358

Absorption cross-section Factors that influence thecapacity of a photosystem to absorb light. 243

Absorption spectrum A plot of the absorbance oflight as a function of wavelength. 96, 97

anthocyanins, 106chlorophyll, 102chlorophyll-protein complexes, 115phycocyanin, 103phytochrome, 374

ACC oxidase, 362ACC synthase, 362Acclimation The capacity of a plant to adjust to and

survive a stress. 216, 234, 242–259low-temperature effects on membranes, 248–249low-temperature-induced changes in gene

expression, 248regulation by light, 249–253

Acclimation to water deficit, effects on shoot–rootratio, 253–254

leaf area adjustment, 253Accumulation ratio, 46Acid-growth hypothesis, 297, 314–316Action spectrum A plot of the efficiency of light in

causing a reaction as a function of wavelength. 97cryptochrome, 103flavonoid biosynthesis, 382leaf photosynthesis, 97photoperiodism, 418–419photosynthesis, 97

phototropism, 393phytochrome, 379stomatal opening, 135quantum yield of photosynthesis, 125UV-B responses, 105

Active transport The energy-dependent transport of asubstance across a membrane. 9, 122, 164, 318

nutrient uptake by roots, 8, 42, 43–52Adaptation heritable modifications in structure or

function that increase the fitness of an organism ina stressful environment. 261–273

Adenine nucleotide transporter A translocatorprotein that transports ATP out of themitochondrion for use elsewhere in the cell. 186

Adenosine phosphate-isopentenyl transferase (IPT)340, 349

Adenosine triphosphate (ATP), coupled phosphatetransfer reactions, 81–83

Adhesion, in capillary rise of water, 5Adiabatic lapse A decrease in temperature without an

exchange of heat, as in changing elevation. 456Aerenchyma, formation by programmed cell death,

287After-ripening Changes that a cold-requiring seed

undergoes during a period of low temperature.453

Agrobacterium tumifaciens, 346–354, 444Alcohol dehydrogenase (ADH), 181Aleurone An outer layer of cells that surrounds

the endosperm in cereal grains. A source ofhydrolytic enzymes that degrade starch and otherstored polymers in the endosperm duringgermination. 332, 334

Alkaloids, 476–479Allantoin, 206–207Allantoic acid, 206–207Allophycocyanin (allophycocyanobilin), 102Alternative oxidase, 187Alternative respiratory pathway, 186–188Ammonification The conversion of organic nitrogen

to ammonia by microorganisms in the process ofdecomposition. 196

Amygdalin, 466Amylopectin, 176α.-Amylase, 176, 332–333Amyloplast A leucoplast (colorless plastid) that forms

or contains starch grains. 86, 177, 402–403Amylose, 175

489

490 Index/Glossary

Amylopectin, 176Anaplerotic pathway, 190Angiosperms [Gk. angion, a vessel + sperma, seed]: A

plant whose seed is borne within a vessel, orcarpel. A flowering plant. 276

Antenna chlorophyll The light-gathering moleculesof a photosynthetic unit that absorb photons andtransfer the energy to the chlorophyll at thereaction center. 114–115

Anthocyanin biosynthesis, control by phytochrome,382

Anthocyanins, 106biosynthesis, 382

Anticlinal cell division Divisions in a planeperpendicular to the surface of the meristem. Asopposed to periclinal divisions which occur in aplane parallel with the surface. 278

Antifreeze proteins (AFPs), 256Antigibberellins See Growth retardant. 329Antiport, 50Aphids, analysis of phloem exudates, 157Apical dominance: role of cytokinin, 344Apical dominance The control of axillary bud

development by auxin and other factorsoriginating in the stem apex. 313

Apoplast The noncytoplasmic continuum in the plantbody, including cell wall space, tracheary elements,etc. 29, 36, 49, 54–55, 164–166, 232, 256, 294

Apoprotein, 100, 374Apparent (net) photosynthesis (AP), 142Apparent free space, 53Apparent rates of photosynthesis (AP), 218Aquaporins Protein channels or pores that control the

selective movement of water across membranes.13–15

Arabidopsis thaliana, use as a model system, 282Arabidopsis, de-etiolation, 387Aromatic amino acids, in phenylpropanoid

biosynthesis, 467–470Asada-Halliwell Pathway, 258ATP synthase (coupling factor) An integral

membrane protein complex responsible for thesynthesis of ATP using the energy of atransmembrane proton motive force.Mitochondrial ATP synthase (F0-F1- ATPase) andchloroplast ATP synthase (CF0-CF1) arestructurally and functionally similar. 90, 116, 122,185

ATPase, calcium-dependent, 301acid-growth response, 315

ATPase-proton pumps, 49iron uptake, 72potassium uptake, 134sugar uptake, 164

Atropine, 477Autotrophic organisms, 61

Auxin A plant hormone; its primary effect is to controlcell enlargement. 305–321

auxin response factor (ARF), 317polar transport, 317–320redistribution in gravitropism, 403–405role in phototropism, 395–396role in seed development, 284transport, 317–320ubiquitin and auxin signaling, 317–318apical dominance, 313AUX/IAA genes, 317axillary bud growth, 313commercial applications, 314conjugation and storage, 310control of cell enlargement, 311, 314–316deactivation, 311discovery, 307distribution, 306PIN genes, 320SAUR (small auxin up-regulated RNAs), 317signalling and transduction, 314–317synthesis, 309–310TIRl (Transport Inhibitor Response1), 317

Available water The water content of the soil betweenfield capacity and the permanent wiltingpercentage. 34

Avenacin A-1, 464Avirulence (Avr) locus, 475Axillary buds Buds that arise in the axil [Gk. axilla,

armpit], or the angle between a branch or leaf andthe stem. 279

cytokinin control of, 344–345

BBacteroids A form of specialized nitrogen-fixing cell.

200Beneficial elements Nutrients that are beneficial to

the growth and development of some plants, buthave not been shown to be requirements of allplants. 66–67

Betacyanins, 106Bioenergetics The application of thermodynamic laws

to the study of energy transformations in livingorganisms. 78–91

Biological clock The internal oscillator that regulatesinnate biological rhythms. 413

Biological clock, 423–428circadian time (CT), 424discovery, 422entrainment, 425free-running period, 423negative feedback loop, 430solar time, 424subjective day, 425subjective night, 425

Index/Glossary 491

temperature-compensation, 426zeitgeber, 425

Biological stress, as modulation of homeostasis, 224Biotic stress A biological insult (e.g., insects, disease)

to which a plant may be exposed. 224, 225–237Bisdesmosidic saponin, 465Blue-light, phototropism, 323Bolting A rapid elongation of the stem, especially in

plants with a rosette habit. Bolting usuallyprecedes flowering. 321–438

Boron, metabolic roles, 73Bound water, 3Boundary layer, 22Brassinolide receptors and signaling, 369Brassinosteroids A class of steroidal plant hormones.

367–369Bud dormancy, 449–451

induction by photoperiod, 450Bud scales, 449Bundle sheath, 263

CC02, uptake by leaf, 132C3 photosynthesis, 136–139C4 photosynthesis, 263–264

adaptation to water stress, 266biochemistry, 263–264ecological consequences, 265regulation, 269temperature sensitivity, 265–266transpiration ratio (TR), 266

C4 syndrome: specific anatomical, physiological, andbiochemical characteristics associated with C4species that increases their fitness in hot, dryenvironments. 263–267

Ca2+-ATPases, 51, 301CAB2::luc reporter system, 429Caffeine, 477, 478Calcium, as a signaling molecule, 301

metabolic roles, 70role in gravitropism, 402–404

Callus cultures The culture of undifferentiated planttissue. Plant tissue cultures. 276–277, 341, 346

Callus tissue [L. callos, hard skin]: A term used todescribe undifferentiated tissue in tissue cultureand wound healing. 276–277

Calvin cycle (see photosynthetic carbon reductioncycle)

Capillary rise, role in water movement, 29Carbon gain and plant productivi1y, 213–221Carbon partitioning The distribution of

photosynthetic carbon and associated energythroughout the plant. 151

Cardenolides, 465Cardiac glycosides, 465

Cardinal temperatures, 218, 448Carotenes, 103, 325, 355–357, 460–461

role in photoprotection, 244–247protecting chlorophyll from photooxidation, 103

Carrier proteins, and nutrient uptake by roots, 43,51–52

Casparian band, 28–29, 54Cavitation A process of rapid formation of bubbles in

the xylem. 31Cell cycle, 292–293

role of cytokinin, 343–344Cell division, 292–294Cell plate: An aggregate of vesicles that align along the

equatorial plane in a dividing cell. The vesicles arederived from the Golgi complex and will fuse toform the plasma membranes and middle lamella ofthe two daughter cells. 293

Cell wall, role in cell growth, 294–298structure and synthesis, 289–290

Cellular respiration, 173–193Cellulose, structure, 289–290

synthesis, 292CF0-CF1 ATPase, 90, 116, 121–122, 230Chalcone synthase (CHS) The enzyme that catalyzes

the first committed step in the synthesis offlavonoid biosynthesis. 472–473

Chaperonins: A class of proteins that direct theassembly of rnultimeric protein aggregates. 235

Chelate (Gk., chele, claw): A stable complex formedbetween a metal ion and an organic molecule. 71

Chelating agents, 71Chemical potential (μ) The free energy per mole of a

substance. Chemical potential is a measure of thecapacity of a substance to react or move. 8–11

Chemiosmosis, synthesis of ATP, 85, 90respiratory synthesis of ATP, 185photosynthetic ATP synthesis, 120–122

Chemiosmotic model for auxin transport, 317–320Chemoautotrophs Organisms that are able to convert

carbon dioxide to organic carbon by using theenergy obtained by oxidizing inorganic substances.196

Chilling sensitive plants, 233–234Chloride ion, metabolic roles, 74Chlorophyll a, 101–102

absorption spectrum, 102Chlorophyll b, 101–102

absorption spectrum, 102Chlorophyll c, 101Chlorophyll d, 101Chlorophyll-protein (CP) complexes, 70, 102, 115, 116Chloroplast, structure, 85–89Chlororespiratory pathway The capacity of

chloroplasts to reduce O2 in the dark. 258Cholodny-Went hypothesis, in phototropism, 395

in gravitropism, 403

492 Index/Glossary

Chromophore The portion of a pigment moleculeresponsible for the absorption of light. 100,374

cryptochrome, 103, 386neochrome, 397phototropin, 105, 396phycocyanin, 102phytochrome, 103, 383phytolases, 105, 386

Chromoplast A plastid containing pigments otherthan chlorophyll, usually carotenoids andxanthophylls. 86, 103

Chromoprotein A protein molecule combined with alight-absorbing moiety called the chromophore.100, 374

CIRCADIAN CLOCK ASSOCIA TED 1 (CCAl),,429–430

Circadian clock (see biological clock), geneticdissection, 429–430

Circadian rhythm [L. circa, about + dies, a day]:Biological rhythms with a periodicity ofapproximately 24 hours. 423

Citric acid cycle (CAC) The second stage of cellularrespiration in which pyruvate is completelyoxidized to CO2, protons, and electrons. TheCAC takes place in the stroma of themitochondrion. Also known as the Kreb’s cycle ortricarboxylic acid (TCA) cycle. 182–183

Clausius’s dictum, 79Climacteric An ethylene-induced burst of respiratory

carbon dioxide during the ripening of certainfruits. 364, 444

Cobalt, as plant nutrient, 67Cocaine, 479Codeine, 477, 478, 479Coenzyme A (CoA), role in citric acid cycle, 183Cohesion-tension theory, for xylem sap movement,

30–33Colchicine, 476Cold acclimation, winter cereals, 254–255Coleoptile A protective sheath that protects the leaf in

the embryo of a grass seed or cereal grain. 276,284, 286, 306–311, 315–316, 391–401

Coleorhiza A protective sheath that protects theradical in the embryo of a grass seed or cerealgrain. 286

Colligative properties, of a solution, 10Columella A group of cells located in the center of the

root cap. Columella cells are the cells that sensegravity. 401–405

Companion cells, 160Coniine, 477, 478CONSTANS (CO), role in dormancy, 451Constitutive protein A protein that is present at all

times. 207Copper, metabolic roles, 73

Corpus (The interior mass of cells in the shoot apicalmeristem. Corpus cells divide in all planes to giverise to volume of the primary plant body. 278

Cotransport, 50Cotyledon A seed leaf. The cotyledon generally

stores food in a dicotyledonous seed and absorbsfood from the endosperm in a monocotyledonousseed. 276

Coumarins A family of phenolic chemicals known fortheir anticoagulant properties as inhibitors ofvitamin K. Coumarin derivatives are usedmedically to thin the blood of patients with heartproblems. 468–470

Coupling factor, 90Crassulacean acid metabolism (CAM) A variant of

the C4 pathway; phosphoenolpyruvate carboxylasefixes C02 in C4 acids at night. During the daytime, the fixed C02 is then transferred to thechloroplast where it is fixed by the Calvin cyclewithin the same cell. Characteristic of mostsucculent plants, such as cacti. 267–270

regulation, 269Cristae Invaginations of the mitochondrial inner

membrane. 89Critical concentration, nutrient requirements, 67–68Crosstalk A term use to describe interactions between

different signaling pathways. 237, 303Crown gall, 345, 346Cryptochrome A small family of photoreceptors that

mediate blue-light responses. 103, 374, 379–380,386–388

as photoreceptor for photoperiodism, 418role in stomatal opening, 136structure, 386signal transduction, 386–387

Culture techniques, for studying plant nutrients,62–64

Cuticle, 20Cuticular transpiration, 20Cutin, 20Cyanide-resistant respiration, 187Cyanobacteria, photosynthesis, 123–124Cyanogenic glycosides, 466Cyclic electron transport Photosynthetic electron

transport through PSI independently of PSII,without the formation of NADPH. 120–122

Cyclic photophosphorylation The formation ofATP in association with cyclic electrontransport. 120

Cyclin A regulatory protein that rises and falls in apredictable pattern during the cell cycle. 293,343–344

Cyclin-dependent kinases (CDKs) Enzymes thatcontrol progression through the cell cycle. 293,343

Cytochrome b6f complex, 119

Index/Glossary 493

Cytochrome complex, in chloroplast, 9Cytokinesis The division of the cell into two daughter

cells, as opposed to karyokinesis, or nucleardivision. 292, 293–294

Cytokinin oxidase/dehydrogenase (CKX), 341in shoot apical meristem, 347–348

Cytokinins A plant hormone primarily responsible forregulating cell division and delayed senescence,among other effects. 55, 339–354

role in seed development, 283–284biosynthesis, 339, 341–343deactivation, 340discovery, 341receptors and signaling chain, 351–353role in cell cycle, 343–344

Cytoskeleton A network of microtubules andmicrofilaments that is responsible for positioningor moving organelles within the interior of thecell. 292, 295, 394, 404

DD1 repair cycle, overcoming damage to PSII. 227Daylength measurement, external coincidence model,

428photophile phase, 427–428role of photoperiodism and biological clock,

428–429scotophile phase, 427–428

De-etiolation The reversal of etiolation caused bylight. 380

DELLA proteins A class of nuclear proteins, ortranscription factors, that function as repressors ingibberellin signaling. 335

Desert biomes, 272–273Desert perennials, adaptations to reduce transpiration

and heat load, 272Designed leakage A hypothesis that attempts to

explain the role of cavitation in times of waterstress. 33

Development, defined, 275Dhurrin, 466Dicotyledons (dicots) One of two classes of flowering

plants; characterized by having seeds with twocotyledons, net-veined leaves, and flower parts infours or fives. 71, 110–111, 131, 160, 167, 256,263, 277, 281, 285, 290

Dicoumarol A coumarin common in moldy hay.470

Dielectric constant, 4Differentiation, defined, 276Diffusion coefficient, 6Digitalis, 465Digitoxin, 465Digoxin, 465Dimethylallyl pyrophosphate (DMAP), 325Dinitrogen fixation (see nitrogen fixation)

Dinitrogenase The enzyme responsible for fixingnitrogen. 74, 200

hydrogen production, 202NIF genes, 203sensitivity to oxygen, 202

Donnan equilibrium, 47Donnan potential, 47Dormancy mechanisms, 449

EEcodormancy Dormancy imposed by limitations in

the environment. 449Ectomycorrhizae, 57Electrochemical gradient, 46Electrogenic proton pump, 49–51Electromagnetic spectrum, 95Electron transport chain The third stage of cellular

respiration in which electrons derived from theoxidation of intermediates in the citric acid cycleare transferred to oxygen with the accompanyingconversion of redox energy to ATP. The electrontransport chain is based in the inner mitochondrialmembrane. 183–185

Elicitor (L. elicere, to entice): Any metabolite isolatedfrom pathogens that evoke a hypersensitive responsein host plants. 474, 475

Embolism An obstruction caused by the formation oflarge gas bubbles in the xylem. 31

Endodermis, 28Endodormancy Dormancy that is an inherent

property of the dormant structure itself. 449Endogenous rhythm [Gk. endon, within + Gk. genesis,

origin]: A rhythm that is not imposed by externalfactors but is generated within an organ ororganism and persists under constant conditions(usually constant light or constant darkness). 112,423–425

Endomycorrhizae, 57Endosymbiosis, 86Energy fluence, 983-Enolpyruvylshikimate-5-phosphate synthase

(EPSPS) A key enzyme in the synthesis ofaromatic amino acids. EPSPE is inhibited by theherbicide glyphosate. 468

Enthalpy The total heat energy of a system, includingenergy available to do work. 80

Entropy A measure of the randomness or disorder of asystem associated with the random movement ofmatter, or a measure of energy that is not availableto do work. 79

Envelope A pair of outer limiting membranes thatdelimit the chloroplast. 85

Environmental stress, 223–238Enzymes, biological catalysts, 146–149Ephemeral plants, 225

494 Index/Glossary

Epigeal germination, 285Epinasty, 391, 406Epinasty The downward curvature of leaves or other

organs, especially in response to the hormoneethylene. 364

Essential nutrient elements Nutrient elements thatare required by plants in order to successfullycomplete their life cycle. 65

Essential oils, 460Ethylene A plant hormone known primarily for its role

in the ripening of climacteric fruit. 362–367biosynthesis, 362–364discovery, 363receptors and signal transduction, 364–367role in climacteric fruits, 444triple response mutants, 364–365

Ethylenediaminetetraacetic acid (EDTA), 71Etiolation A condition of increased stem elongation,

suppressed leaf development, and lack ofchlorophyll. Etiolation is especially characteristicof seedlings grown in darkness or under low lightconditions. 380

Etioplast A specialized plastid present in etiolatedleaves. When exposed to light, etioplasts undergoa reorganization to form chloroplasts. 86

Evapotranspiration The transfer of water vapor fromvegetated land surfaces to the atmosphere,regardless of the source of the vapor. 271

Excitation pressure An energy imbalance due tooverexcitation of PSII by high light. 251

Excited states of molecules, 96Expansins Proteins that induce stress relaxation and

extension of isolated cell walls at low pH.297–298

Extensin, 291Extracellular matrix (ECM) The complex of proteins

and polysaccharides that forms cell walls andother structures external to the plasmamembrane. 289

FF0-F1 ATP-synthase, 184–185Facilitated diffusion, and nutrient uptake by roots, 42F–box protein A protein that recruits a protein

targeted for ubiquitination, the F-box proteincontains a recognition site for the targeted proteinand a recognition site (the ‘‘F-box’’) for theSCF-complex. 302, 317, 476

Feedback inhibition In biochemistry, feedbackinhibition is a mechanism to control metabolicpathways where the end product inhibits theactivity of an enzyme earlier the pathway. Asimilar situation arises in genetics when theproduct of one gene represses the transcription ofanother gene. 148, 217

Fermentation, 181–182Ferredoxin, 70–71, 83, 117, 119–122, 141, 201, 203,

208Fick’s first law of diffusion, 6, 22Field capacity The water that remains in capillary

pores of soil after free (gravity) drainage iscompleted. 34

First law of thermodynamics, 79Flavonoids, 105–106

biosynthesis, 471–473Floral meristem identity genes, 421, 434, 436Floral organ specification, ABC model, 436Floral organ-identity genes, 434, 436Floral repressors, 435Floral-identity gene, 421, 434, 436Florigen A hypothetical flower-promoting hormone.

420–421Flower The reproductive structure of an angiosperm: a

complete flower includes sepals, petals, stamens,and carpels, but all flowers contain at least onestamen or carpel. 279

FLOWERING LOCUS C (FLC), 434, 441FLOWERING LOCUS T (FT) role in dormancy,

451FLOWERING LOCUS T (FT), 434, 441Flowering, 433–442

autonomous pathway, 435effect of temperature, 437–442GA pathway, 435long-day pathway, 435

Flowering-time genes, 434Flowering-time mutants, 434Fluence The quantity of radiant energy falling on a

small sphere, divided by the cross-section of thesphere. 98

Fluorescence and dissipation of excess energy, 96Flux The amount of material crossing a unit area per

unit time. 6Free energy Energy that is available to do work. 80Freezing tolerance, 255–257Fructans, biosynthesis, 156, 487Fruit ripening, in climacteric fruits, 444Fruit set: The first stage in frit development, defined

by the decision to either abort furtherdevelopment or to proceed with further celldivision and cell enlargement in the ovary walls.442–444

role of auxin, 442–444

GGain-of-function mutations, 349, 361Gene-for-gene model A model that attempts to

explain the genetic basis for the hypersensitivereaction in plants. 475

Geranylgeranyl phyrophosphate GGPP, 327

Index/Glossary 495

Gibberellins, 323–337commercial applications, 330deactivation, 329discovery, 325dwarf mutants, 330–332gene expression, 333–336nutrient mobilization, 332role in seed development, 283–284rosette plants, 331signal transduction, 333–336structure and synthesis, 323–329transport, 330

Gluconeogenesis The biosynthesis of glucose fromnoncarbohydrate precursors. 188–189, 283, 286

Glucosinolate, 310, 466Glutamate dehydrogenase (GDH), 205Glutamate synthase (GOGAT), 204Glutamine synthetase (GS), 204Glycolysis The first stage of respiratory carbon

metabolism in which hexose sugars are partiallyoxidized to pyruvic acid. Glycolysis takes place inthe cytosol. 82, 174, 178–180, 189

β-Glycosidase, 466Glyoxylate cycle, role in gluconeogenesis, 188Glyoxysome An organelle that serves as a site for the

conversion of fatty acids to carbohydrate. 188Glyphosate, 170, 468Glyscyrrhizin, 465G-protein-coupled receptor (GPCR), 299Grana Regions in the chloroplast where adjacent

thylakoids appear to be closely appressed. 87–89,122–124, 227, 244

Gravitropism A differential growth response togravity. 398–405

perception, 401–402presentation time, 400reaction time, 400response types, 399role of auxin, 403–405role of calcium, 404–405roots, 401–405starch-statolith hypothesis, 402threshold intensity, 400

Green fluorescent protein (GFP) A fluorescentprotein encoded by a gene from a jellyfish. Whenthe GFP gene is fused to a plant gene of interestand the DNA encoding the fusion protein isintroduced into plant cells, the plant protein canbe located by the fluorescence pattern in amicroscope. 385, 405

Greenhouse effect, 99Gross primary productivity (GFP) The total carbon

assimilated by photosynthesis. 214Ground state The energy state of an unexcited

molecule. 96Growth respiration, 214

Growth retardant A chemical that causes dwarfing byinterfering with gibberellin biosynthesis. 329

Growth An irreversible increase in size. 276Guard cells, 20, 130–131Gutta, 462

HHardiness, 241Heat shock proteins (HSPs), 234–235Heliotropism (solar tracking), 221Heterotrophic organisms, 61High-temperature stress, 234–235Histidine kinase domain, cytokinin receptor, 350Histidine-phosphotransfer proteins (HPTs), 353Holochrome, 100Homeobox (homeodomain) genes/proteins, 348, 451Homeostasis [Gk. homos, similar, + stasis, standing]:

The condition of a relatively stable internalphysiological environment, usually involvingextensive feedback mechanisms. 224

Hormone concept in plants, 305–306Humus The colloidal carbonaceous residue in soils. 40Hydration shells, 4Hydraulic conductance A measure of the capacity of a

tissue or membrane to conduct water; the inverseof resistance to water movement. 32

Hydroactive closure, stomata, 230–233Hydrogen bonding, water, 3Hydrogenase, 75Hydrologic cycle, 271Hydropassive closure, stomata, 230–233Hydroponic culture A soil-less method of plant

culture using controlled nutrient solutions. 62Hydroxynitrile lyase, 466Hyoscyamine, 477Hypersensitive reaction A collection of responses

that serve to limit the spread of invadingpathogens. 235, 475

Hypertonic solution [Gk. hyper, above, over]: Asolution with a higher solute content than a cell oranother solution and, hence, more negativeosmotic potential. 12

Hypocotyl The section of the stem axis below thepoint of attachment of the cotyledons [Gk. hypo,less than or below, + cotyledon]. 285

Hypogeal germination, 286Hyponasty An abnormal bending up of organs such as

leaves and petioles. 391, 406Hypotonic solution [Gk. hypo, less than]: A solution

with a lower solute content than a cell or anothersolution and, hence, less negative osmoticpotential. 12

IInositol triphosphate, 300Imbibition, in seed germination, 282

496 Index/Glossary

Incipient plasmolysis, 12Inducible protein A protein that is present only when

the appropriate substrate is available. 207Insect and disease resistance, 235–237, 474–476Internodes, 286Invertase, 177Ion channels, 43, 44–45Ion pumps A term used in reference to active transport

systems because they move solutes against aconcentration or electrochemical gradient. 43

Iron, metabolic roles, 71–73Irradiance, 98Isoflavones A group of flavones with antimicrobial

properties. 474Isoflavonoids, 106Iso-pentenyl pyrophosphate (IPP), 325, 368Isoprene, 101, 325, 460Isothiocyanates, 70

JJasmonic acid A molecule that, along with its methyl

ester, appears to have an important role in plantdefense against insects and disease. 237, 476

KK+ transport systems, 47–49, 51–52KNOX genes, 348KNOX genes, role in shoot apical meristem, 348–350Kranz anatomy, 263, 265

LLactate dehydrogenase (LDH), 181LATE ELONGATED HYPOCOTYL (LHY), 429Latent heat flux, 271Latex, 462Laticifers, 463Leaf area adjustment A mechanism for reducing leaf

area and transpiration during times of limitedwater availability. 253

Leaf area index (LAI) The ratio of photosynthetic leafarea to covered ground area. 220

Leaf, anatomy, 220LEAFY gene, 434, 436Lectins Proteins that recognize and reversibly bind

with specific carbohydrates. Also known ashemagglutinins because they agglutinate red bloodcells by forming cross-links between sugar residueson the cell surfaces. Because of this property, thelectin ricin, from castor bean (Ricinus communis) isconsidered one of the most potent animal toxinsknown. 199

Leghemoglobin, 202, 204Lenticels, 20Leucoplasts Colorless plastids. 86Ligand The term ligand has two uses. In a strict sense,

a ligand is any molecule that shares an electron

pair with a cation through a coordinate bond, suchas a metal ion or calcium with a chelating agent. Ina more general sense, ligand is used to identifyany, usually smaller, molecule that is bound to amacromolecule. The ligand may itself be anothermacromolecule, as in case of a protein binding to aproteolytic enzyme (a protease) or a regulatorymolecule as in the case of a hormone or othersignal molecule binding to a receptor. 71–72,300

Light compensation point The fluence rate at whichthe rate of photosynthesis is sufficient to justbalance respiration. 216, 262

Light, absorption within the leaf, 110–111measurement of, 98

Light-harvesting complex (LHC), 86, 115–116, 123,244, 249, 255, 262

Phosphorylation, 243Lignin A plastic-like component of some secondary

cell walls; after cellulose, lignin is the mostabundant plant polymer. 298, 470–471

Limit dextrins, 176Lipids, oil droplets, 188, 481Low-temperature stress, 233–234Lumen The intrathylakoid space. 85, 89–90, 116–122,

141Lutein, 104, 244Lupinine, 477

MMacronutrients Nutrient elements that are required

in excess of 10 mmole kg−1 of dry weight. 65Magnesium, metabolic roles, 70Maintenance respiration, 214Maltose, 176Manganese, metabolic roles, 74, 118MAP kinase cascade, 366Maryland Mammoth tobacco, 414Matric potential (M) The contribution to water

potential by the adsorption of water to solidsurfaces. 11, 282

Matrix The unstructured interior of a mitochondrion.89

Mehler reaction, 258Membrane, selectively permeable, 7Meristem [Gk. merizein, to divide]Undifferentiated

tissue that gives rise to new cells. 277Mescaline, 477Mesophile plants, 448Metallothioneins, 287Methylerythritol-4-phosphate (MEP), 326–327Methyljasmonate, 237, 476Mevalonic acid, gibberellin biosynthesis, 326Micronutrients Nutrients that are required in

relatively small quantities (less than 10 mmolekg−1 of dry weight). 65

Index/Glossary 497

Micronutrient, toxicity of, 75Micropropagation, 345Middle lamella The layer of the extracellular matrix

that cements together the primary walls ofadjacent cells. Composed primarily of pecticsubstances. 290, 293

Mitochondria (sg. mitochondrion), structure, 89energy-transducing membranes, 83ATP synthesis, 90

Mitogen-activated Protein Kinase (MAPK), 366Molybdenum, metabolic roles, 74Monocotyledons (monocots) One of two classes of

flowering plants; characterized by seeds with asingle cotyledon, parallel-veined leaves, and flowerparts in threes. 72, 131, 263, 281, 290

Monodesmosidic saponin, 465Morphine, 477, 478Mucigel A mucilaginous coating that lubricates the

growing root tip. 56, 277Multimeric protein A protein made up of multiple

peptides or subunits. 185, 200, 235, 292, 302Mustard oils, 467Mycorrhiza, pl. mycorrrhizae A form of mutualism in

which a root is infected with a symbiotic fungus.57–58

phosphorous uptake, 69

NNADPH: Protochlorophyll oxidoreductase. 101Nastic movement A plant movement that is not

obviously related to any vector in the stimulus.405–410

Natural products: see secondary metabolitesNeochrome, 397Nernst equation, 48Net primary productivity (NPP) The net increase in

carbon, or carbon gain, after accounting for loss ofcarbon due to respiration. 214

effect of CO2 concentration, 216effect of leaf factors, 220effect of light, 215–216effect of nitrogen supply, 219effect of soil water potential, 219effect of temperature, 218

Nickel, metabolic roles, 74Nicotine, 477NIF genes, dinitrogenase, 203Nitrate reductase, 74, 207Nitrification The oxidation of ammonia to nitrate by

nitrifying bacteria. 196Nitrite reductase (NiR), 207Nitrogen assimilation, 195–211Nitrogen cycle The flow of nitrogen between the

major global nitrogen pools. 196Nitrogen cycling The simultaneous import and export

of nitrogen by an organ such as leaves. 208–209

Nitrogen fixation The process of reducing dinitrogen(N2) to ammonia. 196–200

biochemistry, 200–203energy cost, 201symbiotic, 197–200

Nitrogen, metabolic roles, 68Nitrogen-fixing bacteria, 196Nod factors Chitin derivatives

(lipo-chitooligosaccharides) that are secreted intothe soil solution by rhizobia and that prepare thehost roots for rhizobial invasion. 199

Nod genes, in relation to nodulation, 203Nodules, nitrogen-fixing, 198–200Noncyclic (or linear) electron transport The

linear or flow-through process of photosyntheticelectron transport in which electrons arecontinuously supplied from water and withdrawnas NADPH. 120–122

Noncyclic photophosphorylation The formation ofATP in association with noncyclic electrontransport. 120

Nutation A slow rotary or helical pattern described bya growing plant stem. 391

Nutrient depletion zone, 57–58Nutrient functions and deficiency symptoms,

67–75Nyctinastic (sleep) movements Periodic up and

down movements of organs such as leavesregulated by the biological clock. 406–409

OOleosomes Storage droplets of lipid or oil (also called

oil bodies, or spherosomes) that are normallyfound in cells of the cotyledons or endosperm ofseeds. 188, 481

Opines, 346Osmoregulation The process of regulating the

osmotic properties of plant cells. 15–17Osmosensor A device that is able either to detect

changes in the chemical potential of extracellularwater or osmotic properties of the cell. 15

Osmosis The property of water passing through asemipermeable membrane with the tendency ofeventually equalizing the water potential in thetwo compartments. 6–7

Osmotic adjustment A mechanism for regulatingthe water status of a plant cell by accumulatingsolutes and consequently lowering osmoticpotential. 9, 247

Osmotic potential (�s) The change in free energy orchemical potential of water produced by solutes;carries a negative (minus) sign; also called solutepotential. 10

Osmotic pressure (π ) The pressure developed by asolution separated from pure water by adifferentially permeable membrane; it is generally

498 Index/Glossary

an index of the solute concentration of thesolution. 10–11, 153, 296

Oxaloacetate (OAA), first product of C4photosynthesis, 263

β-Oxidation The degradation of fatty acid chains bytwo-carbon units, resulting in the formation ofacetyl-CoA. 188

Oxidative pentose phosphate cycle (OPPC), inchloroplast, 145

Oxidative pentose phosphate pathway, in cellularrespiration, 180

Oxidative phosphorylation The chemiosmoticsynthesis of ATP associated with the transfer ofelectrons through the electron transport chain andthe accompanying consumption of oxygen. 90,185

Oxygen-evolving complex (OEC) A cluster of fourmanganese ions associated with a small complex ofproteins that are responsible for the splitting(oxidation) of water and the consequent evolutionof molecular oxygen. 118

PPapaverine, 477Paradormancy Dormancy imposed by another part of

the plant. 449Paraquat, 124, 127Parthenocarpy (fr. Gk parthenos, virgin): The

development of fruit in the absence of pollinationand fertilization. 443

Pathogenesis-related (PR) proteins Plant proteinsthat are synthesized in response to microbialattack and that serve to limit the growthpathogens. 236, 475

Perforation plates, 26Peribacteroid membrane The membrane that

surrounds bacteroids in the host cell. 200Pericycle, 55, 285Peristomal evaporation, 20Permanent wilting percentage The soil water

content below which a plant is unable to extractsufficient water to maintain turgor. 34

Peroxidase, 471Peroxisome, role in photorespiration, 142–143,

265Phaseic acid, 357–358Phenolics, 467Phenylalanine ammonia lyase (PAL) An enzyme that

diverts aromatic amino acids from protein syn-thesis to the synthesis of phenolic molecules.468

Phenylpropanoids, 467Pheophytin The primary electron acceptor in the

reaction center of PSII. 101, 117–118Phloem [Gk. phloos, bark): The food-conducting

tissue of plants; composed of sieve elements,

various kinds of parenchyma cells and supportingtissues. 156–166

analysis of phloem exudates, 157–159loading, 163structure, 159–160translocation of xenobiotic agrochemicals, 170unloading, 166

Phosphoglycolate, 143–144Phospholipase, 300, 316, 476Phosphorous, metabolic roles, 69Photoacclimation Developmental adjustments in the

structure and function of the photosyntheticapparatus in response changes in irradiance.249–251

Photoassimilate A general term that refers tocarbon compounds; carbon assimilated byphotosynthesis. 151

distribution, 166–170partitioning, 168–170

Photoautotrophs Organisms that convert inorganiccarbon to organic carbon using energy derivedfrom light. 123, 252, 270

Photobiology The study of the effects of light onbiological organisms. 93, 98

Photoblastic seeds Seeds that respond eitherpositively or negatively to light. 379

Photochemical quenching, 228Photoequilibrium, 378Photoinhibition A light-dependent decrease in

photosynthetic rate at high irradiance, such aswhen the light exceeds the requirements for thephotosynthetic evolution of 02 or thephotosynthetic assimilation of CO2. 225–227,244, 247

Photomorphogenesis The control of plantdevelopment by light. 93, 373–388

Photon fluence, 98Photon A discrete units or particle of light. 94Photooxidation of water by PSII, 117Photoperiod, induction of bud dormancy, 450

signal perception by leaf, 419Photoperiodism A response to the timing and

duration of light and dark periods. 93,413–423

critical daylength, 417in nature, 430inductive treatment, 417response types, 415role of dark period, 417

Photophosphorylation The light-dependentchemiosmotic synthesis of ATP in the chloroplast.90, 120–122, 105, 202, 230, 270

Photoprotection The capacity of carotenoids toprotect the photosynthetic system from chronicphotoinhibition. 244–247

Photoreceptors, 100–106

Index/Glossary 499

Photorespiration (PR) A series of reactions in which02 is attached to RuBP, eventually resulting in therelease C02 from the plant. 142–144

Photoreversibility, of phytochrome, 376–377Photostasis The maintenance of cellular energy

balance, which is dependent uponchloroplast-mitochondrial interactions. 251

Photosynthesis [Gk. photos, light, + syn, together +tithenai, to place]: The conversion of light energyto chemical energy; the production ofcarbohydrates from carbon dioxide and water byusing light energy in the presence of chlorophyll.93, 109–149

acclimation to light quality, 252and phototropism, 393–394as oxidation-reduction process, 112discovery, 113energy conversion by, 109–122in cyanobacteria, 123inhibition by light, 225–227inhibitors of electron transport, 124, 127light compensation point, 136sensitivity to water stress, 229–230

Photosynthetic capacity The balance betweencarboxylation capacity and electron transportcapacity. 217

Photosynthetic carbon reduction (PCR) cycle Thepathway by which all photosynthetic eukaryoticorganisms ultimately incorporate CO2 intocarbohydrate. Also known as the Calvin Cycle,after M. Calvin, who was largely responsible forunraveling the reactions of the cycle. 136–142

carboxylation reaction, 137energetics, 139reduction of 3-PGA, 138regenerations reactions, 138–139regulation, 139

Photosynthetic control: The regulation of the rate ofphotosynthetic electron transport by thetransthylakoid �pH. 155

Photosynthetic electron transport chain, 114–119inhibitors, 124, 127

Photosynthetic induction time, 140Photosynthetically active radiation (PAR) That

portion of the electromagnetic spectrum between400 nm and 700 nm. 98

Photosystem I (PSI), 114–117Photosystem II (PSII), 114–117Phototropin The photoreceptor for phototropism.

103, 396–398signal transduction chain, 397structure, 396

Phototropism Differential growth of a plant within alight gradient, e.g., toward or away form a lightsource. 392–398

as a blue-light response, 393

as a response to light gradient, 392auxin redistribution, 395–396Cholodny-Went hypothesis, 395fluence response curves, 394–395in green plants, 398optimizing photosynthesis, 393photoreceptor, 396–397signal chain, 397–398

Phragmoplast A cluster of interdigitatingmicrotubules oriented perpendicular to the planeof the new crosswall during cell division. 293–294

Phycobilins, 102Phycobilisomes, 102Phycocyanin (phycocyanobilin), 102, 383Phycoerythrin (phycoerythrobilin), 102Physiological ecotypes, 431Phytoalexins: Chemicals of a variety of chemical

classes that help to limit the spread of bacterial andfungal infections in plants. 106, 474

Phytochrome A family of photoreceptors that mediatered and far-red light responses. 103, 374–389

as photoreceptor for photoperiodism, 418chromophore, 103, 383discovery, 375high irradiance reactions (HIRs), 378input to circadian clock, 429–430low fluence responses (LFRs), 378photoconversion, 378, 383signal transduction, 384–386structure, 383very low fluence responses (VLFRs), 378

Phytochromobilin The chromophore of thephotomorphogenic chromoprotein phytochrome.383

Phytoecdysones Plant sterols that have a structuresimilar to insect molting hormones. Wheningested by insect herbivores, phytoecdysonesdisrupt the insect’s molting cycle. 462

Phytol, 100Phytosiderophores Iron-binding ligands. 72Phytotropins Inhibitors of auxin transport. 319PIN proteins, in auxin transport, 320

in gravitropism, 405Pistil The central female organ of a flower. Typically

consists of a stigma, style, and basal ovary. 279Pit pairs, 26Plant development, control by temperature, 447–454Plasmodesma, pl. plasmodesmata [Gk. plasma,

form + desma, bond]: Cytoplasmic strands thatextend through cell walls, connecting theprotoplasts of adjacent cells to form theinterconnected cytoplasmic unit referred to as thesymplast. 26, 294

Plasmolysis A condition when the protoplast thenshrinks away from the cell wall, resulting in a lossof turgor. 12

500 Index/Glossary

Plastocyanin, 73, 119–122, 184Plastoquinone, 84, 117–122, 124Plastoquinol, 17Plumule [L. plumule, a small feather]: The first

bud of the embryo or the epicotyl of a dicotseedling. 285

Polar transport The preferential transport of auxin ineither the basipetal (in shoots) or acropetal (inroots) direction. 317–320

Pollen, structure and germination, 280Pollination, 280Polygalacturonase (PG), role in fruit ripening,

442Polymer trap A model to account for symplastic

phloem loading. 165Polyphenols, 467–474Porins Integral proteins in the chloroplast and

mitochondrial outer membranes that serve aslarge, nonselective channels. 89

Porphyrin, 100Potassium , metabolic functions, 69Pressure-flow hypothesis, in phloem translocation,

161–163Primary auxin responsive genes, 317Primary growth Growth, predominantly in length,

that arises from apical meristems. 277–279Primary wall The cell layer deposited during periods

of active cell expansion. 289Primordia, in meristems, 278Programmed cell death (PCD), 236, 287Prolamellar body (PLB) A highly ordered,

paracrystalline structure found inetioplasts. 86

Proplastid A small, self-reproducing vesicularorganelle from which plastids develop. 86

Proteasome A multiprotein protease complex thatdegrade proteins. 302

role in auxin signaling, 317Protein kinase cascade, 300Proteoid roots, 56Protochlorophyll a, 101Protochlorophyllide The colorless chlorophyll

precursor. 86Proton motive force (PMF) An electrochemical

gradient across a membrane, established by thecombination of membrane potential differenceand proton gradient. 49–90

Proton pumps, 90Protoplast, 7, 12, 27, 44–45, 57, 133, 136, 159, 164,

230, 287–289, 294, 296–297, 333Pseudocarpic fruits, 442Psychrophiles, 448Pulvinus, 406–407Pyrethrin A natural insecticide derived from

Chrysanthemum cineraiifolillm flowers. 474Pyruvate dehydrogenase, 183

QQ10Quantum (pI. Quanta): The energy carried by a

photon. 94Quiescent zone A zone of relatively inactive cells at

the center of the root apical meristem. 277Quinine, 479

RRadiationless transfer and dissipation of excess energy,

97Rain forest biomes, 270Raoult’s law, 21Reaction center The chlorophyll plus associated

proteins and redox carriers that are directlyinvolved in light-driven redox reactions. 86, 102,114–118, 175, 218, 227, 228–229, 230, 234,244–247

Reactive oxygen species (ROS) Toxic forms ofoxygen, such as free radicals and peroxides, whichcan lead to cell damage. 244

Redox couple A reduced/oxidized pair such asNADPH and NADP+. 83

Redox potential The tendency to accept or donateelectrons. 83

Reducing sugars Reducing sugars have a free aldehydegroup that reduces mild oxidizing agents. Thestandard test is the reduction of silver ion (Ag+) toyield a metallic silver mirror lining on the inside ofthe reaction vessel. 56, 158–159, 486

Resistance (R) locus, 475Respiration A series of metabolic reactions for

retrieving the energy and carbon stored byphotosynthesis. 173–194

alternative pathway, 186–188as a source of carbon skeletons for biosynthesis,

189–191ATP synthesis, 185–186citric acid cycle, 182–183cyanide-resistant, 187electron transport chain, 183–185glycolysis, 178–180impact of temperature on, 248–249pentose phosphate pathway, 180–181response to environmental conditions, 192–193role of mitochondrion, 182–188SHAM-sensitive, 187

Respiratory climacteric, 191, 364, 442, 444Respiratory quotient (RQ), 174Retrograde gene regulation The regulation of

nuclear genes by organelles such as thechloroplast. 251

Rhizobia: A group of nitrogen fixing bacteria. 197Rhizosphere, 56Rhythmic leaf movements, 422–424Riboflavin, 84, 103, 105, 311, 379, 393

Index/Glossary 501

Ribulose-l,5-bisphosphate carboxylase/oxygenase(Rubisco), 88, 133, 138–145, 148, 202, 209–210,217, 219, 225–226, 234–235, 249–250, 254–255,262–267, 429

Rieske iron-sulfur (FeS) protein Iron-bindingproteins in which the iron complexes with sulfurresidues rather than a heme group as in the case ofthe cytochromes. 71, 119–121

Root apical meristem (RAM), 277, 281, 285, 401Root cap, 275

role in gravitropism, 401–405Roots, and water uptake, 34–37

role in nutrient uptake, 42–59response to gravity, 401–405

Rosette plants, control by gibberellin, 330–332Rotenone-insensitive dehydrogenase, 186Rubber, 462Rubisco activase, 141, 234Rubisco An acronym for the enzyme

ribulose-l,5-bisphosphate carboxylase/oxygenase.88, 133, 138–145, 148, 202, 209–210, 217, 219,225–226, 234–235, 249–250, 254–257, 262–267,429.

SS-adenlosylmethionine (SAM), 362–363Salicylhydroxamic acid (SHAM), 187Salicylic acid, 236, 475Sapogenins, 464Saponins, 464Saturation vapor pressure, 21Scarification The process of mechanically disrupting

or removing the seed coat. 284, 452SCF complex The multiprotein enzyme 3 (E3)

complex that mediates ubiquitination of proteinstargeted for degradation by the proteasome.302

role in auxin signaling, 317Scopolamine, 477Second law of thermodynamics, 79Second messenger A mobile signaling molecule that

is formed or released inside the cell in response tothe binding of a first messenger—e.g., a hormoneor other ligand—to a receptor or the activation ofa receptor by an environmental signal. 300

Secondary growth, 279Secondary metabolites Molecules that may have no

obvious role in normal cell function. 459–479Secondary wall The highly organized innermost layers

of cell walls that are laid down after the cell hasstopped expanding. 289, 298

Seed coat, as dormancy mechanism, 451Seed development, 279–281, 283–284Seed dormancy, 451–454

impact of temperature, 453role of inhibitors, 452

Seed germination, 281–285Seismonasty, 406, 409–410Selenium, as plant nutrient element, 67Senecionine, 477Senescence associated genes (SAGs), 286Senescence, 286

delay by cytokinins, 346Shade avoidance syndrome A condition of increased

stem elongation, reduced branching, and earlyflowering characteristic of plants growing inshade. 381

Shikimic acid pathway, in aromatic amino acidbiosynthesis, 468, 470, 472, 475

in flavonoid biosynthesis, 472Shoot apical meristem (SAM), 24, 277–279, 285, 420,

429, 433, 435maintenance by cytokinins, 347–348role of gibberellins, 349

Sieve element, 159–170Sieve-tube members, 159Silicon, as plant nutrient, 67Singlet state, 96Sodium, as plant nutrient, 66Soil colloids, 40Soil, composition and structure, 33–34, 40–42Soil, ion exchange capacity, 41Soil-plant-atmosphere continuum The integrated

flow of water from the soil, through the plant, andinto the atmosphere. 33

Solute potential (see osmotic potential)Solution culture, 62Spectral energy distribution (SED) A measure of

light quality. 98Spring cereals, temperature and photoperiod, 438Stamen The male organ of a flower. The site of pollen

production in a flower. 279Starch phosphorylase, 177Starch, biosynthesis, 152, 154–156

mobilization (breakdown), 175–178Statocysts, 402Statocytes, 402Statoliths, 402Stele The central vascular core of a root. 285Stem elongation, gibberellin, 330–332Steroids, 462Sterols, 462Stilbenes A phenylpropanoid derivative that has

antifungal properties. 472Stoma, pl. stomata [Gk. stoma, mouth]: A pore in the

epidermis of leaves and stems. Also used to refer tothe entire stomatal apparatus which includes thesurrounding guard cells. 130–131

hydroactive closure, 231hydropassive closure, 231mechanism of opening and closure, 133–136regulation by endogenous rhythm, 136

502 Index/Glossary

response to water deficit, 230–233structure, 20

Stomatal complex, 130Stomatal conductance, 232Stomatal transpiration, 20, 130Stratification The process of enhancing seed

germination by subjecting moist seed to lowtemperature. 453

Stress, acclimation, 223–238high-temperature, 234–235low temperature, 233–234monitoring, 228–229role of oxygen 258

Stroma The unstructured background matrix in achloroplast. 85–90

Suberin, 28, 54Substomatal space, 20Substrate-level phosphorylation The direct synthesis

of ATP through the transfer of a phosphate groupfrom a substrate to ADP. 183

Sucrose carrier genes (SUTI , SUC2), 165Sucrose, biosynthesis, 153–156Sulfur, metabolic roles, 70Sun and shade species, 262Sunflecks, 100, 243Superoxide dismutase (SOD), 258Superoxide radical (O−

2 ), 258Surface tension, 4Symbiotic nitrogen-fixation, 197–200Symplast Protoplast continuity through

plasmodesmata. 164–166, 294Symport, 50Systemic acquired resistance: The development of a

general immune capacity throughout the entireplant following an initial invasion by a pathogen.236, 475

TTemperature and flowering response 437–442

breaking dormancy, 453distribution of plants, 454–457role in defining biomes. 454

Temperature stress, acclimation to high temperatures,257

cold acclimation, 255–257Terpenes A diverse family of molecules built on the

5-carbon isoprene unit. 325–327, 460–465synthesis, 325–327

The photorespiratory glycolate pathway, 143Thermal deactivation and dissipation of excess energy,

96Thermodynamics, 78–80Thermogenesis, 187Thermonasty Plant movements, e.g., opening and

closure of petals, due to differential growth inresponse to alternating temperature. 406, 454

Thermoperiodism The growth response of plants toalternating high and low temperature. 454

Thermophiles, 448Thermotolerant plants, 257Thigmonasty, 406Thiocyanates, 70Thioglucosides (thio, sulfur), 467Thioredoxin, 141, 145, 155Thylakoids, lateral heterogeneity of photosynthetic

complexes, 122–123Thylakoids A highly structured internal system of

membranes in the chloroplast. 88Ti plasmid The tumor-inducing plasmid of

Agrobacterium tumifaciens. 345TIRl (Transport Inhibitor Response1) 317toc1 (timing of CAB) 429–430Totipotency The concept that cells are capable of

assuming an embryonic state and developing intoany tissue of a mature plant. 276

Trace elements Another term for micronutrients.Nutrients that are required only in trace amounts.65

Tracheary elements, 24flow rates, 27

Transcription factor A protein that binds to DNAand regulated transcription of the gene. 303

Transfer cells Specialized parenchyma cell with wallin-growths that increase the surface area of theplasma membrane; apparently functioning in theshort-distance transfer of solutes. 160

Translocation The transport of photoassirnilates overlong distances. 151

Transpiration ratio (TR) The ratio the uptake of C02to the loss of water by evaporation (transpiration)from the leaf. 266

Transpiration The loss of water vapor by plants; mosttranspiration occurs through open stomata. 20–27

effect of relative humidity (RH), 23effect of temperature, 23effect of wind, 24quantity, 19

Triplet oxygen, 244Triplet state, of an excited molecule, 97Tropism A differential growth response that is related

to the direction of the stimulus. 391True (gross) photosynthesis (GP), 142Tunica One or more peripheral layers of the shoot

apical meristem. Tunica cells give rise to surfacegrowth by anticlinal divisions. 278

Turgor pressure The pressure component that arisesfrom the force exerted outwardly against the cellwalls by the expanding protoplast. 12

role in cell expansion, 296–297role in calcium channels, 301

Two-component systems Sensory signalingmechanisms consisting of a transmembrane sensor

Index/Glossary 503

protein and a cytoplasmic protein called theresponse regulator. 15–17, 351–353, 384

cytokinin receptor, 350ethylene receptor, 365osmosensing receptor, 15phytochrome, 384

UUbiquitin A small protein that is linked to other

proteins targeted for degradation by proteasomes.302–303

auxin signaling, 317cryptochrome signaling, 387gibberellin signaling, 335in cell cycle, 293in gravitropism response, 405jasmonate signaling, 476phytochrome degradation, 377role in heat stress, 235senescence-associated gene, 286

Ultraviolet radiation, 94, 100Uniport system, 50Urease, 66, 75UV-B radiation, as a developmental signal, 105

VVapor density, 21Vapor pressure, 21Vascular cambium, 279Vernalin: A hypothetical plant hormone thought to be

involved in the vernalization response. 440Vernalization (L. ver, plus a suffix meaning ‘‘to make’’

or ‘‘become.’’): The promotion of floweringbrought about by subjecting the shoot apex to aperiod of low temperature. 438–442

induced state, 439perception by shoot apex, 440

Vesicular-arbuscular mycorrhiza (VAM), 57Vessel members, 26Vessels, structure, 26–27Vinblastine, 479Violaxanthin, in ABA synthesis, 356Violaxanthin, role in photoprotection, 244Vivipary Precocious germination, i.e., while the seed is

still on the mother plant. 358Volatile organic carbon (VOC), 463

WWall pressure The inward pressure exerted by the cell

wall against an expanding protoplast; equal butopposite to turgor pressure. 12

Water potential (�) The algebraic sum of the solutepotential and the pressure potential, or wallpressure; the chemical potential of water in acell. 11–13

Water potential gradient, as the driving force for watermovement, 11

Water stress, 229–230Water-use efficiency (WUE), 266Water, chemical potential, 8–11

cohesion, 4heat of fusion, 3heat of vaporization, 4hydrogen bonding, 3movement by bulk flow, 5physical and chemical properties, 2–5solvent properties, 4specific heat, 3surface tension, 4tensile strength, 5thermal properties, 3

Winter cereals, freezing tolerance, 254–255temperature and photoperiod, 438tolerance to photoinhibition, 254

XXanthophyll cycle, role in photoprotection, 228–229,

244Xanthophylls, 86, 103–104, 244, 355–357

in LHCII, 116Xanthoxin, 356Xylem vessels, the ascent of sap, 27Xylem, cohesion-tension theory, 30–33Xyloglycans, in cell wall, 290, 292

YYang cycle, 362–363Yield threshold, 296

ZZeaxanthin, 104, 244Zinc, metabolic roles, 73Zygote, 280