TEACHING TOOLS IN PLANT BIOLOGY™: LECTURE NOTES

Light-Dependent Reactions of Photosynthesis

INTRODUCTION AND OVERVIEW

Photosynthesis in plants converts the energy of sunlight into

chemical energy and through this process releases O2. Although

photosynthesis involvesmanyproteinsandcatalyticprocesses, it

often is described as two sets of reactions, the light-dependent

reactions and the carbon-fixing reactions, which are also occa-

sionally described as the “dark reactions.”

Plants do not have a monopoly on photosynthesis, which also

occurs in other photosynthetic eukaryotes and several species

of bacteria. In most cases, bacterial photosynthesis does not

release oxygen. Oxygenic (O2-producing) photosynthesis is

restricted to cyanobacteria, green, red, and brown algae, and

plants. Oxygen is produced when water is split to release

protons (H1), electrons, and oxygen. Oxygenic photosynthesis

requires the involvement two photosystems working in series,

photosystem I (PSI) and photosystem II (PSII).

The light-dependent reactions of photosynthesis in plants can

bedescribed as a series of steps that take place in and across the

thylakoid membranes of chloroplasts. Light harvesting involves

the capture of photonsbypigmentmolecules and the funneling of

light energy to reaction center chlorophyll molecules. Reaction

center chlorophylls are positioned so that the captured light

energy induces a charge separation across the membrane. Light

excitation of the reaction center chlorophyll of PSII causes it to

give up electrons and become oxidized. Electrons stripped from

water with the accompanying production of oxygen reduce the

oxidized reaction center chlorophyll in PSII. Electrons are ulti-

mately passed by way of the electron transport chain of cyto-

chrome b6f and through PSI to NADP1, reducing it to NADPH.

During these reactions, protons are translocated to the interior

lumen compartment of the thylakoid membranes. The resulting

charge and pH gradients, collectively known as proton motive

force, drive the synthesis of ATP by the ATP synthase complex.

This lesson examines how the multiprotein complexes that

make up the photosynthetic apparatus carry out these steps,

their evolutionary origins, and how their inherent flexibility and

feedback loops enable photosynthetic reactions to acclimate to

changing light andmetabolic conditions. Finally, we explore how

current insights are enabling scientists to develop improved algal

strains for biofuels and plants that are more resilient to abiotic

stresses and are inspiring the design of artificial systems for the

conversion of sunlight into energy.

Our understanding of photosynthesis rests on the work of

thousands of scientists working in physics, engineering, and

chemistry and across the disciplines of biology. Pioneers include

JosephPriestley, who recognized the ability of plants to “restore”

air that had been “injured” by a burning candle (in other words,

to produce oxygen). Key discoveries came from the work of

Willstatter (Nobel Prize in Chemistry, 1915), Warburg (Nobel

Prize in Physiology or Medicine, 1931), Hill (1930s), Emerson

(1950s), Arnon (1950s–1970s), Calvin (Nobel Prize in Chemistry,

1961), Woodward (Nobel Prize in Chemistry in 1965 for the total

synthesis of chlorophyll and other molecules), Deisenhofer,

Huber, and Michel (Nobel Prize in Chemistry in 1988 for the

determination of the three-dimensional structure of a bacterial

photosynthetic reaction center), and others too numerous to

mention. For comprehensive histories of the pioneers of pho-

tosynthetic research, seeGovindjee andGest (2002), Govindjee

et al. (2003), and Govindjee et al. (2004).

EVOLUTION AND DIVERSITY OF PHOTOSYNTHESIS

It is difficult to set a date for the beginning of photosynthesis.

We knowwith certainty that there is a layer of insoluble iron that

was deposited in rocks ;2.3 to 2.5 billion years ago, which

resulted from a dramatic increase in the level of atmospheric

oxygenand theconcomitantdecrease in solubility of irondue to

its oxidationbyoxygen to the formof iron(III) oxide. This sets the

most recent date for the evolution of oxygenic photosynthesis,

but evidence from stromatolites (ancient fossils) suggest that

oxygen-producingcyanobacteriamayhave evolved longbefore

this, perhaps as long as 3.4 billion years ago. The time lag

between the potential origin of oxygenic photosynthesis and

the accumulation of atmospheric oxygen that occurred millions

of years later can be attributed largely to the ability of oceans to

accumulate and buffer oxygen. Suffice it to say, photosynthesis

has beenmaking a tremendous contribution to the atmospheric

gas composition and the entry of energy into Earth’s biosphere

for a long time.

Although photosynthetic prokaryotes are found in several dif-

ferentphyla, there arebasically threedifferent typesofchlorophyll-

based photosynthesis. Anoxygenic photosynthetic prokaryotes

use either a Type I or a Type II reaction center (RC), described

further below. As the name indicates, these organisms do not

oxidizewaterbut insteaduseanelectrondonorother thanwater.

Oxygenic photosynthesis, which is restricted to cyanobacteria

and the eukaryotic organisms that have cyanobacteria-like

chloroplasts, requires both Type I and Type II reaction centers

working in series to bridge the energy gap between water

oxidation and NADP1 reduction.

The evolution of photosynthesis has been described as,

“a complex process that cannot be described by a simple, linear,www.plantcell.org/cgi/doi/10.1105/tpc.15.tt1115

The Plant Cell, November 2015, www.plantcell.org ã 2015 American Society of Plant Biologists. All rights reserved.

branching evolutionary diagram.Rather, photosynthesis emerged

by recruiting and modifying genes encoding components of

several other pre-existing metabolic pathways, along with a

few key innovations and probably several lateral gene-transfer

events. The resulting view is that, likemanymetabolic pathways,

photosynthesis is a mosaic process that has no single well

defined evolutionary origin,” (Blankenship, 2001).

Anoxygenic Photosynthesis

Photosynthetic bacteria have been found in six phyla, of which

five contain only anoxygenic photosynthetic bacteria. The wide-

spread occurrence of photosynthesis and the fact that only some

members of these phyla are photosynthetic lends support to the

idea that lateral gene transfer (the direct movement of genes

across species, not via a common ancestor) has been a major

factor in the distribution of photosynthetic capacity.

Anoxygenic photosynthetic prokaryotes use either a Type I or

Type II reaction center (and it is likely that Type II reaction centers

are derived from Type I). Both types share the fundamental

property of using light energy to induce charge separation

across a membrane using bacteriochlorophyll as the primary

electron donor. They differ in the first stable electron acceptor,

which in Type I RCs are Fe4-S4 clusters and in type II RCs are

quinone molecules.

When only one reaction center is present, photosynthetic

electron transfer is cyclic and the light-driven reaction mainly

promotes the establishment of a proton motive force for the

production of ATP.

Oxygenic Photosynthesis

Cyanobacteria

Cyanobacteria are a monophyletic group but are also among

the most morphologically diverse and successful prokaryotes.

They are important contributors to Earth’s oxygen supply and

the conversion of solar energy to chemical energy. Many cya-

nobacteria are also able to fix nitrogen (convert chemically inert

N2 into NH41, a form that can be used by organisms). Most

cyanobacteria are free-living organisms in aquatic ecosystems.

Some cyanobacteria livewithin plant tissues as endophytes and

can contribute to the plant’s nitrogen nutrition, and some form

symbiotic associations with fungi in the form of lichen or even

with marine animals (see below). Cyanobacteria provide an

outstanding model for the study of oxygenic photosynthesis.

Eukaryotic Photosynthesis: Primary and Secondary

Endosymbiosis

With a single known exception (Paulinella chromatophora;

describedbelow), photosynthetic eukaryotes aredescendants

of a primary endosymbiotic event that took place ;1 to 1.5

billion years ago, in which an ancestral eukaryotic cell engulfed

anancestral cyanobacterium.Thisprimaryendosymbioticevent

resulted in three lineages: Viridiplantae (green algae including

Chlamydomonas reinhardtii and land plants), rhodophytes (red

algae), and glaucophytes (rare freshwater algae).

In themillions of years since this primary endosymbiotic event,

many of the endosymbiont’s genes were transferred to the

nucleus, making photosynthetic function dependent on both

nuclear and organelle genomes. Many plastid proteins are

encoded by nuclear genes; their genes are transcribed in the

nucleus, their mRNAs are translated in the cytosol, and the

resulting proteins are transported into the plastid through

a two-layered envelope that resembles the cyanobacterial

inner and outer membranes. The development of machinery

for the import of proteins from cytosol to plastid was essential

for the transition from free-living cyanobacteria to plastid. The

selective advantage for genemigration from endosymbiont to

nucleusmaybe that thenuclear locationof thesegenesprovides

a better integration of their function or because nuclear genes,

unlike plastid-localized genes, can self-correct during meiosis

via recombination.

Although green algae and terrestrial plants descend from the

same ancestor, there have been some changes in their photo-

synthetic machinery since they diverged;400million years ago.

The core reaction center complexes of PSI and PSII are more or

less conserved among cyanobacteria, algae, and plants, but the

light-harvestingcomplexes (LHCs) that capturephotons toenergize

PSI and PSII differ in different photosynthetic organisms.

Finally, secondary and tertiary endosymbiosis has extended

the ability to carry out photosynthesis much further than the

progeny of the primary endosymbiotic event. Secondary and

tertiary endosymbiosis refers to the process in which green or

red algae (the products of primary endosymbiosis) were them-

selves engulfed within other cells. Products of these events

include euglenoids, diatoms, dinoflagellates, and other algae.

Collectively, these organisms belong to a broad spectrum of

eukaryotic clades, and although they are important in global

energy transfer, they are less commonly used asmodels for the

study of light-dependent photosynthetic reactions.

Nascent Primary Endosymbiosis in P. chromatophora

Recently, a second example of primary endosymbiosis was

identified in thephotosynthetic amoebaP.chromatophora,which

is in the kingdom Rhizaria. This second primary endosymbiotic

event occurred relatively recently, ;60 million years ago; recall

that the primary endosymbiotic event that gave rise to plants

occurredmore thanabillionyearsago.P.chromatophoraharbors

photosynthetic organelles called chromatophores, which resem-

ble plastids. Chromatophores in P. chromatophora are derived

from a-cyanobacteria, whereas all other known plastids in pho-

tosynthetic eukaryotes are derived from b-cyanobacteria. The

chromatophore is clearly an endosymbiotic organelle because it

cannot live independently outside of the host and it divides with

the host cell. Like plant plastids, some of the P. chromatophora

genes have been transferred to the nucleus, although to a lesser

extent as those in chloroplasts probably due to their younger

evolutionary age. As in plants and green algae, the proteins

encoded by these transferred nuclear genesmust be transported

into the organelle. This exciting discovery offers researchers

a system in which to explore an early stage of plastid evolution

and ultimately to understand better the evolution of eukaryotic

photosynthetic organelles.

2 The Plant Cell

LIGHT AND PIGMENTS

The Nature of Light

Photosynthesis converts light energy into useful biochemical

products. Light is composed of energy-carrying particles known

asphotons. Photons can alsobe represented as electromagnetic

waves that travel at a fixed speed of;3 3 108m/s (denoted as c)

with a frequency (n; cycles per second) andwavelength (l; dis-

tance between peaks) in the relationship c 5 n l. Since c is

a constant, as l decreases, n increases; the shorter the wave-

length, the higher the frequency. Furthermore, since E 5 hn

(whereE is a photon’s energy, h is Planck’s constant, and n the

photon’s frequency), as wavelength decreases, frequency

increases and so does energy. Long-wavelength light such

as that in the infrared region of the spectrum is low in energy,

whereas short-wavelength light such as that in the UV region

of the spectrum carries much more energy. Thus, the wave-

length of light determines whether or not it has enough energy

to drive the reactions of photosynthesis. In plants, PSII has an

energy requirement for light of 680 nm or shorter, and PSI

requires light of 700 nm or shorter.

Visible light is defined by the properties of the human eye and

falls between ;400 and 700 nm. Electromagnetic radiation of

wavelengths less than 400 nm include high-energyUV light and

x-rays, which are largely blocked from reaching the earth’s

surface by its atmosphere. Electromagnetic radiation of wave-

length longer than 700 includes low-energy infrared light and

radio waves.

Photosynthetic Pigments and Accessory Pigments

Pigments are compounds that absorb visible light and are often

characterized by networks of double bonds. As a consequence

of the pigments they produce, plant chloroplasts preferentially

absorb blue and red light, which is why leaves and other pho-

tosynthetic tissues of plants appear green. When a pigment

molecule’s conjugated electron system absorbs light, an elec-

tron becomes excited to a higher energy state that can be

passed on to anothermolecule. In photosynthetic systems, light

is absorbed by accessory pigments present in protein-pigment

arrays arranged in LHCs. A system of accessory pigments sur-

rounds each reaction center in what is described as an antenna

complex. Each antenna complex passes the energy of collected

light to a reaction center at its center. The size and composition of

the antenna complex is sensitive to the environment; for example,

a heavily shaded leaf has a larger antenna complex surrounding

each reactioncenter thana leaf in full sunlight.Accessorypigments

include chlorophylls, carotenoids, and phycobilins (in cyanobac-

teria and red algae).

Chlorophyll

Most chlorophyll-dependent photosynthetic organisms produce

chlorophyll a, which is always present in the reaction center,

and one or more additional forms of chlorophyll that serve as

accessory pigments. Chlorophyll comprises a cyclic tetrapyr-

role moiety with a central Mg ion and an acyl (lipid) tail through

which it is anchored. Chlorophyll is produced in plastids through

the tetrapyrrole biosynthetic pathway, which is a branched

pathway that also leads to heme, siroheme, and phytochromo-

bilin, which is a linear tetrapyrrole that is the chromophore of the

photoreceptor phytochrome. Structurally related to chlorophyll

are pheophytins that are the first electron acceptor from the

excited chlorophyll of PSII. Pheophytin is simply chlorophyll

without the Mg.

Substitutions on the tetrapyrrole ring structure affect chloro-

phyll’s absorption spectra. Chlorophyll b is found in land plants,

green algae, and cyanobacteria. Chlorophyll c lacks the phytol

tail and is primarily found in brown algae and dinoflagellates.

Chlorophyll d, e, and f are minor components and found in

diverse organisms, mostly cyanobacteria. Bacteriochlorophylls

are structurally similar to plant chlorophylls and are produced by

a conserved biosynthetic pathway. Plant chloroplasts produce

chlorophyllaandb,which show twopeaksof light absorbance in

blue and red light. Chlorophyll a absorbsmaximally at;430 and

660 nm, and chlorophyll b at ;460 and 650 nm, with peak

absorbance wavelength varying slightly with solvent. Chloro-

phyll a and b are differentially represented in core reaction

centers and antennacomplexes,so therelativeabundanceofeach

is an indicator of the proportional representation of each type of

complex.

When chlorophyll a at the reaction center absorbs light energy,

an electron moves to a higher energy level, and the pigment

is denoted as Chl*. The excited electron can fall back to its

lower energy level, releasing energy as longer wavelength

light (fluorescence), the energy can be released as heat, or the

electron can be transferred to another molecule as the first

step of photochemistry; when this happens, the resulting

chlorophyllmolecule is denoted asChl1. The rest of this article

describes what happens after the charge transfer event that

initiates photochemistry.

Carotenoids

Carotenoids are accessory pigments that absorbmainly in the

blue range and appear yellow or orange; they capture photons

that would not be absorbed by chlorophyll and pass the

energy to the reaction center chlorophyll. Carotenoids have

essential functions in photoprotection, including the dissipa-

tion of excess energy when light levels exceed the plant’s

ability to process it, and they also structurally stabilize the

photosystems.

There are two classes of carotenoids: hydrocarbon carotenes

andoxygen-containingxanthophylls.Bothare40-carbon lipophilic

pigments made from isoprenes. Carotenes such as a-carotene,

b-carotene, and lycopene appear orange; they are named after

the orange color of carrots but are also found in sweet potato,

pumpkin, and other orange-colored tissues. Xanthophylls

appear yellowish (xanthos in Greekmeans yellow) and include

lutein, zeaxanthin, and violaxanthin; the latter will be discussed

later in the context of the xanthophyll cycle, which is involved in

nonphotochemical quenching (NPQ) of excess light energy. Ca-

rotenoids are important nutrients for humansaswell;b-carotene is

the precursor for vitamin A, and lutein accumulates in the retina

November 2015 3

of the eye where it protects photopigments from damage. Carot-

enoids are abundant in photosynthetic tissues but their color is

masked by that of chlorophyll; when chlorophyll breaks down

during senescence, they are revealed as the familiar yellow and

orange colors of autumn leaves.

Phycobilins

Phycoblilins are linear tetrapyrroles that comprise the chromo-

phore moiety of phycobiliproteins, which are water-soluble

accessory photosynthetic proteins in cyanobacteria, red algae,

and other non-green algae. In cyanobacteria, phycobilins are

found in phycobilisomes, which act as light-harvesting antenna.

Thephycobilins areespecially efficient at absorbing red, orange,

yellow, and green light, wavelengths that are not well absorbed

by chlorophyll a. Small modifications to the phycobilin chromo-

phore or to their orientation relative to the apoprotein affect phy-

cobiliprotein absorption spectra. Phycobilin pigments include

phycocyanin (abundant in cyanobacteria, absorbs red and orange

light, and appears blue) and phycoerythrin (abundant in red algae,

absorbs blue and green light, and appears red). The chromophore

of phytochrome, which regulates light responses, is a linear

tetrapyrrole related to phycobilins.

THE LIGHT RESPONSE CURVE AND QUANTUM

EFFICIENCY

The light response curve is an important tool for describing the

rate of photosynthesis and is produced by plotting the rate of

photosynthesis (measured as O2 evolution or CO2 consumption)

at varying light intensities. At low light levels, photosynthesis is

limited by light, so there is a linear increase in photosynthesiswith

increasing light. However, this linear relationship only holds at

fairly low light levels. The light saturation point is the light level at

which the rate of photosynthesis is limited by factors other than

light; it depends on many factors including the species of plant,

the light intensity towhich the plant has acclimated, temperature,

etc. In thedarkor at very low light levels,mitochondrial respiration

can be observed as a net consumption of O2 (or production of

CO2). The light level at which respiration and photosynthesis

balance and net O2 production is zero is defined as the light

compensation point; it also depends onmany factors including

the species of plant, temperature, etc.

Quantum efficiency measures the energetic efficiency

of photosynthesis. The quantum yield of oxygen evolution

describes the amount of oxygen evolved per photon. Theo-

retically, it requires eight photons to evolve onemolecule ofO2

(as described further below), so the theoretical quantum

efficiency is 0.125 mol O2/mol photons. The measured value

is always lower, as there are inevitably energy losses due to

heat production, fluorescence, and other factors. Quantum

efficiency is a useful indicator of stress, for example, as stress

induces nonphotochemical quenching, which lowers quan-

tum efficiency. The quantum efficiency is not very dependent

on light wavelength as long as the light has sufficient energy to

drive photosynthesis. Although plants do not absorb much

green light, it is capable of initiating photochemistry.

PLASTIDS AND CHLOROPLASTS

Plastids are essential organelles in most plant cells although

generally not found in the spermcells of pollen. Severalmetabolic

processes besides photosynthesis occur in plastids including

nitrogen and sulfur assimilation and the synthesis of secondary

metabolites, pigments, fatty acids, and various hormones. It is

important to remember that plastids are descendants of endo-

cytosed prokaryotes. Many features of the ancestral prokaryote

can still be detected, including the vestigial circular plastid

genome, division by fission, and the chemical composition of

the lipid membranes.

Chloroplast Development and Differentiation

Plastids are never synthesized de novo but are always inherited

by daughter cells during cell division. Some single-celled algae

have only a single chloroplast. Plant cells have more; undifferen-

tiated meristematic cells may have as few as 10 proplastids that

are small and have minimal expansion of internal membranes,

whereas leaf mesophyll cells may have 100 or so chloroplasts.

Proplastids differentiate into several forms depending on the

cell type. In some root cells, proplastids differentiate into amy-

loplasts thataccumulatestarchgranulesandhaveaspecial role in

gravity perception. In fruits and flowers, some plastids differen-

tiate as carotenoid-filled chromoplasts. In the prephotosynthetic

tissues of dark-grown or etiolated seedlings, plastids partially

differentiate into etioplasts, which are primed to differentiate into

chloroplasts upon illumination. Protochlorophyllide, the precur-

sorof chlorophyll, is attached to theprolamellar body,which is the

thylakoid precursor in etioplasts.

When etioplasts are exposed to light, within just a few hours

there is a light-induced conversion of protochlorophyllide to

chlorophyllide and then to chlorophyll, and theprolamellar bodies

reorganize into thylakoids. Light also triggers changes in gene

expression in the nucleus. Nuclear genes encoding photosyn-

theticapparatusare induced, asare thegenesencodingenzymes

required forpigmentsynthesis.These transcriptional changesare

accompanied by large-scale metabolic changes to prepare the

organelle for its role as a photosynthetic entity.

Ultrastructure of Chloroplasts

Chloroplasts of higher plants are typically oval in shape and;5 to

10 mm long. They are bounded by an outer layer that is a double

membrane (meaning that it has two lipid bilayers) known as

the envelope. Inside the envelope is an aqueous layer called the

stroma, which is largely filled by the photosynthetic membranes

called thylakoids. These membranes anchor the photosynthetic

apparatus and are needed for the establishment of a proton

gradient to drive ATP synthesis. The space inside the thylakoid

membranes is known as the thylakoid lumen. Plastoglobules are

lipoprotein particles permanently attached to the thylakoid mem-

branes, which contain biosynthetic enzymes responsible for the

production of tocopherols (reactive oxygen scavengers, also

known as vitamin E) and phylloquinone (polycyclic aromatic

ketones, part of the electron transport chain of PSI and also

4 The Plant Cell

known as vitamin K1). Of the 3500 to 4000 different proteins

found in the chloroplast, only ;75 to 80 are encoded by the

plastid genome; genes encoding all the rest are located in

the nucleus.

Chloroplast Envelope

Chloroplasts are enclosed by a double membrane called an

envelope. Lipid composition, particularly enrichment in galac-

toglycerolipids, and the presence of distinctly prokaryote-like

membrane proteins indicate that both the outer and inner

envelope membranes are derived from the outer and inner

membranes of ancestral prokaryote. Furthermore, the lipid

composition of the inner envelope membrane is similar to that

of the thylakoid membrane. Membrane proteins in the envelope

include protein import machinery through which the photosyn-

thetic complex proteins are translocated, as well as numerous

transporters for the metabolites that move into and out of a

functioning chloroplast.

Thylakoid Membranes

The inner structures of land plant chloroplasts are beautiful

and distinctive. Thylakoids serve as anchors for large protein

complexes of light-dependent reactions and serve as barriers

to generate the proton gradient required for ATP synthesis.

Undifferentiated plastids have little or no thylakoid mem-

branes. Upon light stimulation, internal thylakoid membranes

proliferate, forming discs of membranes known as grana that

resemble stacks of coins and are connected by regions of

membrane known as the stromal thylakoid. Some regions of

the thylakoid membranes are pressed against each other and

known as appressed membranes. PSII is mainly found in the

appressed regions of themembrane. Themargins of the grana

and the stromal membranes are not pressed against other

membranes and are therefore known as unappressed mem-

branes. The bulkier protein complexes PSI and ATPase predom-

inate on the unappressed membranes, and cytochrome b6f is

uniformly distributed throughout. This physical separation

of photosystems allows a greater fine-tuning of photosynthe-

sis and responses to changing light intensity and metabolic

demands.

STRUCTURE AND FUNCTION OF PHOTOSYNTHETIC

COMPLEXES

Each of the photosynthetic apparatus described below aremem-

brane anchored and comprise multiple proteins and pigments or

cofactors. Classically, the components of these complexes were

characterized by their physical separation through density-

gradient centrifugation and gel electrophoresis, and these

methods are still important. In recent years, our understanding

of photosynthesis has been greatly enhanced through high-

resolution structural analysis based on crystallography of the

key components. Comparisons of the structures obtained

from plant, algae, and cyanobacteria reveal insights into

how evolution has shaped these complex machines.

Linear Electron Transport

Here, we describe the machinery that carries out the light

reactions of photosynthesis in plants, initially focusing on the

linear electron transport (LET) that predominates in most con-

ditions. In LET, light energy transferred from the LHCs to PSII

and PSI oxidizes their reaction centers, driving the linear transport

of electrons fromH2O to NADPH. Specifically, electrons flow from

water through PSII to the mobile electron carrier plastoquinone

(PQ), through cytochrome b6f, then to PSI by another mobile

electron carrier plastocyanin (PC), then via ferredoxin to reduce

NADP1 to NADPH.

Concomitant to the reduction of NADP1 to NADPH, the elec-

tron transfer events also contribute to a proton gradient across

the thylakoid membrane, which is used for the synthesis of ATP

by ATP synthase. Together, ATP and the reducing power of

NADPH are used in the biosynthetic reactions of the carbon-

fixing reactions of photosynthesis.

Structure and Function of PSII-LHCII Supercomplex

PSII carries out light-energized electron transport that leads

to water splitting and oxygen release. PSII is a multi-protein,

multi-pigment complex that spans the thylakoid membrane.

PSII operationally includes the core PSII complex along with

the oxygen-evolving complex (OEC) that is highly conserved

between plants, algae and cyanobacteria, and the peripheral

antenna or light-harvesting complexes (LHCII) that are more

divergent.

Core PSII Complex

The structure of PSII from cyanobacteria has been resolved at

1.9A,whichallowsmostoftheproteinsandcofactorstobeidentified.

Thecyanobacterialcomplex ismadeupof17membrane-spanning

proteins, three peripheral proteins, 35 chlorophyll molecules,

and 11 b-carotenes. Genes encoding PSII proteins are named

Psb genes.

PSII functions as a dimer. Themost highly conserved elements

are those in the reaction center involved in light capture and

electron transport as well as those involved in water splitting.

Eachmonomer has a distinct reaction centermade upof proteins

D1 (PsbA) and D2 (PsbD) as well as an inner light-harvesting

complex composed of chlorophyll binding proteins CP43 (PsbC)

and CP47 (PsbB). The function of the reaction center proteins is

mainly to anchor the pigments and electron carriers in place and

to provide an appropriate environment for efficient energy trans-

fer. As described below, the D1 protein of PSII is subject to light-

dependent turnover and repair, a property that may provide

a safety valve for the protection of other sensitive photosynthetic

components.

A pair of chlorophyll molecules forms the core of the reaction

center. Light excites chlorophyll to form Chl*. Charge transfer

occurswhenanelectron leavesChl*, formingChl1. Theelectron is

passed to pheophytin (chlorophyll that lacks Mg; Pheo) to pro-

duce Pheo2. Pheophytin passes the electron to the primary

acceptor, QA, which is a tightly bound PQ. PSII contains two

(possibly three) PQ cofactors. One, QA, is fixed within the PSII

November 2015 5

structure andservesasaconduit for electrons frompheophytin to

the second PQmolecule. The second PQmolecule binds to PSII

at theQB site. Plastoquinone is a two-electron carrier, which after

accepting twoelectrons is in the formPQ22. Twoprotons fromthe

stroma bind to it to form PQH2 (plastoquinol), an uncharged

molecule. This reduced, uncharged form can dissociate from

thePSII complex, physically carryingelectrons to thecytochrome

b6f complex and simultaneously transferring protons from the

stroma to the lumen.

A pool of PQ/PQH2 molecules exists within the thylakoid

membrane to shuttle electrons between PSII and cytochrome

b6f. Once PQH2 is released from PSII, another PQ molecule will

bind to PSII in its place to accept electrons from another round of

excitation. It should be noted that PSII can only give rise to one

charge separation event at a time. Thus, twophotonsare required

for the complete reduction and release of a single PQH2molecule

from the complex.

Extrinsic Proteins and the Oxygen-Evolving Complex

The OEC resides on the luminal surface of PSII and includes the

catalytic center, loops of intrinsic proteins, and some extrinsic

proteins. There is one OEC per PSII monomer. The catalytic

center core is an inorganic Mn4CaO5 cluster that performs the

mechanistically challenging reaction of removing four tightly

bound electrons and four protons from water to form O2. The

cluster cancycle throughdifferent oxidationstates tosequentially

provide four electrons and reach the oxidation status required to

split two water molecules and release O2. The four H1 released

from the water molecules on the luminal side of the membrane

contribute to the proton gradient across the thylakoid.

Amino acid residues from the PSII membrane proteins D1, D2,

andCP43coordinate theassociationof theMn4CaO5cluster. The

removalof the fourelectrons fromwaterby theMn4CaO5cluster is

a four-step process. One electron is removed from water with

each photon-induced charge separation at the reaction center

and then passed via a redox-active tyrosine residue on the D1

protein (Yz) to fill the “hole” left in the PSII reaction center. Overall,

the removal of four electrons from two water molecules requires

the absorption of four photons by the reaction center that results

in the release of four H1 into the lumen. The electrons transferred

during the turnover of PSII are ultimately used to reduce two PQ

molecules to two PQH2 molecules, a process that also uses H1

from the stromal side of the thylakoid membrane.

Depending on the type of photosynthetic organism, at least

three extrinsic proteins are associatedwith the luminal face of the

complexwhereoxygenevolutionoccurs.Theseextrinsicproteins

are not required for oxygen production but do enhance it. These

proteins contribute to the stability of the Mn4CaO5 cluster,

facilitate the catalytic rate under physiologically relevant con-

ditions, andcontribute to theassemblyof thePSII enzyme.PsbO

is the only extrinsic protein conserved among cyanobacteria,

algae, and plants. This protein is also known as the manganese

stabilizing protein. Other extrinsic proteins include PsbP (plants

only), PsbQ (plants, cyanobacteria, and red algae), PsbU, and

PsbV (cyanobacteria and red algae). Thus, the complement of

extrinsic proteins associated with PSII varies among photosyn-

thetic organisms.

PSII Peripheral Antenna Complex or LHCII

Onemajor difference between plants and cyanobacteria is in the

structure of the peripheral protein/pigment complex that funnels

light to the photosynthetic core complex.

In cyanobacteria and red algae, the PSII accessory light-

harvesting system is composed of phycobilisomes, made up

of proteins and phycobilins. A central core anchors six rods

made up of phycobiliproteins and linker proteins that radiate

outwards. The entire phycobilisome structure is attached to the

stromal side of PSII and extends into the stroma.

In plants and green algae, the light-harvesting complex of PSII

sits in the thylakoid membrane. In vascular plants, it consists of

two layers: major, more abundant trimeric LHCII proteins and

minor, less abundant monomeric LHCII proteins. Based on

acrystal structure fromspinach (Spinacia oleracea), eachmono-

mer is made up of a single polypeptide chain, 14 chlorophylls,

and four carotenoids. In addition to the chlorophyll amolecules

found in the core complex, the outer LHCs also bind chlorophyll

b molecules and xanthophylls. The arrangement of the outer

layer varies with light and other conditions. Several additional

proteins are involved in the dynamic interaction between PSII

and LHCIIs. Under certain conditions LCHII complexes can

move away from PSII to decrease the amount of light funneled

into the PSII reaction center, providing flexibility to the photo-

synthetic system (see below). Furthermore, the ratio of light-

harvesting complexes to reaction centers varies with the light

intensity to which a plant is acclimated.

Q Cycle and Cytochrome b6f: Electron Transport from

Plastoquinol to Plastocyanin

Cytochrome b6f is a membrane-embedded complex that func-

tions as a dimer, with each dimer made up of eight subunits.

Some of the subunits are very similar to those in the cytochrome

bc1 complex that carries out electron transport inmitochondrial

respiration, and these complexes clearly share an evolutionary

origin. Theprotein subunits are cytochromeb6 andcytochrome f

proteins (hence the name of the complex), an iron-sulfur protein

also known as a Rieske protein (after its discoverer, John Rieske),

a 17-kD protein, and four shorter proteins. Unlike the related

complex in mitochondria, cytochrome b6f also binds chlorophyll

and b-carotene, as well as an unusual heme not found in the

mitochondrial complex.

Cytochrome b6f is sometimes referred to as plastoquinol-

plastocyanin oxidoreductase. Electrons from PSII are carried

by plastoquinol (PQH2) to cytochrome b6f where the Q-cycle

takes place. The Q-cycle (which also occurs in mitochondrial

electron transport) is a series of oxidation and reduction reactions

of that allows for maximal contribution to the proton gradient

and transfers electrons from a two-electron carrier (PQH2) to

a single-electron carrier (plastocyanin, a small protein). As elec-

trons are passed through cytochrome b6f, protons are passed

from the stroma to the lumen.

The Q cycle can be described as a series of steps. Reduced

PQH2 generated by PSII moves to the cytochrome b6f complex

where it is oxidized to PQ. During this initial oxidation, the release

of two protons into the thylakoid lumen results in one electron

6 The Plant Cell

transferred to PC, and the other electron to one of the two b-type

hemes borne by the complex and located on the opposite side of

themembrane.Another roundof oxidationoccurswhenasecond

PQH2 molecule delivers its electrons to cytochrome b6f. Again,

the release of one of its electrons results in the transfer of two

protons to the thylakoid lumen and one electron to plastocyanin.

Theother electron is transferred to another b-heme.Next, the two

reduced hemes cooperate to reduce a PQ into a PQH2, which is

then released from cytochrome b6f into the thylakoid membrane.

Thus, there is a recycled pool of PQ/PQH2 between PSII and

cytochrome b6f for the transfer of electrons between these two

complexes. The stoichiometry of the Q cycle is: two PQH2 enter,

two electrons are transferred to plastocyanin, four protons are

pumped into the lumen, and onePQH2 is regenerated; thus, each

molecule of PQH2 delivers four protons to the lumen and two

electrons to plastocyanin.

On the luminal side of the cytochrome b6f complex, electrons

are passed to themobile electron carrier plastocyanin, a copper-

containingenzyme. InChlamydomonasandcyanobacteria,butnot

plants,plastocyanincanbereplaced functionallybycytochromec6whencopper is scarce. Plastocyaninor (cytochromec6) transports

electrons from cytochrome b6f to PSI.

Structure and Function of PSI-LHCI

PSIuses light energy to transfer electrons fromthesolubleelectron

carrier plastocyanin on the luminal side of the thylakoidmembrane

to ferredoxinon thestromal side. It isamulti-protein,multi-pigment

complex that spans the thylakoid membrane and operationally

includes the core PSI complex and the peripheral antenna or light-

harvesting complex that is highly divergent.

Core PSI Complex

In terrestrialplants,PSI isacomplexof17proteinsubunitsandover

200 prosthetic groups, mainly, chlorophylls but also three Fe4S4

clusters and a few carotenoids and phylloquinones. The structure

is dominated by two large and related proteins, PsaA and PsaB.

Like in PSII, the heart of PSI is a pair of chlorophyll molecules,

one of which is excited and oxidized by light energy transduced

fromthesurroundingnetworkofpigmentmolecules.Thiselectron

moves through a chlorophyll to a phylloquinone and from there

through threeFe4S4 clusters andfinally to ferredoxin (Fd), a stable

electron acceptor. Ferredoxin is a soluble electron carrier that

binds reversibly to PSI and transfers electrons to NADP1 by the

action of the enzyme ferredoxin:NADP1 oxireductase. The electron

hole created when PSI absorbs a photon is filled by electrons

donated by the soluble electron transporter plastocyanin (or

cytochromec6). In cyanobacteria,PSI isusually foundasa trimer

but can also be present in tetramers or dimers, and it is thought

that this multimer arrangement increases the functional size of

the light-harvesting antenna for dim-light conditions.

PSI Peripheral Light-Harvesting or Antenna Complexes

As described for PSII, a key difference between plant, cyanobac-

terial, and algal PSI complexes is in the nature of the peripheral

light-harvesting complex. In plants and green algae, PSI is present

asamonomer that issurroundedbyanouterantennasystemmade

up of LHCI complexes that are embedded in the thylakoid mem-

brane and arranged in a half-moon shape. This crescent-shaped

arrangement ismadeupofasingle layerof fourproteins invascular

plants but a double layer of eight proteins in Chlamydomonas.

Bacterial PSI has not been purified with associated phycobili-

somes, but recently single-particle electron microscopy images

have been obtained and their energy transfer has been observed

by fluorescence spectroscopy.

Structure and Function of ATP Synthase

Proton translocation coupled to light-driven electron transfer

builds up the pH gradient (DpH) and electrical potential (DC)

across thylakoids. DpH and DC form the transthylakoid proton

motive force (pmf), which is the essential driving force for the

phosphorylation of ADP to produce ATP.

ATP synthase couples the synthesis of ATP fromADP andPi to

the movement of protons from inside the lumen (high concen-

tration; low pH) to the stroma (low concentration; high pH) across

the thylakoid membrane. The enzyme is also referred to as an

ATPase, as the reactioncanoccur inbothdirections (hydrolysis of

ATP linked to pumping protons against the gradient). The chlo-

roplast ATP synthase that carries out photophosphorylation is an

F-ATPase that is related to the mitochondrial ATPase and the

bacterial F-ATPase. The plant VH1-ATPase that is involved in

proton pumping across endomembranes is also an evolutionarily

related enzyme.

F-ATPases are large enzymes with more than 10 subunits that

assemble into two complexes. F0 is a membrane-embedded com-

plex that conducts protons and is made up of 10 to 15 copies of

subunit c, and one each of a, b, and b’ (in the older literature these

were referred to as subunits I to IV). F1 is a peripheral complex that

hydrolyzes/synthesizes ATP, is attached to F0 by a stalk, and is

made up of three copies each of thea andb subunits and one each

of the g, d, and e subunits. F1 is sometimes referred to as “coupling

factor 1” because when it is stripped off the membranes proton

efflux isuncoupled fromATPsynthesisduetothe lossof thecatalytic

complex of ATP synthase. Restoring F1 to stripped membranes

restores the coupling of proton efflux to ATP synthesis.

Paul Boyer and John Walker were awarded the 1997 Nobel

Prize in Chemistry “for their elucidation of the enzymatic mecha-

nism underlying the synthesis of adenosine triphosphate (ATP).”

Boyer laterwrote, “All enzymesarebeautiful, but theATPsynthase

is one of themost beautiful as well as one of themost unusual and

important,” and it is beautiful, particularly when seen in electron

micrographs. The enzyme uses a rotary catalytic mechanism

through which proton efflux provides energy for ATP synthesis.

Briefly, proton efflux through the F0 channel causes the a3b3

hexamer of F1 to rotate around the g subunit, and the conforma-

tional changes energize bond formation between ADP and Pi.

Thegandesubunitsworktogether toregulatethe levelofenzyme

activity, particularly in response to shifts between light and dark.

After a dark-to-light transition, the actions of PSII, cytochrome b6f,

and PSI cause protons to accumulate in the thylakoid lumen

and reduced ferredoxin and thioredoxin to be produced.

The chloroplast enzyme requires an acidified thylakoid lumen

November 2015 7

and reduced thioredoxin to be active, which prevents the

enzyme from running backward (hydrolyzing ATP) in the dark.

PATHWAYS OF ELECTRON TRANSPORT

There are at least three routes of photosynthetic electron trans-

port along the thylakoid membrane: (1) linear electron transport

that includes PSII, cytochrome b6f, and PSI; (2) cyclic electron

transport that includes PSI and cytochrome b6f; and (3) the

water-water cycle that uses PSII, cytochrome b6f, and PSI but

does not produce NADPH. The major pathway, linear electron

transport, is the onemost often taught, but alone it is inadequate

to meet the metabolic needs of the plant. The other pathways

serve to balance production of NADPH and ATP (both cyclic

electron transport and the water-water cycle produce a proton

motive force for ATP production but do not yield NADPH).

Furthermore, these alternative electron transport pathways

can provide a photoprotective function.

Linear Electron Transport Involves PSII and PSI

As described above, in LET, light energy transferred from the

LHCs to PSII and PSI oxidizes their reaction centers, driving the

linear transport of electrons from H2O to NADPH. The route of

linear electron transport can be summarized as:

H2O~PSII~PQ~Cyt  b6 f~PC~PSI~Fd~NADP1~NADPH:

As a consequence of the Q-cycle, during LET the lumen gains

three protons for each electron that is transported: one proton

for each electron released by the oxidation of H2O at PSII and two

protons for each electron transferred through cytochrome b6f

to PSI. Therefore, linear electron transport has a proton/electron

(H1/e2) ratio of 3:1. The ratio of e2/NADPH is 2:1 (it requires two

electrons to reduceoneNADP1 toNADPH); thus, the ratio ofH1 to

NADPH is six. The stoichiometry of theATP synthase is such that it

takes 14 H1 to produce 3 ATPs (a ratio of H1/ATP of 4.67).

Consequently, the ATP/NADPH output ratio of LEF is 1.28 (which

equals 6/4.67). The pooled energy requirements for CO2 fixation in

theCalvin-Bensoncycle togetherwithphotorespirationandnitrate

assimilation inC3plantsare;1.43ATP/NADPH.Thus, LEF results

in a deficit of;0.15 ATP per NADPHproduced. This deficit can be

alleviated by cyclic electron transport, which passes electrons

cyclically throughPSIandcytochromeb6f leadingtothemovement

of protons into the lumen and subsequent ATP synthesis without

NADPH production.

Cyclic Electron Transport around PSI

Cycle electron transport (CET) uses all of the machinery of LET

except PSII. CET is a series of reactions in which electrons from PSI

reducePQtoPQH2,whichdeliversprotonstothelumenintheQcycle

ofcytochromeb6f.TheelectronsaretransferredtoPCandreturnedto

PSI to keepCETgoing; theelectronscycle. Protons released into the

thylakoid lumen contribute to the transthylakoid protonmotive force

(pmf). Therefore, CET contributes to the synthesis of ATP but not

NADPHandsoadjusts theATP/NADPHratioduringphotosynthesis.

There are two possible CET pathways. In both routes, PQH2

recycles electrons back to PSI through cytochrome b6f complex

and plastocyanin. The two routes of CET can be summarized as:

ð1ÞPSI~Fd~PQ~Cyt  b6 f~PC~PSI  ðdependent  on  PGR5=

PGRL1  in  plantsÞ

ð2ÞPSI~Fd~NADP1~PQ~Cyt  b6 f~PC~PSI  ðdependent 

on NDH  complex Þ

The first route is sensitive to antimycin A and transports electrons

from PSI to Fd and then to PQ by ferredoxin plastoquinone re-

ductase. In Arabidopsis thaliana, the regulatory proteins PGR5

(PROTON-GRADIENT REGULATED5) and PGRL1 (PGR-LIKE1)

form a complex with PSI and mediate this antimycin-A sensitive

CET. This pathway appears essential for photosynthesis because

pgr5 and pgrl1 mutants have compromised photosynthesis. The

second CET route transfers electrons fromPSI to NADP1 through

Fd and then to PQ by NAD(P)H dehydrogenase complex (NDH

complex). The secondpathway is not essential for photosynthesis

because mutants deficient in NDH complex grow as well as the

wild-type plants in greenhouse conditions; however, the NDH-

dependent CET pathway may function under stress or becomes

essential when the first pathway is absent. Due to the difficulties to

measure CET accurately in vivo, controversies exist about the two

pathways of CET and how they are regulated.

The relative contribution of CET to photosynthesis depends on

themetabolicneedsof theplant.Thephotosyntheticneeds forATP

are higher in organisms or cells that concentrate CO2 such as

Chlamydomonas and C4 plants such as maize (Zea mays), so the

proportion of CET is greater than in C3 plants. Nevertheless, in C3

plants, CET seems to have a protective role in some conditions,

such as during the induction of photosynthesis or during low-CO2,

high-light, or drought conditions. Furthermore, by contributing to

the accumulation of H1 in the lumen, elevated CEF contributes to

photoprotective processes such as nonphotochemical quenching.

The Water-Water Cycle and Chlororespiration

The water-water cycle, also called the Mehler reaction, involves

the LET chain but uses oxygen instead of Fd as an electron

acceptor at PSI. Transferring electrons to O2 reduces it to super-

oxide (O2), which is subsequently reduced to H2O by superoxide

dismutase and ascorbate peroxidase. The water-water cycle

functions like a cycle because electrons are extracted from

H2O at PSII, transferred through PQ, cytochrome b6f, PC, and

PSI to reduce O2 and ultimately produce water. The water-water

cycle uses the linear transport chain and contributes to trans-

thylakoid pmf, but because the electron eventually goes to O2

instead of NADP1, it contributes to the production of ATP but not

NADPH. This pathway can be summarized as:

H2O~PSII~PQ~Cyt  b6 f~PC~PSI~O2~H2O2~H2O:

The water-water cycle is thought to function for photoprotection.

During the light-induction phase (dark-adapted plants in light

condition), enzymes in the Calvin-Benson cycle initially are

8 The Plant Cell

not fully activated, so there is no consumption of NADPH for

CO2 fixation. Thewater-water cycle serves as a safety valve to

reduce the production of NADPHandbuild up a transthylakoid

proton motive force to trigger light dissipation through NPQ.

The water-water cycle has the disadvantage of generating

reactive oxygen species, which often lead to significant cel-

lular damage and must be quenched before damage occurs.

Chlororespiration is a light-independent reaction that transfers

electrons to oxygen and is also considered a mechanism to

alleviate excitation pressure on electron transport. Physiologically,

there are conditions in which the PQH2 pool becomes overly re-

duced, leavingnowhere for theelectronsproduced fromPSII togo.

Plastid terminal oxidase is a plastid enzyme, related to mitochon-

drialalternativeoxidase,whichoxidizesPQH2toPQbyreducingO2

to H2O. In other words, it draws electrons out from the electron

transport chain to prevent overreduction. Chlororespiration lowers

the excitation pressure on electron transport and likely reduces

overall oxidative stress. Chlororespiration links the redox state of

thePQpoolwith thatof thestroma,placingplastid terminal oxidase

at the crossroads of many metabolic processes.

In cyanobacteria, flavodiiron proteins (encoded by Flv genes)

are involved in photoprotection. Flv1 and Flv3 are necessary for

growth in fluctuating light conditions and function in aMehler-like

reaction that reduces O2 to H2O. Flv2 and Flv4 are thought to

protect PSII from damage by accepting electrons from the

electron transport chain. Furthermore, a thylakoid membrane-

associated terminal oxidase is also required to protect the cells

from damage during fluctuating light regimes.

DAMAGE AVOIDANCE AND REPAIR: ACCLIMATIONS TO

LIGHT STRESS

Photosynthetic organisms have to function in variable light

regimesandconditions. Theyare regularly subjected tovariation in

light intensity, light quality (relative amount of light of each wave-

length), and angle of light incidence.Someof these changes occur

over long time periods (seasons), others diurnally, and others on

the order of minutes, for example, the periodic sunflecks experi-

enced by leaves beneath a canopy. Variable light occurs against

a backdrop of variations in metabolism (e.g., energy demand and

nutrient availability) and environment (e.g., temperature and water

availability). Ashasbeendiscoveredbyscientists trying todevelop

artificial photosynthesis, it is rather amazing that such a high-

energy process can be maintained. Key to its success is the

ability to repair damage and to avoid it. Plants display numerous

strategies for both.

Photosynthetic organisms manage photon interception and

usage in the variable light environment to enable efficient pho-

tosynthesis and to minimize photodamage. As described pre-

viously, the rate of photosynthesis increases with light only at

relatively low light intensities, demonstrating that photosynthetic

organisms are regularly exposed to excess excitation energy.

Anytime when light energy exceeds the light saturation point,

plantsaresusceptible todamagedue to theoverreducedelectron

transport chain. When the system is light saturated, there is an

increased probability that excited chlorophyll will convert to the

excited triplet state (3Chl*),which iscapableof transferringenergy

tooxygen toproduce reactiveoxygenproducts.Therefore, plants

are exquisitely sensitive to the indicators of excess excitation

pressure, including a buildup of protons in the thylakoid lumen

and an overly reduced PQH2 pool; low lumen pH and high PQH2

levels each trigger regulatory responses to promote energy dis-

sipation. The timescale of responses can be extremely rapid (on

the order of seconds) and involve conformational or covalent

changes to the light harvesting machinery or occur more slowly

and involve changes in gene expression. Rapidly reversible re-

sponses to excess light are described as photoprotection,

whereas photodamage is reversed more slowly, and this differ-

ence is measureable.

Movements to Optimize Light Interception

Oneof themostdirectways tooptimize lightharvesting is tomoveor

to change orientation relative to sunlight. Single-celled organisms

can move toward light under light-limiting conditions to maximize

photosynthesis or away from excess light to prevent photodamage

inaprocesscalledphototaxis.Similarly,chloroplasts in leafcellscan

move to increase or decrease their interception of light through

accumulation or avoidance responses respectively. In many plant

species, leaf angle is sensitive to light, with the leaves rotating or

lowering at midday or other times when light incidence can exceed

photosynthetic capacity. In most cases, these movements are

mediated by light perception by photoreceptors such as photo-

tropins and phytochromes that are distinct from the pigments that

capture light forphotosynthesis,although there isalsoevidence that

the activity of PSII can affect leaf movement.

Acclimation via Stoichiometric Changes in Complex

Abundance

Arelativelyslowand long-termresponsetoprolongedlight intensity

changes is a change in the size of the light-harvesting systems, in

which their size is inversely proportionally to light intensity. These

changes arise through transcriptional and posttranscriptional con-

trolsofLHCgenesasa resultofsignaling fromthechloroplast to the

nucleus and involve both induction of expression of LHC genes in

low light and turnover of LHC proteins in high light. Similarly,

acclimation to high light involves an increase in the amount of

PSI, PSII, cytochrome b6f, and ATP synthase relative to the light

harvesting systems, a change that is reflected in an increased ratio

ofchlorophylla tochlorophyllb.Finally, lightspectralqualitiesaffect

the relative levels of PSII to PSI; specifically, in shaded canopies,

light is enriched for the longer wavelength light that is preferentially

absorbed by PSI, so the plant acclimates by increasing the abun-

dance of PSII. These acclimations can be quantified bymeasuring

the relative abundance of chlorophyll a to chlorophyll b and by

measuring the light saturation point.

Excess Light Energy Dissipation through

Nonphotochemical Quenching

Light energy absorbed by antenna or light-harvesting complexes

can have three fates. It can be processed for photochemistry,

November 2015 9

emitted via fluorescence, or dissipated byNPQ. The termNPQ

comes from the standard method for measuring photosyn-

thetic activities. By measuring the amount of chlorophyll

fluorescence under various conditions, it is possible to mea-

sure the amount of light that is quenched (i.e., not fluoresced)

due to photochemistry (termed photochemical quenching

[qP]) or NPQ.

NPQ can be divided into at least three different components

that can be distinguished by the relaxation kinetics of chlorophyll

fluorescence.Energy-dependentquenching (qE) isdependenton

the acidification of the lumen. It is the major and most rapid

component of NPQ in most algae and plants and relaxes within

seconds to minutes. State-transition quenching (qT) involves

movement of the mobile light-harvesting LHCII complex and

relaxes within tens of minutes. Photoinhibitory quenching (qI) is

caused by photoinhibition of photosynthesis and relaxes very

slowly in the range of hours. Each of these is described below.

Energy-Dependent Quenching (qE)

One consequence of excess light energy is a buildup of protons

in the thylakoid lumen that accumulate faster than they can be

processed by ATP synthase. Thylakoid lumen acidification is

sensed by special members of the LHC protein family that are

protonated on their luminal surface: PsbS in plants including

bryophytes, andLHCSR (light-harvestingcomplex stress related)

in Chlamydomonas and bryophytes. The acidic lumen also in-

duces the activity of an enzyme, violaxanthin deepoxidase,which

converts the carotenoid violaxanthin to zeaxanthin. This change

induces conformational changes in the PSII light-harvesting sys-

tem, causing intercepted light energy to be dissipated as heat

rather than transferred to the reaction center. The exact photo-

protective nature of this response continues to be debated and

may involve both transfer of energy from excited chlorophylls to

carotenoids in LHCII as well as charge transfer occurring in the

antenna complex that prevent light from reaching reaction center

chlorophylls.When the thylakoidpHreturns tonormal, zeaxanthin

is enzymatically convertedback toviolaxanthinand light energy is

again channeled into the reaction center.

Energy-dependent quenching is usually the dominant compo-

nentofNPQinplants.Thecontrol ofqEbyDpHallows inductionor

relaxation of qE within seconds in response to changing light

intensity to prevent excessive excitation of PSII centers under

high light or tomaximize light harvesting for photosynthesis in low

light. The acidification of the thylakoid lumen that results from

alternative electron transport, such as cyclic electron transport

and water-water cycle, promotes qE.

State Transition (qT)

Photosynthesis ismostefficientwhenPSI andPSII areoperating in

a balanced way, yet their antenna complexes have different com-

plementsofchlorophyllaandbandcarotenoids (thereforedifferent

absorption spectra), and light quality is variable, meaning that the

excitation energy transferred to PSI and PSII can be unbalanced.

One way to address this is through conformational changes in the

LHCII light-harvesting machinery that are referred to as a state

transition (qT). To some extent, this causes a channeling of light

energy fromLHCII to PSI, but it appears as though themajor effect

is to decrease light energy transfer to the reaction center of PSII.

State transition is thought to affect;20%of LHCII in Arabidopsis,

but as much as 80% of LHCII in Chlamydomonas.

Themolecular basis for state transition rests on the redox state

of the plastoquinone pool. Overexcited PSII leads to an accu-

mulation of reduced plastoquinone, PQH2.WhenPSII gets ahead

of PSI, the binding of reduced PQH2 to the cytochrome b6f

complex activates LHCII kinase; conversely, when PSI gets

ahead of PSII, the plastoquinone pool becomes oxidized and

LHCII kinase is inactivated. LHCII kinase was first identified in

Chlamydomonas and named Stt7, and later in Arabidopsis and

named STN7. Stt7 and STN7 are structurally and functionally

related and both are attached to thylakoid membranes. When

activated, they phosphorylate LHCII to induce state transition.

This reaction is reversible by the actionof a specificphosphatase.

Photoinhibition (qI)

Photoinhibition (qI) is light-induced reduction in photosynthetic

quantum yield that occurs as a consequence of photodamage

to the D1 protein of PSII (see below) or other slowly reversible

damage to the photosynthetic machinery. Light of any wave-

length can lead to photodamage, but plants can be particularly

sensitive to short wavelength blue or UV light hitting and dam-

aging the Mn4Ca cluster in the oxygen-evolving complex. This

damage slows the rate of electron flow from theOEC into the PSII

reaction center, which can lead to the production of reactive

oxygen species, which in turn decreases the rate of repair of the

D1 protein of the PSII reaction center. Photoinhibition can be

measured through fluorescence methods and is a slowly revers-

ible source of NPQ.

PSII Photodamage and Repair

While PSII requires light for its catalytic activity, this complex is

irreversibly damaged by light, more so than any of the other

complexes of the light-dependent reactions. PSII damage in-

creases with light intensity, but occurs to some extent at all light

intensities. The primary site of damage is the core D1 protein. To

recover activePSII complexes, thedamagedD1proteinmust be

recognized, proteolytically removed from the complex, and

replaced with a newly synthesized D1 protein. This involves

migration of the damaged PSII complex from the appressed

grana stacks to the unappressed regions, insertion of a newly

synthesized D1 polypeptide, and reassembly of the PSII com-

plex. To maintain a steady state level of functional PSII com-

plexes, photosynthetic organisms have dedicatedmachinery to

recognizeD1damageand repairPSII. It hasbeensuggested that

rather than being a defect, PSII’s sensitivity to photodamage

serves as a safety valve to protect PSI from damage.

Sensitivity of Photosynthesis to Heat, Drought, and Other

Stress

As described above, many short-term and long-term acclima-

tions enable photosynthetic processes to be maintained even as

10 The Plant Cell

light intensity changes. Essentially any kind of stress, ranging

fromnutrient stress,drought stress, temperaturestress, andeven

pathogen attack, can affect carbon fixation, which in turn affects

the flow of electrons through the photosystems as well as the

cycling of ATP/ADP, NADPH/NADP1, and PQ/PQH2. Changes in

DpHand redox state of thePQpool are important in regulating the

core photosynthetic processes as well as pathways to confer

protection from photooxidative damage.

However, there are additional ways that some stresses can

affect photosynthesis. For example, high temperatures increase

thefluidityof the thylakoidmembrane,which inturnaffectselectron

transport, proton efflux through ATP synthase, and ionmovement

across thylakoid membranes. One response to increased thyla-

koid leakiness is an increase in CET to maintain proton motive

force. As another example, stomatal closure in response to water

deficitdecreases theuptakeofCO2,meaning that thepoolsofATP

and NADPH accumulate, resulting in an overreduced electron

transport chain and the accumulation of protons in the thylakoid

lumen. This increase in transthylakoidprotonmotive force induces

an increase in energy-dependent quenching, which helps to dis-

sipateexcess lightenergyandprevent light-induceddamage.Cold

temperatures slowmetabolic processes but do not affect the rate

of light interception, and cold temperatures are frequently asso-

ciated with an increase in photooxidative damage. One of the big

ongoing research areas is to integrate what we know about the

light-dependent reactions under optimal conditions with the

whole-plant physiological changes that occur with stress.

Regulation and Retrograde Signaling

Many of the biochemical shifts and responses described above

occur autonomously in plastids and can be measured in isolated

chloroplasts. However, other photosynthetic acclimations and

stress responses involve changes in nuclear gene expression.

Not only are most of the chloroplast-localized proteins encoded

by genes in the nucleus, but the chloroplast operates within the

larger context of the cell and the rest of the cell’s complement of

nuclear-encoded proteins. Thus, there is a clear need for in-

tegration between the functions of the plastid, the nucleus, the

cytosol, and the mitochondrion.

Information flows from chloroplast to nucleus. This has been

shown in several ways, for example, through the observation that

plastid ribosomal protein mutants, which have abnormal plastid

activities, also show a downregulation of nuclear-encoded pho-

tosynthetic genes, an observation that reveals that information

about plastid functions is signaled to the nucleus. Along with

similar signals from the mitochondria, these are known as retro-

grade signals; signals from the nucleus to the organelles are

known as anterograde signals.

Other than their existence, the nature of retrograde signals

remains uncertain. A current view is that multiple signals convey

different types of information, as indicated by different tran-

scriptional responses. Furthermore, distinct signals are thought

to be involved in nuclear processes during chloroplast devel-

opment (biogenic controls) versus those involved in fine-tuning

mature chloroplast functions (operational controls) in response

to changing conditions.

Several different retrograde signals have been identified and

there isconflictingevidence for the relative importanceofdifferent

types of signals. One type of signal was first identified from a

genetic screen for an uncoupling of the nuclear and plastid ge-

nomes; the identifiedmutants are knownasgenomeuncoupled or

gun mutants, and several affect heme biosynthesis. Because

tetrapyrroles, including chlorophyll or heme or their biosynthetic

intermediates, can cause light-mediated damage, their levels are

necessarily subject to tight regulation. However, the nature of the

heme-derived signal, whether an intermediate or an indication of

flux rate, remains obscure. Another signal identified through ge-

netic approaches is PAP (3#-phosphoadenosine 5#-phosphate).PAP is synthesized in plastids andPAP levels increase inwild-type

plants exposed todrought or high light intensity. PAP is an inhibitor

of exoribonuleases and is thought to affect RNA metabolism and,

therefore, gene expression. Finally, there is evidence to support

various reactive oxygen species and even a chloroplast-envelope

bound transcription factor (which is cleaved off the plastid and

translocated to the nucleus under stress) as having roles as

retrograde signals. The nature and roles of retrograde signals

continue to be very actively investigated.

MONITORING LIGHT REACTIONS

Insights into the light-dependent reactions can be obtained

from gas exchange studies or analysis of the fluorescence/

absorbance properties of the reaction center chlorophyll and

other pigments.

Gas Exchange Analysis of Photosynthesis

Photosynthesis involves the consumption of CO2 and the pro-

duction ofO2, and achange in the concentration of these gasses

provides a convenient way to measure photosynthesis. The

measurements can be made in a chamber clamped to a leaf

or by placing an entire plant into a sealed chamber. CO2 absorbs

infrared light so its concentration is relatively easy to measure

spectroscopically, whereas oxygen can be measured using oxy-

genelectrodes.CO2measurementsarequite informativeabout the

finalprocessesofphotosynthesis, carbonfixation,butarenot ideal

for the study of light-dependent reactions as there are so many

additional steps and variables (e.g., CO2 concentration, stomatal

conductance, Rubisco activity, etc.) between carbon fixation

and the reactions that take place in the thylakoid membranes.

Furthermore, both O2 and CO2 levels are affected by mitochon-

drial and peroxisomal metabolism, so net gas exchange mea-

surements represent sums of many processes.

Spectroscopic Measurements of Light-Dependent

Reactions

Several spectroscopic measurements are available to monitor

light reactions of photosynthesis in vivo using light-adapted,

intact leaves. PSII activity can be measured by chlorophyll

fluorescence, PSI activity can be measured by the absorbance

change at 810 to;830nm, and the transthylakoid protonmotive

November 2015 11

force and proton flux can be measured by electrochromic shift

(ECS). More details are provided below.

Chlorophyll Fluorescence Measurement of PSII Activity

When a chlorophyll amolecule absorbs light, it is excited from

its ground state to its singlet excited state (Chl*), which can

return to the ground state via three processes: (1) the excita-

tion energy can be transferred to reaction centers to drive

photosynthesis through photochemistry (qP); (2) the energy

can be released as heat through NPQ; (3) the energy can be

reemitted as chlorophyll fluorescence. At room temperature,

chlorophyll fluorescence mainly originates from PSII, so chlo-

rophyll fluorescence conveys quantitative information about

PSII photochemistry.

Fluorescence yield is usually low (0.6 to 3% of total light

absorbed); however, because the fluorescence emission peaks

at longer wavelength than absorbed light, it can be measured by

illuminating a leaf with light of a defined wavelength and mea-

suring theamountof fluorescenceemissionat longerwavelength.

In order to measure fluorescence in daylight, a modulated mea-

suring system can be used, in which the light used to induce

fluorescence (known asmeasuring light) is modulated (switching

on andoff at high frequency) and thedetector is tuned todetect at

the same frequency.

First, a baseline level of fluorescence (Fo) must bemeasured in

dark-adapted leaves (in which the photosynthetic enzymes and

intermediates and nonphotochemical adaptations are all as-

sumed to be negligible), using light intensity that is too low to

initiate photochemistry. Next, the maximum yield of fluores-

cence can bemeasured. A short saturating pulse of light causes

all of the photosynthetic reaction centers to give up an electron

and become reduced, transiently preventing further photo-

chemistry; the reaction centers in this state are described as

closed. Therefore, this pulse is followed by the maximal level of

fluorescence, denoted Fm. The difference in fluorescence signal

between Fm and Fo is the variable range of fluorescence, denoted

Fv, and the ratio of Fv to Fm indicates the maximum quantum

efficiency of PSII.

Next, actinic light (light sufficient to support photosynthesis)

canbeswitchedon.Again, a rapid fluorescence response follows

(due to the closing of reaction centers), but the fluorescence

decays as the photosynthetic enzymes are activated and pho-

tochemical and nonphotochemical energy dissipation processes

ramp up. This quenching of fluorescence is due to a combination

of nonphotochemical and photochemical quenching.

Todistinguish between photochemical and nonphotochemical

quenching of chlorophyll fluorescence, a brief (#1 s) flash pulse is

used to transiently saturateandclose (reduce) all thePSII reaction

centers, resulting in zerophotochemical quenching. It is assumed

that the flash is short enoughso that no (or a negligible) increase in

the NPQ occurs and no long-term change of photosynthesis is

induced. The flash causes the chlorophyll fluorescence to reach

a maximum value (Fm#, the prime denotes the fluorescence in

light-adapted leaves) which is attained in the absence of any

photochemical quenching.

The difference between Fm (dark-adapted leaves) and Fm#(light-adapted leaves) is ameasure of NPQ, and the rate at which

maximal fluorescence is restored in the dark varies by the type of

NPQ. For example, qE reverses more quickly than qI.

The difference between Fm# (maximal fluorescence in light-

adapted leaveswhen reaction centers are closed) and the steady

state yield of fluorescence (F#, also written as Fs#) indicates the

extentof photochemistry occurring in the light (which ismeasured

here as photochemical quenching). In other words, Fm#2 F# is anindicationof theperformanceofPSII, andwhennormalized toFm#is described as the PSII operating efficiency or FPSII.

Under optimal conditions,FPSII is usually linearly proportional

to carbon fixation rate measured by gas exchange. However,

under stressful conditions (e.g., light, drought, and temperature

stress), this linear relationship often does not hold and FPSII

overestimates the rate of carbon fixation, because stress-

induced alternative electron transports (e.g., water-water cycle;

described above) can contribute to the total electron transport

throughPSII but theproduct isnotused for carbonfixation.Thus,

chlorophyll fluorescence in combination with gas exchange is

a powerful and accurate way to monitor photosynthesis under

diverse conditions.

P700 Measurement for PSI Activity

PSI photochemistry is initiated by light energy transfer from

antenna pigments associated with PSI to its reaction center

chlorophyll (P700), which absorbs light and goes to the excited

state (Chl*). The electron is then transferred from the excited

state Chl* to a downstream primary electron acceptor, eventually

producing oxidized P700, which is denoted as P7001. P7001

absorbs 810-nm light (peak at 810 to ;840 nm), while P700

does not, so the oxidation status of P700 can be determined by

absorbance at 810 nm.

To measure the percentage of oxidized P700, the maximum

oxidizable P700 pool is attained by applying a saturation flash

along with far-red light (which preferentially excites PSI), which

together ensures the complete oxidation of the P700 pool. The

P700 oxidation ratio is determined by the ratio of P7001 induced

by actinic light to the maximum amount of P7001 (induced by far

red and flash). Upon turning off the light, the reduction rate of

P7001 can be measured.

An increase in the relativepoolof reducedP700can indicate the

occurrence ofCEF. The rate ofCEF is a key acclimation to several

environmental variables; for example, plants inwhichCO2fixation

is limited tend to showan increase inCEF,which canbeobserved

by PSI activity measurements.

Electrochromic Shift to Monitor Transthylakoid Proton

Motive Force

Charge movements across the thylakoid membrane affect the

transthylakoid electric field, which in turn affects carotenoid ab-

sorbance. Thepeakchange incarotenoidabsorbanceat518nm is

knownastheECS,and itcanbeusedtomonitor thepmfacross the

membrane. The full amplitude of the ECS absorbance is propor-

tional to the total light-induced pmf, and following a dark interval,

theECSdecay rate reflectshowfastprotonsmoveoutof the lumen

through ATP synthase. ECS provides a noninvasive, direct, in situ

measurement of the thylakoid energization and deenergization,

12 The Plant Cell

transthylakoid pmf, and proton conductance of the thylakoid

membrane. Furthermore, by comparing the full amplitude of

ECS absorbance (which is proportional to pmf) in the absence

and presence of PSII inhibitors, the contribution of cyclic electron

transport to pmf can be estimated.

ECS measurements can be extended to measure the two

components of pmf. During a short dark interval (;500 ms),

ECS is dominant and measurement at 518 nm is sufficient to

measure the transthylakoid pmf; however, during longer periods,

other components that occur on the minutes timescale, like zeax-

anthin formation (absorbance change peak at 505 nm) and light

scattering (absorbance change peak at 535 nm), overlap and

interfere with the ECS. To circumvent this, multiwavelength

measurements and deconvolution are necessary to subtract

the effects of zeaxanthin and light scattering from that of the

ECS. Multiwavelength measurements can be deconvoluted so

that ECS can bemonitored during a longer dark interval (25 s) to

separate the two components of the transthylakoid pmf, DpH,

and DC.

Fluorescence Imaging

By combining light emitting diodes and sensitive digital cameras,

it is possible to incorporate spatial dimensions into fluorescent

measurements and represent these measurements as images.

These methods can be particularly useful in understanding how

photosynthetic parameters vary across a leaf. Furthermore,

although they lack the precision of fine-scale imaging systems,

it is possible to engineer systems capable of imaging fluorescence

over larger scales such as fields, which can help to automate the

screening process when breeding for enhanced resilience to

stress. In some cases, these systems rely on solar-induced fluo-

rescence,which isnotascontrolledaspulseamplitude-modulated

measurements and so more difficult to interpret.

OPTIMIZING AND IMPROVING PHOTOSYNTHESIS

Natural selection produced plants that survive in their natural

environments. We depend on plants or algae that produce high

yields of specific products in what may be very unnatural envi-

ronments. Given that plants in natural environments are often

growth-limited by water or nutrients rather than sunlight, it is

possible that they are not genetically optimized for high photo-

synthetic capacity. Investigations into improving light-dependent

reactions can be summarized as: (1) harvesting otherwavelengths

of light more efficiently, (2) decreasing shading, and (3) minimizing

photooxidativedamageandaccelerating repair.At thispoint,most

of these explorations are quite preliminary or theoretical, but

nevertheless interesting.

Harvest Light More Efficiently

Plants use only a narrow range of light wavelengths for photosyn-

thesis (400 to;700 nm), and green light is only weakly absorbed

(reflected/transmitted green light gives leaves their color).

Engineering plants to express phycoerythrin, a phycobiliprotein

from red algae that absorbs green light, might increase the

capture and flow of photons from the light-harvesting complex

to the reaction center chlorophyll, provided that the pigment

could be properly anchored and oriented. Another possibility

could be to introduce bacteriochlorophylls that absorb longer

wavelengths on the order of 1000 nm. However, using this

longer wavelength light would require extensive remodeling of

the photosynthetic apparatus, as it does not have enough

energy to drive reactions in the current system of oxygenic

photosynthesis.

Truncate Light-Harvesting Antenna Systems

In any photosynthetic system, shading occurs. Is it possible to

decreaseshading formoreeffective lightharvesting?Perhaps the

easiest system in which to apply this idea is a cell culture system

like those grown for the production of biofuels. Cells on the

surface may intercept too much light and need to dissipate the

energy, while at the same time those on the interior of the culture

vessel can be light-limited. In cyanobacteria cultures with trun-

cated light-harvesting antenna, overall levels of photosynthesis

were enhanced due tomore light being available to shaded cells.

Can this strategy be applied in multicellular plants? We know

that plants acclimate to a light gradient in several ways, for

example, by differences in the size of the antenna complexes

relative to the reactioncenters inupperversus lower leaves (which

is easily measureable by a change in the ratio of chlorophyll a to

chlorophyll b). It might be possible to engineer plants with sig-

nificantly decreased light absorption capacity in their upper

leaves (through both molecular and anatomical changes such

as amore vertical orientation) and similarly increased capacity for

light absorption in the lower leaves. Could photosynthetic ca-

pacity be increased by truncating antenna complexes in the

complex three-dimensional leaf canopy?

Minimizing Damage and Accelerating Repair

Currently the most feasible strategy for optimizing photosyn-

thesis may be to decrease photooxidative damage. When

excess excitation energy is not dissipated efficiently, reactive

oxygen species can be produced, and the damage caused can

be metabolically costly to repair. Several studies suggest that

damage can be minimized by engineering plants to be more

sensitive to excess excitation energy and to more effectively

control it through enhancing the xanthophyll cycle or by

reducing the PQH2 pool via CEF or chlororespiration. For

example, a protein present in cyanobacteria and bryophytes,

LHCSR, senses excess light energy and triggers NPQ. It has

been suggested that introducing this protein into vascular

plants could similarly decrease photooxidative damage and

so enhance photosynthesis. Similarly, photosynthesis could

be enhanced by increasing the rate of replacement of the D1

protein of PSII. Further explorations of these strategies will

require an analysis of the metabolic costs associated with

enhanced photoprotection and to what extent they decrease

quantum efficiency under optimal conditions.

November 2015 13

ARTIFICIAL PHOTOSYNTHESIS

The energy that reaches the earth from the sun in an hour is

equivalent to “all the energy humankind currently uses in a year”

(Barber and Tran, 2013). An early proponent of harnessing this

energywas the Italian scientistGiacomoCiamician (1912),whose

vision for a smokestack-free future sounds remarkably contem-

porary, “And if in a distant future the supply of coal becomes

completely exhausted, civilization will not be checked by that, for

life and civilization will continue as long as the sun shines!” More

than 100 years later, chemists and engineers are still working to

develop an affordable, effective, and efficient way to fuel civili-

zation with energy obtained from sunlight.

Artificial Photosynthesis for Chemical Energy Production

Thecostofconvertingsunlight toelectricalenergy isbecomingmore

affordablewith thedevelopmentof improvedphotovoltaic systems.

However, the limitationof thesesystemsis that thecurrentproduced

is difficult to store, and they are most effective when the energy is

used immediately; in other words, your solar-powered light is less

efficient after dark, which is when you need it most.

An alternative approach is to use solar energy to produce

chemical energy, which can be stored more easily and used

when and where it is most needed. This process is often de-

scribed as artificial photosynthesis. As in biological photosyn-

thesis, the process can be broken down into four steps: (1) light

harvesting, (2) using light energy to separate charge across

a membrane, (3) the oxidative step, using that charge separation

to oxidize water, and (4) the reductive step, using that charge

separation to reduce a substrate (such asH1 to H2 gas, or CO2 to

formic acid and other organic compounds). Artificial photosyn-

thetic systems can be entirely synthetic or can incorporate prop-

erties of living cells and synthetic biology.

One challenge is to develop semiconductingmaterials that can

support charge separation and the catalysis ofwater splitting and

hydrogen generation. Some materials can do this without addi-

tional electrocatalysts, or these can be provided separately in

a hybrid system.Difficulties include findingmaterials that are able

to capture sufficient energy using a broad spectrum of light (as

opposed to just themost highly energetic UV light), materials that

are abundant enough in the earth’s crust to be feasible for large-

scale production, andmaterials that are durable and persistent in

the functioning device. Some of the ideas being explored are

directly inspiredbyphotosynthesis, for example, thecoupling two

light-harvesting devises in series, one to drive water splitting and

one to drive hydrogen reduction, and the use ofMnCacomplexes

as catalysts for the water-splitting reaction.

Another approach that can be described as semisynthetic in-

corporates enzymes or even living cells into an energy production

system. As examples, hydrogenase enzymes can increase the

efficiency of hydrogen productionwhen incorporated into synthetic

systems, natural chromophores can enhance light harvesting in

synthetic systems, and hybrid systems composed of catalysts

introduced into cell culture systems can increase the efficiencies

by which light is converted to biofuels.

Alternatively, a synthetic biology approach can be used, in

which the properties and functions of living cells are altered so

they make desired products more efficiently. As an example,

Chlamydomonas can be engineered to more efficiently produce

H2 for use as fuel, and bacteria or algae or diatoms can be

metabolically engineered to increase their production yield of

useful biofuels. Regardless of the approach, the Second Law of

Thermodynamics must be followed. In any energy utilization

process, it is imperative that getting rid of entropic waste does

not exceed the value of the captured light energy. Currently none

of these strategies are highly efficient, but the goal of cheap and

abundant energy is propelling this research at a fast pace.

Rhodopsin-Based Phototropy

Finally, energy from the sun can be harnessed using systems

based on rhodopsins. It is important to note that rhodopsins are

entirely unrelated to the photosystems we have been describing

that arebasedonchlorophyll, but theyare introducedhereas they

show promise as energy-harnessing systems. Rhodopsins are

light receptorsmade up of an opsin protein and retinal, a pigment

related to vitamin A. Rhodopsins act as light sensors (they are

the photoreceptors found in animal eyes), but in prokaryotes they

can harness light energy through the process of phototropy.

Rhodopsins do not participate in electron transfer reactions

and therefore do not permit full photosynthesis. Bacteriorhodop-

sins use light energy to pumpprotons across amembrane, which

can be used for secondary transport or ATP synthesis; other

systems pump chloride. Rhodopsin systems have spread widely

throughout prokaryotes throughgene transfer andare genetically

simpler than chlorophyll-based photosynthetic systems, requir-

ing only a few genes to produce retinal and opsin. This genetic

simplicity makes them attractive targets for synthetic biology

approaches. However, rhodopsin-based systems use light less

efficiently than chlorophyll-based systems, due to both a nar-

rower light action spectrum and a lower efficiency of coupling

protons to chemical energy. Nevertheless, several efforts are

underway to harness them as light-energized pumps.

PHOTOSYMBIOSIS: PHOTOSYNTHETIC FUNGI AND

ANIMALS

Not all photosynthetic organisms are plants, algae, or bacteria.

We conclude this lesson by describing members of fungal and

animal kingdoms that have acquired the ability to live symbiot-

ically with photosynthetic organisms in a process that has been

described as photosymbiosis. A few are oddities, but others are

important contributors to Earth’s ecosystems; in fact, it is esti-

mated that 50%of theocean’sphotosynthetic output is a result of

photosymbiosis.

Lichen: Fungus1 Algae or Cyanobacteria

Lichen are widespread and provide a valuable food source for

many animals. Lichen are symbiotic organisms that consist of

a fungus (themycobiont) and an extracellular photobiont that can

becyanobacteria (in10%of lichens) orgreenalgae (in90%); a few

are tripartate and include both an algae and cyanobacterium.

14 The Plant Cell

The symbiosis can persist through vegetative propagation or be

reestablished following spore formation. Nearly 20,000 lichen

species are known. They are classified by the name of the

dominant fungal partner and have been described as the sym-

biotic phenotypeof the fungus. In thismutualistic association, the

photobiont provides reducedcarbonand themycobiont provides

water, nutrients, and a sheltered environment for the photobiont.

The whole is greater than the sum of the parts and lichen are able

to live in a wide range of environments and grow all across the

world on an amazing variety of substrates.

Photosynthetic Animals

Many types of marine invertebrates are able to serve as hosts

for photosynthetic algae, although it is not always clear if this

relationship is truly beneficial to both partners. Many marine

plankton that form the foundation of the ocean’s food web are

protists from the taxa Rhizaria, Alveolates, and Stramenopiles

that harbor microalgal symbionts.

Some of the more familiar photosynthetic animals are the

photosynthetic corals, which are simple animals in the phylum

Cnidaria. Some corals form calcium carbonate structures,

including huge reefs that provide important ecosystems for

marine organisms. The ability of corals to produce these vast

carbon-based structures is largely dependent on their mutual-

istic relations with a type of algae known as dinoflagellates or

zooplankton. These symbioses can be passed on maternally or

become established in vegetative tissues. Warming and acid-

ifying oceans can cause the symbiont to die or be expelled from

the host, resulting in bleaching and death of the coral. A recent

study indicates thatwe are currently in themidst of a global coral

die-off as a consequence of warming oceans and El Nino

weather events.

A tidal acoel flatworm, Symsagittifera roscoffensis (formerly

known as Convoluta roscoffensis), is providing a good model for

investigating photosymbiosis, in part because it lacks coral’s

calcified structures. Larvalworms ingest greenalgae (Tetraselmis

convolutae) a few days after hatching. Once ingested, the algae

divide mitotically and persist extracellularly within the worm’s

body. Adults can host 40,000 algal cells and appear bright green

(an early researcher describe them as “plant-animals”). These

organisms are providing insights not only into the biochemistry of

photosymbiosis but also how the symbiont affects the host’s

behavior; like other photosynthetic organisms, these worms

exhibit positive phototactic behavior.

Most of the known algal-animal symbioses involve inver-

tebrates, but there is one example that involves a vertebrate.

Eggs of the spotted salamander Ambystoma maculatum appear

green due to the presence of a single-celled algaOophila amblys-

tomatis (oophila means egg-loving). The algae are found only in

association with the salamanders and are thought to benefit

from the nitrogenous waste produced by the developing

embryo. The animal is thought to benefit from the oxygen

released as a consequence of photosynthesis. Although this

was thought to be an extracellular symbiosis, algae have

recently been identified living transiently within the animal

cells, the first occurrence of an intracellular endosymbiosis in

a vertebrate host.

Kleptoplastic Sea Slugs

In each of the above cases, the animals host intact unicellular

algae, in what is clearly a symbiosis of two organisms. The sea

slug Elysia chlorotica reveals a different approach. These

animals eat algae and then maintain the algal chloroplasts in

an intact and functional state within their digestive tissues, in

a process known as kleptoplasty (plastid-stealing). The stolen

plastids have been shown to remain photosynthetically active

for several months after removal from the algal cell. Although it

was once proposed that some of the algal genes had migrated

to the sea slug nucleus, this idea has largely been discredited,

so this interestingsymbiosis raises thequestionofhowaplastid

stays viable when it cannot be restored by gene products from

the nucleus.

Although salamanders and sea slugs do not contribute

much to the global energy economy, they provide novel re-

search models and novel opportunities to fuel imaginations

and ignite interest in the vitally important, life-sustaining

reactions of photosynthesis.

SUMMARY AND ONGOING RESEARCH

The question of how plants convert light energy into chemical

energy has fascinated scientists since it was first recognized.

Todaywehaveagoodunderstanding of the chemical processes

as well as the structure of the molecular machines that carry

them out. Great strides in obtaining structural information of the

photosynthetic complexes has led to atomic-level understand-

ing of the core reactions, which has been augmented by com-

parative studies of photosynthesis in organisms that diverged

from each other millions of years ago. New approaches to the

study of photosynthesis are refining our understanding of the

extremely rapid events that occur as light is captured, such as

quantum coherent energy transfer that excites physicists and

can contribute to more efficient light harvesting efficiency in

solar energy cells.

As we continue to delve more deeply into defining the core

reactions that harvest light, we also are expanding our un-

derstanding of how they respond to fluxes of light and me-

tabolism. In multicellular organisms, we have the additional

questions of how developmental processes and intercellular

signals and fluxes influence and are influenced by the light-

dependent reactions of photosynthesis. Genetic approaches

through mutants and new approaches including metabolo-

mics and systems approaches are increasingly important in

building a complete understanding that integrates processes

that span broad scales of time and space.

Opportunities are being explored for improvements in

photosynthetic efficiency, to enhance production from plants

and cultured cells. Opportunities are also being explored

for the development of alternative energy sources such as

biofuels from single-celled algae or cyanobacteria, or bio-

inspired artificial photosynthesis. Perhaps the biggest chal-

lenge for the 21st century will be to learn from plants how to

harness and use the abundant energy that the sun provides to

the Earth.

November 2015 15

Ru Zhang

Department of Plant Biology

Carnegie Institute for Science

rzhang@carnegiescience.edu

Johnna Roose

Louisiana State University

jroose@lsu.edu

Mary Williams

Features Editor, The Plant Cell

c/o Laboratory of Plant Physiology and Biophysics

University of Glasgow

mwilliams@aspb.org

ORCID ID: 0000-0003-4447-7815

RECOMMENDED READING

(This is a representative list of sources to help the reader access

ahugebodyof literature.Weapologize inadvance to thosewhose

work is not included.)

INTRODUCTION ANDOVERVIEW

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Beerling, D. (2007). The Emerald Planet. (Oxford, UK: Oxford University

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Blankenship, R.E. (2014). Molecular Mechanisms of Photosynthesis,

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millennium: historical highlights of photosynthesis research, part 3.

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highlights of photosynthesis research. Photosynth. Res. 73: 1–6.

doi:10.1023/A:1020169502548

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