Lipid Bio Synthesis

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The Plant Cell, Vol. 7, 957-970, July 1995 O 1995 American Society of Plant Physiologists

Lipid Biosynthesis

John Ohlroggeav and John Browseb

a Department of Botany and Plant Pathology, Michigan State University, East Lansing, Michigan 48824lnsti tute of Biological Chemistry, Washington State University, Pullman, Washington 99164

INTRODUCTION

Lipids are an essential constituent of all plant cells. The vegeta-tive cells of plants contain -5 to 10% lipid by dry weight, andalmost all of this weight is found in the membranes. Althougheach square centimeter of a plant leaf may contain only 0.2mg of lipid, this amount can account for -400 cm2 of mem-brane, reflecting the fact that membrane lipids are arrangedin layers ust two molecules hick (5 to 8 nm). If a leaf mesophyllcell were expanded a million times, the membranes would stillbe less than 1 cm thick. Despite heir slight dimensions, how-ever, the lipid membranes are the major barriers hat delineatethe cell and its compartments, and they form the sites wheremany essential processes occur, including the light harvest-ing and electron transport reactions of photosynthesis.

Some plant cells produce much more lipid than does a leafmesophyll cell. Lipids are the major form of carbon storagein the seeds of many plant species, consti tuting up to .u6O%of the dry weight of such seeds. Epidermal cells produce cutic-ular lipids that coat the surface of plants, providing he crucialhydrophobic barrier that prevents water loss and also forminga protection against pathogens and other environmentalstresses. In addition to the abundant cellular lipids, minoramounts of fatty acids are important as precursors o the hor-mone jasmonic acid and in acylation of certain membraneproteins.

Unlike the other major constituents of plants (proteins, car-bohydrates, and nucleic acids), lipids are defined on the basisof their physical properties rather than their common chemi-cal structure. Thus, lipids are often loosely defined as thosecompounds that are insoluble in water and that can be extractedfrom cells by nonpolar organic solvents (such as chloroform).As such, this class of compound s extremely diverse n struc-ture and actually constitutes the products of severa1 distinct

biosynthetic pathways. The most abundant types of lipid n mostcells, however, are those that derive from the fatty acid andglycerolipid biosynthetic pathway, and these lipids constitutethe major subject of this article. Other recent reviews includeOhlrogge et al. (1993b), Kinney (1994), Miquel and Browse(1994), and Topfer et al. (1995).

Other classes of lipid include many types of compoundsderived from the isoprenoid pathway. Over 25,000 differentisoprenoid-derived compounds have been described in the

To whom correspondence should be addressed.

plant kingdom, making his probably he richest store of chem-ical structures in the biosphere. Most of these compounds areconsidered secondary metabolites because they are notfound in all cells and are probably not essential o cell growth.However, the sterols, gibberellins, abscisic acid, and the phytolside chain of chlorophyll are also derived from this pathway.A recent book by Moore (1993) and articles in this issue byBartley and Scolnick (1995) and McGarvey and Croteau (1995)provide more detailed nformation on some of these other lipidclasses.

The fatty acid biosynthesis pathway is a primary metabolicpathway, because it is found in every cell of the plant and isessential o growth. nhibitors of fatty acid biosynthesis are le-thal to cells, and no mutations that block fatty acid synthesishave been isolated. The major fatty acids of plants (and mostother organisms) have a chain length of 16 or 18 carbons andcontain from one to three cis double bonds. Five fatty acids(18:1,18:2,18:3,16:0, and in some species, 16:3) make up over90% of the acyl chains of the structural glycerolipids of almostall plant membranes (Figure 1).

Fatty acids in cells are almost never found in the form offree fatty acids. Instead, their carboxyl group is esterified orotherwise modified. In membranes, almost all the fatty acidsare found esterified to glycerol; this class of lipid is termedglycerolipids. Membrane glycerolipids have fatty acids attachedto both the sn-1 and sn-2 positions of the glycerol backboneand a polar headgroup attached to the sn-3 position (Figure1). The combination of nonpolar fatty acyl chains and polarheadgroups eads to the amphipathic physical properties ofglycerolipids, which are essential to formation of membranebilayers. If all three positions on glycerol are esterified withfatty acids, a triacylglycerol structure results that is not suit-

able for membranes but instead constitutes the major formof lipid storage in seeds. The cuticular lipids, which are foundon the surface of all terrestrial plants (von Wettstein-Knowles,1993), contain cutin, which is a polymer of primarily 16- and18-carbon hydroxy fatty acids cross-linked by esterification oftheir carboxyl groups to hydroxyl groups on neighboring acylchains. Wax esters in the cuticular lipids are formed by esteri-fication of fatty acids to fatty alcohols. Finally, many fatty acidsare reduced o aldehydes and alcohols hat remain embeddedin the complex cuticular lipid matrix.

Although fatty acids are major constituents of every mem-brane n a cell and are also found outside cells in the cuticular


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958 The Plant Cell

DIACYLGLYCEROL-

Y Y Y

? Y HH-C-C-C-HEAD GROUP

I I I

RI Rz

R, = Fatty Acid:

OII

18:3-c--w. CH, LlNOLENlC11

-C

O LlNOLElCV c H 3 8:2

-\CH3

II 16:O-C CH, PALMlTlC

-

O

CH, 16:l-transA3

1 :OO

-CCH STEARIC

-PO LAR LlPlD HEAD GROUPS

GLYCOLIPIDS I PHOSPOLI PI DSI

MGDGmonogalactosyldiacylglycerol

HO-CH. H

DGDGO-CH, digalactosyl

H~ diacylglycerol

H d

PCH H HI I I +-o-P-o-c-c-N(cH,), phosphatidyl-I; ; I A choline

PE?H Y T-O-P-O-C-C-NH, phosphatidyl-d A A ethanolamine

yH ? 7 Y PGI; A H h glycerol

-O-P-O-C-C-C-OH phosphatidyl-

OH OHOHPI

OH

PSH H HI I I Isuphoquinovosyl I -O-!-O-F-F-NH2 phosphatidyl-diacylglycerol I O H COOH serine

HO III O\Hp Yq H VI -0-P-O-C-F-C-HI !A !A A b

CLcardiolipin

I ? ? II CHFCHCH~O- P=O

H

Figure 1. Structures of the Major Fatty Acids and Glycerolipids of Plant Cell Membranes.

The fatty acid and glycerolipid structures are arranged in approximate order of their abundance in plant leaves. Note that the fatty acids arereferred to by the number of carbon atoms (before the colon) and the number of double bonds (after the colon).

lipids, their major site of synthesis is within the plastid. In thisregard, the process of lipid biosynthesis n plants s fundamen-tally different from that in animals and fungi, which producefatty acids primarily in the cytosol. The plastid localization offatty acid synthesis means hat unlike animals and fungi, plantsmust have mechanisms to export fatty acids from the plastidto other sites in the cell. Furthermore, there must be mecha-nisms by which the rest of the cell controls the production andexport of fatty acids from the plastid. How the demand for fattyacids for assembly of extraplastidial lipids is communicated

to the plastid is a major unknown in plant lipid metabolism.

SUBSTRATES FO R FATTY AClD SYNTHESIS

All the carbon atoms found in a fatty acid are derived fromthe pool of acetyl-coenzyme A (COA) present in the plastid.The concentration of acetyl-COA in chloroplasts is only 30 to50 BM (Post-Beittenmiller et al., 1992), which is sufficient tosupply the needs of fatty acid synthesis or only a few seconds.

Nevertheless, acetyl-CoA pools remain relatively constant, evenwhen rates of fatty acid synthesis vary greatly, as in ight (whensynthesis is relatively high) and dark (when synthesis is low).Thus, a system must be available that rapidly produces acetyl-COA n the plastid for fatty acid production.

A major unresolved question in plant metabolism is howthis pool of acetyl-COA is generated. The most straightforwardpathway would be through the action of plastidial pyruvate de-hydrogenase (PDH) acting on pyruvate, either derived fromthe glycolytic pathway or perhaps produced as a side reac-

tion of ribulose bisphosphate carboxylase activity (Andrewsand Kane, 1991). However, this route has been questioned onsevera1 grounds. First, although PDH activities are generallyhigh in nongreen plastids, PDH activity in isolated chloroplastsof some species is insufficient to account for rates of fatty acidsynthesis (Lernmark and Gardestrom, 1994, and referencestherein). In addition, chloroplasts contain an extremely activeacetyl-COA synthetase, and free acetate has been found super-ior to pyruvate and other substrates as a precursor of fatty acidsynthesis by isolated chloroplasts (reviewed by Roughan andSlack, 1982). These considerations have led to suggestions


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Lipid Biosynthesis 959

BI BC BCCP CT MF-ACCase

- MS-ACCaseGRAMINEAE

MF-ACCase MF-ACCase

H F - A C C a u

Figure 2. The Acetyl-COA Carboxylase Reaction

(A) ACCase has three functional regions: biotin carboxylase, whichactivates COz by attaching it to biotin in an ATP-dependent reaction;biotin carboxyl carrier protein; and carboxyltransferase, which trans-fers activated COz rom the biotin carboxylase region to acetyCCoA,producing malonyl-COA.(B) Two forms of ACCase occur in plants. A multifunctional structure(MF-ACCase) has the three functional regions shown in (A) encodedin a single, large (>200 D)polypeptide. A multisubunit structure MS-ACCase) consists of three or more subunits that form a large com-plex. The MF-ACCase s believed o occur in the cytosol of dicots andin both he plastid and cytosol of graminaceous plants. The MS-ACCaseoccurs in the plastids of most other plants, ncluding all dicots exam-ined so far.

n

of a number of alternate pathways, including production of

acetyl-COA by a mitochondrial PDH followed by transport offree acetate or acetylcarnitine to the plastid. Free acetateentering plastids is activated o acetyl-COA by acetyl-COA syn-thetase, an enzyme with 5- to 15-fold higher activity than the invivo rate of fatty acid synthesis (Roughan and Ohlrogge, 1994).In addition, cytosolic malate and glucosed-phosphate havebeen proposed as precursors of the plastid acetyl-COA poolin oilseeds (Smith et al., 1992; Kang and Rawsthorne, 1994).

Thus, our understanding of how carbon moves from pho-tosynthesis into acetyl-COA is clouded by an abundance of

potential pathways. Furthermore, essentially all of the sugges-tions on the origin of plastidial acetyl-COA are based on in vitroanalyses of enzyme activities or precursor incorporations. Inno case do in vivo data address how photosynthate s metabo-lized to produce acetyl-COA for fatty acid synthesis. Because

of the central role of acetyl-COA in many metabolic pathways,it is likely that more than one pathway may contribute to main-taining the acetyl-COA pool, and which pathway is used mayvary with tissue, developmental stage, IighVdark conditions,and species.

STRUCTURE AND ROLE OF ACETYL-COACARBOXYLASE

The enzyme acetyl-COA carboxylase (ACCase) is generallyconsidered to catalyze the first reaction of the fatty acid bio-synthetic pathway-the formation of malonyl-COA rom acetyl-

COA and C02. This reaction actually takes place in two steps,which are catalyzed by a single enzyme complex, as shownin Figure 2A. In the first reaction, which is ATP dependent,C02 from HC03-) is transferred by the biotin carboxylaseportion of ACCase to a nitrogen of a biotin prosthetic groupattached o the eamino group of a lysine residue. In the secondreaction, catalyzed by the carboxyltransferase, the activatedC02 is transferred from biotin to acetyl-COA to formmalonyl-COA.

The structure of the plant ACCase has been a subject ofconsiderable confusion in the past, but recent evidence fromseveral laboratories s starting to provide new insights into theorganization of this complex key regulatory enzyme (Figure26). The confusion arose in large part because plants containdifferent forms of the enzyme, one of which easily loses activ-ity during attempts to characterize it.

It is now understood hat there are at least two different typesof ACCase structures. In one type of organization (frequentlyreferred to as prokaryotic), ACCase consists of several sepa-rate subunits assembled into a700-kD complex (Sasaki et al.,1993; Alban et al., 1994). At present, we know some detailsabout three of the subunits. The biotin carboxylase is an-50-kD polypeptide that is nuclear encoded (Shorrosh et al.,1995). The biotin carboxyl carrier protein (BCCP) is a 34- to38-kD protein that is almost certainly also nuclear encoded.A gene for a third subunit (-65 to 80 kD) has been identifiedin the plastid genome by its homology to one of the carbox-

yltransferase subunits of Escherichia coli ACCase. This is theonly component of plant lipid metabolism known to be encodedin the plastid genome. Furthermore, it may be unusual amongplastome-encoded proteins n hat its expression does not seemhighly regulated by light. Antibodies to this carboxyltransfer-ase subunit inhibit ACCase activity and coprecipitate he BCCPsubunit (Sasaki et al., 1993). This result indicates that, unlikein E. coli, the separate subunits associate n acomplex whosecomponents can be coprecipitated. At present, it is not clearif the three known subunits are sufficient to produce an active


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960 The Plant Cell

ACCase complex. Most likely, other components of the ACCasecomplex exist that have yet to be characterized, because evendimers of the described subunits do not add up to a 700-kDcomplex. The 700-kD complex remains associated during gelfiltration experiments, but attempts by several groups to purify

it to homogeneity have resulted n the loss of activity, presum-ably due to dissociation of the subunits. An intensive researcheffort is currently under way to characterize urther the struc-ture of this complex.

In the second type of ACCase organization, the threecomponents of the reaction are present on a single large mul-tifunctional polypeptide. This structure is termed eukaryoticbecause it is similar to that found in the cytosol of yeast andanimals. Severa1 genes and cDNA clones have been isolatedfor this type of ACCase from plants, animals, and fungi, allof which encode proteins with the biotin carboxylase domainat the N terminus, the BCCP domain in the middle, and thecarboxyltransferase at the C terminus.

The two ACCase isozymes have several important differ-ences in their biochemical properties. The multifunctionalenzyme has a much lower K,,, for acetyl-COA than the mul-tisubunit complex and has the ability to carboxylatepropionyl-CoA at substantial rates, which the multisubunit com-plex does not. In addition, the multifunctional enzyme issensitive to several important herbicides of the aryloxyphenoxy-propionic acid and cyclohexane-l,9dione classes that haveno effect on the multisubunit ACCase.

Which type of ACCase structure is present depends on itssubcellular ocalization and the typeof plant. Dicots have bothtypes of enzyme. The prokaryotic multisubunit form is foundin plastids, whereas the eukaryotic multifunctional polypep-tide structure occurs outside the plastids, most likely in the

cytosol. The plastidial isozyme of ACCase is involved primar-ily, if not exclusively, in supplying malonyl-COA or de novo fattyacid synthesis. A second isozyme of ACCase is presumablyneeded in the cytosol to supply malonyl-COA for a variety ofpathways, including fatty acid elongation for cuticular lipidproduction and flavonoid biosynthesis. 60th pathways arefound primarily in leaf epidermal cells, and the epidermis isindeed the main location of the multifunctional ACCase inleaves (Alban et al., 1994). In addition, the elongation of oleicacid (C18) to erucic acid (C22) s a major malonyl-COA-depen-dent pathway in some oilseeds, such as Brassica napos. Thiselongation OCCUIS outside the plastid and presumably dependson the cytosolic ACCase isozyme. Finally, substantial concen-trations of malonic acid that may derive from a cytosolicACCase isozyme occur in the leaf and root of soybean (Stumpfand Burris, 1981). Although a cytosolic location Seems mostreasonable for the multifunctional ACCases that have beencloned from dicots (Shorrosh et al., 1994), such a location hasyet to be demonstrated directly.

Although the evidence is still fragmentary, many monocotsshare with dicots the occurrence and localization of the twotypes of ACCase. However, the Gramineae family of plants isdifferent n that both the plastid and cytosolic ACCase isozymesare large multifunctional polypeptides Egli et al., 1993; Konishi

and Sasaki, 1994). Coincident with this evolutionary difference,the chloroplast genomes of rice and maize have lost the genethat encodes the putative carboxyltransferase subunit of theprokaryotic-type ACCase. The difference in ACCase organi-zation in the Gramineae has now provided an explanation for

the action of the grass-specific herbicides, which inhibit onlythe eukaryotic form of the enzyme (Konishi and Sasaki, 1994).Although both the cytosolic and plastid eukaryotic ACCasesare inhibited by these herbicides, the plastid form in theGramineae s much more sensitive than is the cytosolic form,and the plastid fatty acid synthesis pathway is more essentialto growth than are the secondary pathways dependent uponthe cytosolic ACCase.

REGULATION OF FATTY AClD SYNTHESIS

In animals, yeast, E. coli, and plants, ACCase is a regulatoryenzyme that controls, at least in part, the rate of fatty acid syn-thesis. Light/dark regulation of ACCase activity is responsiblefor the light/dark modulation of fatty acid synthesis rates ofspinach leaves (Post-Beittenmiller et al., 1991, 1992). In addi-tion, fatty acid synthesis n tobacco suspension cells is subjectto feedback inhibit ion by lipids provided exogenously in themedia, and this feedback appears to act at the leve1 of ACCaseactivity (Shintani and Ohlrogge, 1995). Although the regula-tory role of ACCase is well established in some tissues, severalimportant questions remain about how flux through the fattyacid synthesis pathway is controlled.

(1) What regulates ACCase activity? Although ACCase ac- ,tivity may determine the rate of fatty acid synthesis,

understanding the regulation of lipid metabolism requires anunderstanding of what factors control ACCase. In animals andfungi, ACCase is regulated by several biochemical mecha-nisms, including phosphorylation, activation by citrate, andfeedback inhibition by acyl-COA. None of these mechanismshas yet been shown to occur in plants; nevertheless, clearlybiochemical regulation occurs. The rate of fatty acid synthe-sis in leaves is six-fold higher in the light than in the dark.Although part of the light/dark control in vivo is likely to arisefrom alterations in cofactor supply, ACCase rapidly extractedfrom light-incubated chloroplasts is two- to fourfold more ac-tive than that from dark-incubated chloroplasts, even when invitro conditions and cofactors are identical (Ohlrogge et al.,1993a). At present, we have no explanation for this differencein activity.

(2) What other enzymes control he flux of fatty acid synthe-sis? ACCase may be only one of a number of enzymes thatcan be considered rate limiting. The condensing enzymes (seelater discussion), in particular, 3-ketoacyl-ACP synthase I I I(KAS III),are also logical control points. In some metabolic path-ways, control is spread over several regulatory enzymes, andthe flux control coefficient of each varies with the conditions.Now that clones are available for ACCase and most of the otherenzymes of fatty acid synthesis, transgenic plant experiments


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CH,-CB/ cetyl:EiA Acetyl-COACarboxylase

Malonvl-ACP MalonvCCoA

OCondensationCh-CHz- CH2-C-S-ACP

Butyryl-ACP

Reduction ofdouble bond

CH3-C- CH2-C -S-ACP::3-Ketobutyryl-ACP

CO2

NADPH + H+

R-C- CH2-C-S-ACP

Reduction 6of 3-Ketoacyl-ACP

3-keto group

continuesOCH3-C- CH2-C-S-ACP

IOH

3-Hydroxybutyryl-ACP OC%-CH=CH- C-S-ACP

trans-A*-Butenoyl-ACP\

Dehydration

Figure 3. The Reactions of Saturated Fatty Acid Biosynthesis.

Acetyl-COA s the basic building block of the fatty acid chain and enters the pathway both as a substrate for acetyl-COA carboxylase (reaction1) and as a primer for the initial condensation reaction (reaction 3). Reaction 2, catalyzed by malonyl-CoA:ACP transacylase, transfers malonyl

from COA o form malonyl-ACP, which is the carbon donor for allsubsequent elongation reactions. After each condensation, the 3-ketoacyl-ACPproduct is reduced (reaction 4), dehydrated (reaction 5), and reduced again (reaction 6), by 3-ketoacylACP eductase, 3-hydroxyacyl-ACP dehy-drase, and enoyl-ACP reductase, respectively.

will provide crucial in vivo tests of the role of each enzymein controlling flux through the pathway.

THE FATTY AClD SYNTHESIS PATHWAY

Plants are fundamentally different from other eukaryotes in

the molecular organization of the enzymes of fatty acid syn-thesis. Overall, to produce a 16- or 18-carbon fatty acid fromacetyl-COA and malonyl-COA, at least 30 enzymatic reactionsare required. In animals, fungi, and some bacteria, all of thesereactions are catalyzed by a multifunctional polypeptide com-plex located in the cytosol. In plants, the individual enzymesof the pathway are dissociable soluble components locatedin the stroma of plastids. Although the component enzymesof plant fatty acid synthesis can be separated easily in vitro,an intriguing question is whether they may be organized invivo into a supramolecular complex.

The central carbon donor for fatty acid synthesis is themalonyl-COA produced by ACCase. However, before enteringthe fatty acid synthesis pathway, the malonyl group is trans-ferred from COA o a protein cofactor, acyl carrier protein (ACP).From this point on, all the reactions of the pathway nvolve ACPuntil the 16- or 18-carbon product is ready for transfer toglycerolipids or export from the plastid (Figure 3). ACP is asmall(9 kD) acidic protein that contains a phosphopantethein

prosthetic group to which the growing acyl chain is attachedas a thioester. After transfer to ACP, the malonyl-thioester entersinto a series of condensation reactions with acyl-ACP (or acetyl-COA) acceptors. These reactions result in the formation of acarbon-carbon bond and in the release of the COn hat wasadded by the ACCase reaction. Removal of this C 02 helpsto drive this reaction orward, making it essentially irreversible.

At least three separate condensing enzymes (also knownas 3-ketoacyl-ACP synthases) are required to produce an 18-carbon fatty acid. The first condensation of acetyl-COA andmalonyl-ACP to form a four-carbon product is catalyzed by


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962 The Plant Cell

Prokaryotic Pathway - Plastidial compartment

Eukaryotic Pathway - Extraplastidial compartmentactivated

m,m=dgroup,i["6&2;2,OA61 COA [8:1(160)@ CTP - headgroup , PGl,m

PPi@IG3-PI m m

Figure 4. The Prokaryotic and Eukaryotic Pathways of Glycerolipid Synthesis.

The prokaryotic pathway occurs in plastids, uses acyl-ACPs as substrates, and esterifies predominantly palmitate 16:O) at position 2 f glycerol.The eukaryotic pathway occurs outside the plastid (primarily at the ER), uses acyl-CoAs as substrates, and positions 18-carbon atty acids atposition 2 f glycerol-3-phosphate. The amount of lipid synthesized by the prokaryotic pathways varies in angiosperms from 5 to 40% dependingon the plant species and the tissue. Reactions 1 and 2 re catalyzed by glycerol-9phosphate acyltransferase and lysophosphatidic acid acyltrans-ferase, respectively. CDP-DG, cytidine diphosphate-diacylglycerol; DAG, diacylglycerol; DGDG, digalactosyldiacylglycerol; GSI? glycerol3phosphate;LPA, monoacylglycerol-Sphosphate; MGDG, monogalactosyldiacylglycerol; PA, phosphatidic acid; PC, phosphatidylcholine; PE, phosphatidylethanola-mine; PG, phosphatidylglycerol; PI, phosphatidylinositol; SL, sulfoguinovosyldiacylglycerol.

KAS III(Jaworski et al., 1989). A second condensing enzyme,KAS I, s believed responsible for producing chain lengths romsix to 16 carbons. Finally, elongation of the 16carbon palmitoyl-ACP to stearoyl-ACP requires a separate condensing enzyme,KAS II. The initial product of each condensation reaction isa 3-ketoacyl-ACP. Three additional reactions occur after eachcondensation to form a saturated fatty acid (Figure 3). The3-ketoacyl-ACP s reduced at the carbonyl group by the en-zyme 3-ketoacyl-ACP reductase, which uses NADPH as theelectron donor. The next reaction is dehydration by hydroxyacyl-ACP dehydratase. Each round of fatty acid synthesis is thencompleted by the enzyme enoyl-ACP reductase, which usesNADH or NADPH to reduce the trans-2 double bond to forma saturated atty acid. The combined action of these four reac-tions leads to the lengthening of the precursor fatty acid bytwo carbons while it is still attached to ACP as a thioester.

The fatty acid biosynthesis pathway produces saturated fattyacids, but in most plant tissues, over 75% of the fatty acidsare unsaturated. The first double bond is introduced by thesoluble enzyme stearoyl-ACP desaturase. This fatty acid

desaturase is unique to the plant kingdom in that all otherknown desaturases are integral membrane proteins. The clon-ing of this soluble enzyme has recently led to its crystallizationand the determination of its three-dimensional structure by x-raycrystallography (J. Shanklin, personal communication). Thisand other structural studies have led to the first detailed in-sights into the mechanism of fatty acid desaturation and thenature of the active site. The enzyme is a homodimer in whicheach monomer has an independent active site consisting ofa diiron-oxo cluster. The two iron atoms are coordinated withina central four helix bundle in which the motif (D/E)-E-X-R-His represented in two of the four helices. During the reaction,

the reduced iron center binds oxygen and a high valent iron-oxygen complex likely abstracts hydrogen from the C-H bond(Fox et al., 1993).

The elongation of fatty acids in the plastids is terminatedwhen the acyl group is removed from ACP This can happenin two ways. In most cases, an acyl-ACP thioesterase hydro-lyzes the acyl-ACP and releases free fatty acid. Alternatively,one of two acyltransferases in the plastid transfers the fattyacid from ACP to glycerol-3-phosphate or to monoacylglycerol-3-phosphate. The first of these acyltransferases is a solubleenzyme that prefers oleoyl-ACP as a substrate. The secondacyltransferase resides on the inner chloroplast envelopemembrane and preferentially selects palmitoyl-ACP Whetherthe fatty acid is released from ACP by a thioesterase or anacyltransferase determines whether it leaves the plastid. If athioesterase acts on acyl-ACP, then the free fatty acid is ableto leave the plastid. It is not known how free fatty acids aretransported out of the plastid, but it may occur by simple diffu-sion across the envelope membrane. On the outer membraneof the chloroplast envelope, an acyl-COA synthetase s thought

to assemble an acyl-COA thioester that is then available foracyltransferase reactions o form glycerolipids in the endoplas-mic reticulum (ER). How the acyl-COA moves from the outerchloroplast envelope to the ER is also unknown, but it mayinvolve acyl-COA binding proteins, small abundant proteins re-cently found to be present in plants (Hills et al., 1994).

GLYCEROLIPID SYNTHESIS

The major fate of 16:O and 18:l acyl chains produced in theplastid is to form the hydrophobic portion of glycerolipid


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Lipid Biosynthesis 963 ,

molecules, which are components of all cellular membranes.The first steps of glycerolipid synthesis are two acylation reac-tions that transfer fatty acids to glycerol-3-phosphate to formphosphatidic acid (PA; Figure 4). Diacylglycerol (DAG) isproduced from PA by a specific phosphatase; alternatively, a

nucleotide-activated orm of DAG (CDP-DAG) is produced fromthe reaction of PA with cytidine 5'4riphosphate (CTP). Theenergy to drive attachment of the polar headgroup during denovo glycerolipid synthesis is provided by nucleotide activa-tion. When DAG is the lipid substrate, it is the headgroup thatis activated. Thus, cytidine 5'diphosphate (CDP)choline, CDP-ethanolamine, and CDP-methylethanolamine an be substratesfor phospholipid synthesis, and UDP-galactose and UDP-sulfo-quinovose are substrates for monogalactosyldiacylglycerol(MGDG) and sulfoquinovosyldiacylglycerol (SL) synthesis,respectively. Conversely, when CDP-DAG s the lipid substrate,reactions with myo-inositol, serine, and glycerol-3-phosphateresult in formation of the phospholipids phosphatidylinositol(Pl), phosphatidylserine (PS), and phosphatidyl glycerolphosphate (the precursor of PG), respectively. Digalactosyl-diacylglycerol (DGDG) is synthesized from MGDG (Joyard etal., 1993).

Although the synthetic pathways are presented here as alinear series of simple enzymatic steps, the actual biochemis-try involved is complicated by the possibilities of headgroupmodification (for example, the synthesis of phosphatidylcho-line [PC] from phosphatidylmethylethanolamine PE] by tworounds of methylation) and headgroup exchange. The detailsof the reactions involved and the synthetic routes that proba-bly operate in higher plants have been reviewed previously(Browse and Somerville, 1991; Joyard et al., 1993; Kinney,1993).

Two Pathways for Membrane LipidSynthesis

As outlined in Figure 4, higher plants possess wo distinct path-ways for the synthesis of glycerolipids: he prokaryotic pathwayof the chloroplast inner envelope, and the eukaryotic pathway,which begins with phosphatidic acid (PA) synthesis in he ER.Because of the specificities of the plastid acyltransferases orcertain acyl-ACP substrates (Frentzen, 1993), the PA made bythe prokaryotic pathway has 16:O at the sn-2 position and, inmost cases, 18:l at the sn-1 position. This PA is used for thesynthesis of PG or is converted to DAG by a PA-phosphataselocated in the inner plastid envelope. This DAG pool can actas a precursor for the synthesis of the other major plastid mem-brane lipids, MGDG, DGDG, and SL (Joyard et al., 1993). Incontrast with the plastid isozymes, the ER acyltransferasesuse acyl-COA substrates to produce PA that is highly enrichedin 18-carbon atty acids at the sn-2 position; 16:0, when pres-ent, is confined to the sn-1 position. This PA gives rise tophospholipids such as PC, PE, and PI, which are characteris-tic of the various extrachloroplast membranes. In addition,however, the DAG moiety of PC can be returned to the chloro-plast envelope, where it enters the DAG pool and contributes

to the synthesis of plastid lipids (Figure 5). Evidence fromsevera1 Arabidopsis mutants indicates that lipid exchangebetween the ER and the chloroplast is reversible to some ex-tent (Miquel and Browse, 1992; Browse et al., 1993) becauseextra chloroplastic membranes in mutants deficient in ER

desaturases see ater discussion) contain polyunsaturated attyacids derived from the chloroplasts.

In many species of higher plants, PG is the only productof the prokaryotic pathway, and the remaining chloroplast ipidsare synthesized entirely by the eukaryotic pathway. In otherspecies, including Arabidopsis and spinach, both pathwayscontribute about equally to the synthesis of MGDG, DGDG,and SL (Browse and Somerville, 1991), and the leaf lipids ofsuch plants characteristically contain substantial amounts ofhexadecatrienoic acid (16:3), which is found only in MGDGand DGDG molecules produced by the prokaryotic pathway.These plants have been termed 16:3 plants to distinguish themfrom the other angiosperms (18:3 plants), whose galactolipidscontain predominantly linolenate. The contribution of the eu-karyotic pathway to MGDG, DGDG, and SL synthesis is reducedin lower plants, and in many green algae the chloroplast isalmost entirely autonomous with respect to membrane ipid syn-thesis. One problem presented by the two-pathway model isthe need to move hydrophobic lipid molecules from the ER toother sites, particularly the chloroplast. Until recently, a class ofsoluble proteins characterized (from in vitro experiments) aslipid ransfer proteins had been considered o be the intracellulartransporters. However, biochemical and immunohistochemicalevidence has made it clear that these proteins are extracellu-lar and therefore cannot fulfill this proposed role (Sterk et al.,1991; Thoma et al., 1993).

Membrane Desaturases

In all plant tissues, the major glycerolipids are first synthesizedusing only 16:O and 18:l acyl groups. Subsequent desatura-tion of the lipids to the highly unsaturated forms typical of themembranes of plant cells (Figure 1) is carried out by membrane-bound desaturases of the chloroplast and the ER (Browseand Somerville, 1991; Heinz, 1993). lnvestigation of these de-saturases by traditional biochemical approaches has beenlimited because solubilizing and purifying them have provenvery difficult (Schmidt et al., 1994). Our understanding of themechanisms and regulation of the chloroplast and ER desatur-ases has benefited considerably from the characterization ofseven classes of Arabidopsis mutants, each deficient in a speci-fic desaturation step (Browse and Somerville, 1991; Somervilleand Browse, 1991). The loci defined by four of these classeswere originally called fadA, fad6, fadC, and fadD (forfatty acid- esaturation), but these have now been renamed fad4, fad5,fad6, and fadi: respectively. Mutations in two oci, fad2 andfad3, primarily affect desaturation of the extrachloroplast ipids,whereas mutations in the remaining five loci, fad4, fad5, fad6,fadi: and fad8, affect chloroplast lipid desaturation (Figure 5).


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964 The Plant Cell

ENDOPLASMIC RETICULUM

PI, PG COPDAG

t tr o d l el a

fad2 fad3PA

G3 P (1 : O ) (16:O) (16:O) (16:O) (16:O)16:O-COA 18: I -COA

A

718:l 16:O

18 :Z t16:lI1 6 : O )

18:3t16:1

PA

16:O 18:l 16:O-I--*

MGDG-+-

18:l 1 6 : O 18:l 1 6 : O

78:l 1 6 : O

18:l 16:l

1 1

1

18:2 16:2 18:2 16:O 18:Z 16:O1 1 1 1 ad7 .

E2

DAGlr8:2 18:2

-f fad6T fad8 T T T T7 - 78:3 16:3 18:3 16:O 18:3 1 6 : O 18:3 18:3 18:3 18:3 1 6 : O 18:3

T f T- - -(16:O)

PLASTID

Figure 5. An Abbreviated Diagram of Fatty Acid Synthesis and Glycerolipid Assembly in Arabidopsis Leaves.

Widths of the lines show the relative fluxes through different reactions. The breaks indicate the putative enzyme deficiencies in various mutants.Adapted from Browse and Somerville(1991) and used with permission of Annual Reviews. Abbreviations or lipid structures are defined in Figures1 and 4. ela, enhanced linolenate accumulation; mdl, reduced oleate desaturation.

Two of the chloroplast desaturases are highly substrate spe-cific. The fad4 gene product controls a A3 desaturase thatinserts a trans double bond into the 16:O esterified to positionsn-2 of PG, whereas the fad5 gene product is responsible orthe synthesis of A7 16:l on MGDG and possibly DGDG (Browseet al., 1985; Kunst et al., 1989a). In contrast, the other twochlo-roplast desaturases act on acyl chains with no apparentspecificity for the length of the fatty acid chain (16 or 18 car-bon), its point of attachment to the glycerol backbone (sn-1o1 sn-2), or the nature of the lipid headgroup. The 16:1/18:1desaturase s encoded by the fad6gene (Browse et al., 1989),

whereas two 16:2/18:2 isozymes are encoded by fadiand fad8(Browse et al., 1986; Gibson et al., 1994; McConn et al., 1994).The ER 18:l (fad2) and 18:2 (fad3) desaturases (Miquel andBrowse, 1992; Browse et al., 1993) also act on fatty acids atboth the sn-1 and sn-2 positions of the molecule. These en-zymes have been characterized as PC desaturases, but it ispossible that they act on other phospholipids as well.

The overall leve1 of glycerolipid unsaturation s also affectedby mutations that control other steps of fatty acid biosynthe-sis. For instance, the fabl and fab2 (forfattyacid biosynthesis)mutants of Arabidopsis are characterized by increased levels

of 16:O or 18:0, respectively, in seed and leaf tissues (Jamesand Dooner, 1990; Lightner et al., 1994a; Wu et al., 1994). Inthe fabl mutant, the biochemical defect appears to be a reduc-tion in the activity of the condensing enzyme (KAS II)responsible or the elongation of 16:O to 18:O. In fab2, it is as-sumed that 18:O-ACP desaturase activity is reduced. In bothmutants, saturated fatty acids are incorporated nto all the majormembrane glycerolipids (although MGDG contains relativelylow proportions of them). 60th mutants appear to be leaky,so the changes in overall membrane fatty acid compositionare moderately small. Nevertheless, the changes have pro-

found effects on the growth and development of these plants(see later discussion).

Characterization and cloning of the membrane-bound attyacid desaturases were for many years unattainable and frus-trating goals. However, solation of most of the genes encodingmembrane-bound desaturases has been possible due to re-cent advances in molecular biology and genetics, especiallywith the creation of T-DNA tagged mutants of Arabidopsis(Feldmann et al., 1989). This T-DNA tagging method allowedFADP to be cloned (Okuley et al., 1994). fAD3 was isolatedby both map-based chromosome walking (Arondel et al., 1992)


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and T-DNA tagging. The FADG (Hitz et al., 1994), FAD7 (Ibaet al., 1993; Yadav et al., 1993), and FADB(Gibson et al., 1994)genes were all cloned based on their homology to FADP andFAD8 The homology of these genes with gene products romother organisms hat have been characterized at the biochem-

ical level indicates that the FAD loci represent the structuralgenes for desaturases rather than proteins hat control he de-saturation reactions in any other way. The power of theapproaches available to Arabidopsis researchers is evidencedby the fact that clones are available now for more than six differ-ent desaturases, whereas research in animal lipid metabolismhas yielded only a single, membrane-bound desaturase clone.

Regulation of Membrane Lipid Synthesis

Analyses of Several of the Arabidopsis lipid mutants (Browseand Somerville, 1991), particularly act7, demonstrate thatpowerful mechanisms mediate the regulation of glycerolipidbiosynthesis o meet the demands for particular ipid molecu-lar structures for optimal membrane function. For example, act7mutants are deficient in activity of the chloroplast acylACPsn-glycerol-3-phosphate acyltransferase - he first enzyme ofthe prokaryotic pathway (Figure 5). This deficiency s compen-sated for by increased synthesis of chloroplast lipids via theeukaryotic pathway (Kunst et al., 1988). There remain the fun-damental questions of precisely what signals communicatethis demand o the lipid biosynthetic machinery and what mech-anism transmits the signal from the many different membranesof the cell to the major sites of lipid and fatty acid synthesisin the ER and the chloroplast envelope and stroma. Currently,no aspect of these regulatory mechanisms is understood n

higher plants, so this area represents a challenging and poten-tially rewarding subject for future research.

FURTHER METABOLISM OF MEMBRANEGLYCEROLIPIDS

Little is known about the cell biology of lipid turnover anddegradation. The enzymes involved, ncluding phospholipases,galactolipases, and lipoxygenases, have been thoroughly in-vestigated for a number of years (Galliard, 1980; Vick, 1993;Wang, 1993). However, it is not at all clear how these and otherenzymes are regulated during the senescence of an organ.For example, up to half of the lipid content of a leaf is metabo-lized during senescence, presumably through P-oxidation ofthe fatty acyl chains. Lipids are probably not transported outof an organ; therefore, the whole process of lipid turnover mustbe controlled to bring about the orderly dismantling of cellsand organelles and to facilitate the export of those lipid break-down products that can be used as precursors elsewhere inthe plant.

The further metabolism of membrane lipids can resultnot only in nonspecific breakdown products but also in the

production of specific cellular signals. One of these is phos-phatidylinositol 4,5-bisphosphate, whose presence in higherplants has been established (Cote and Crain, 1993). By anal-ogy to the well-studied phosphoinositide system in animal cells(Majerus, 1992), his product may act as a precursor or signal

molecules that is, DAG and inositol 1,4,5-trisphosphate, or IP3).The action of IP3 in releasing Ca2+ nto the cytoplasm andthereby regulating cellular processes has been demonstratedin severa1 plant systems (Alexandre and Lassalles, 1992; Coteand Crain, 1993). On the other hand, it is not at all certain thatthe DAG arm of the phosphoinositide signaling pathway is pres-ent in plants (Cote and Crain, 1993).

Another specific signal is asmonate, a plant growth regula-tor derived from 18:3. Recent investigations of jasmonate actionhave revealed that at very low concentrations it is able tostrongly induce specific genes, including proteinase inhibitorsand other plant defense genes (Farmer and Ryan, 1990;Gundlach et al., 1992) and vegetative storage proteins (Masonand Mullet, 1990). The structure and biosynthesis of jasmonatehave intrigued plant biologists because of their parallels tosome eicosanoids central to inflammatory responses and otherphysiological processes n mammals (Creelman et al., 1992).In plants, jasmonate is synthesized from linolenate which pre-sumably is released from membrane ipids by a phospholipaseA2) by a pathway that starts with the action of lipoxygenaseon 18:3 (Figure 5). The proposed pathway has not been fullycharacterized at the level of purified enzymes. Several lipoxy-genase isozymes have been purified and thoroughly studied(Vick, 1993), but it is not yet clear which of these acts in jas-monate synthesis. Allene oxide synthase has also been purifiedand the corresponding genes cloned. 60th the biochemistryand cell biology of jasmonate synthesis are now being given

more attention so that we can look forward to having a morecomplete picture of this extremely potent lipid-derived signalmolecule.

PHYSIOLOGY AND CELL BIOLOGY OF MEMBRANELlPlD COMPOSITION

The central issue with respect to the role of glycerolipids inmembranes is framed by the observations that each membraneof the cell has a characteristic and distinct complement ofglycerolipid types and that, within a single membrane, eachclass of lipids has a distinct fatty acid composition (Browseet al., 1993). Furthermore, he composition of each membraneand lipid class is largely conserved hroughout the plant king-dom. This diversity and its conservation hroughout evolutionimply that differences n lipid structure are important or mem-brane unction. However, despite considerable effort, the detailsof this structure/function relationship have remained elusive,in large part because there have been no instructive biologi-cal examples available n which changes in lipid compositionhave been clearly shown to have specific effects on membraneprocesses.


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966 The Plant Cell

Figure 6. Som e Phenotypic Consequences of Changes in MembraneLipidComposition.

(A )Compared with wild-type plants (left),the Arabidopsis fade mutant (right) becomes chlorotic after3 weeks at 5C.(B ) fed2 A rabidopsis plantsdie after 7 weeks at 6C.(C) Compared with wild-type plants (left),the f ab l mutant (right)is unaffectedby up to 1week of exposureto chillingat 2C.(D ) After4 weeksat 2C, however,fabl plants (right) show clear symptomsof chlorosisand reduced growth.(E) Thefab2 mutant (right)is a severe dwarfat 22C compared withthe wild type (left),as a result of an increase in stearic acid (18:0)in itsmembrane lipids.

In the last few years, investigationsof several Arabidopsislipid m utants have begunto provide suc h examp les(Hugly et

al., 1989; Kunst etal., 1989b; Hugly andSomerville,1992).The fad2 mutants of Arabidopsis are deficient in the activityof the ER 18:1 desaturase resp onsiblefo rproductionof poly-unsaturated lipidson the eukaryotic pathwayof lipid synthesis(Miqueland B row se, 1992). When gro w n at 22C,fad2 plantsare similarin growthand appearance to the w ild type. However,these m utants show a dramatic chilling-sensitive phenotypew hen plants are transferred to6C (Miquel etal., 1993). A t thistemperature, w ild-type A rabidopsis co ntinues to grow and de-velop normally,but the leaves of three allelic fad2 mutantsgradually deteriorate, displaying patchesof necrosis and ex-tensive accumulationofanthocyanins. Deathof the leaves andof the whole plants eventually follows(Figure 6). Therefore,even though the lesions at thefad2 locus are largely irrele-vant to thegrowthand survivalofplants at norm al tem peratures,polyunsaturated membranesare essential fo rmaintaining cel-lular func tion and plant viability at low tem peratures. A lthoughthe deficiency inpolyunsatu rated lipids has severe conse-quences , sev eral lines of ev idence indicate that the c hange(s)in membrane functionin fad2 plants at 6C begins to c o m -prom ise cell viability onlyafter several days (M iquelet al., 1993).This gradual developm ent of sym ptom s at6C argues againsta general collapse of membrane integrityas the cause of thelethal phenotype. Rather, it indicates that the decrease in poly-unsaturated membrane lipids may initially have relatively

limited effects in disrupting cellular function.

Additional Arabidopsis mutants have helped to clarify therole of lipid composi t ionin chilling sensitivity,a phenomenon

of considerable agricultural significance. The increased 16:0content in leaves of f a b l plants includes the synthesis of mo-lecular s pecies of chloroplast PG that contain o nly 1 6:0 plus16:1-frans plus 1 8:0, and these m olecular species acco unt for42% of the PG in leaves of the mutant. Such"high meltingpoint" molecular speciesof PG have been correlated withchilling se nsitivity of plants (M urata, 1983). Ex perim ents withtransgenic plants engineered to express increased levelsof high-melting-point PG have also been interpretedas sup-porting a major rolefo r high-melting-point PG species indetermining plant chilling s ensitivity (M urata etal., 1992; Wo lteretal. , 1992). However,f a b l m utants are able to growand com -plete their life cy clesnormally at 10C. They are also unaffected(as com pared with wild-type c ontrols) by m ore severechillingtreatments that lead q uickly to the death of c ucum ber and otherchilling-sensitive plants. These treatments include exposureto 2C for up to 7days and freezingto -2C for 24 hr (Wuand Browse, 1995). Following these treatments, mutant plantsreturned to 22C remain indistinguishable from wild-type co n-trols (Figure 6). If grown continuouslyat 2C, fab l plantseventually show damage (Figure 6; Wu and Browse, 1995),but the f a b l m utant does not sho w classicchilling sensitivity,despite its increased 16:0 content. This suggests thathigh-melting-point species of PG cannot alone account for chillingsensitivityof plants.

The fab2 m utant, w hich co ntains elevated levels of 18:0 in


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membrane ipids, shows an extreme dwarf growth habit, withmutant plants growing to less than one-tenth the size of thewild type (Figure 6). Many cell types in the mutant, includingleaf mesophyll and epidermal cells, fail to expand, whereasothers, including stomatal guard cells and leaf trichomes, de-velop to the normal size. The relationship between the dwarfand high 18:O phenotypes was established by identifying a mu-tation, a suppressor of high stearate, that causes the fab2mutant to grow to normal size and to have normal levels of18:O. Moreover, when fab2 plants are grown at high tempera-tures, which ameliorates the effects of increased saturationof membrane lipids, tissue 18:O levels remain high but plantgrowth is more normal (Lightner et al., 1994b). This result sug-gests that increased saturation of membrane ipids is the directcause of the dwarf phenotype of fab2 mutants.

BIOSYNTHESIS OF STORAGE TRIACYLGLYCEROLS

A plant stores reserve material (oil, protein, and carbohydrate)in its seed to allow growth of the next generation. The amountof oil in different species may vary widely, from as little as 1%to as much as 60% of the total dry weight of the seed. In al-most all plant species, he chemical orm in which oil is storedis triacylglycerol TAG). Unlike he glycerolipids ound in mem-branes, TAGs do not perform a structural role but instead serveprimarily as a storage form of carbon.

In the mature seed, TAG is stored in densely packed lipidbodies that are roughly spherical in shape, with an averagediameterof 1 pm (Huang, 1992; Murphy, 1993; Herman, 1994).This sim does not change during seed development, and ac-cumulation of oil is accompanied by an increase in the numberof lipid bodies. These organelles appear to be surrounded bya type of membrane that is only a monolayer rather than anormal bilayer membrane. The membranes of isolated lipidbodies contain both phospholipids and major characteristicproteins, ermed oleosins. However, these constituents are lessthan 5% of the weight of the oil body, with TAG constitutingby far the major component (90 to 95%).

The Pathway of Triacylglycerol Synthesis

TAG biosynthesis, at least in some species, has been proposedto take place by the comparatively straightforward Kennedypathway. As in the eukaryotic pathway for synthesis of the majormembrane glycerolipids (Figure 4), fatty acids are sequentiallytransferred rom COA o positions 1 and 2 of glycerol-3-phos-phate, resulting n the formation of the central metabolite, PA.Dephosphorylation of PA releases DAG. In the final step ofTAG synthesis, a third fatty acid is transferred to the vacantposition 3 of DAG. This step is catalyzed by diacylglycerolacyltransferase (DAGAT), the only enzymatic reaction uniqueto TAG biosynthesis.

The assembly of three fatty acids onto a glycerol backboneis not always as straightforward as outlined for the Kennedypathway, however. In many oilseeds, most fatty acids producedin the plastid are not immediately available or TAG biosynthe-sis. Instead, the acyl chains enter into phosphatidyl choline(and PE to a lesser extent), where they become desaturatedor otherwise modified. Fatty acids from PC may then becomeavailable or TAG synthesis by one of two mechanisms. In thefirst mechanism, a fatty acid attached to COA and a fatty acidon PC may essentially trade places. Such an acyl exchangeprobably occurs by the combined reverse and forward reac-tions of an acyl-CoA:PC acyltransferase (Stymne and Stobart,1987). The resulting acyl-COA may then be used as an acyldonor in TAG synthesis. The exchange reaction allows 18:lnewly produced and exported from the plastid to enter PC,while desaturated or otherwise modified fatty acids depart forTAG or other lipids. The second mechanism by which PC canparticipate in TAG synthesis is by donation of its entire DAG

portion (Stymne and Stobart, 1987). In many plants, the syn-thesis of PC from DAG and CDP-choline appears o be rapidlyreversible as catalyzed by the CDP-choline phosphotransfer-ase (Slack et al., 1983). The reversibility of this reaction allowsthe DAG moiety of PC, including any modified fatty acids, tobecome available for TAG synthesis.

Because DAG is a common precursor to both membranePC and storage TAG, identification of these enzymatic reac-tions responsible or TAG biosynthesis does not satisfactorilyaddress two potentially related questions of plant ipid metab-olism: How do plants control which fatty acids are stored intriacylglycerol and which are restricted o membranes? 1s thesynthesis of TAG spatially distinct from the synthesis of mem-brane glycerolipids?

The first question is particularly relevant to those oilseedsthat store unusual fatty acids. The fatty acid composition ofstorage oils varies much more than that of membrane glycero-lipids. More than 300 different fatty acids are known to occurin seed TAG (Harwood, 1980; van de Loo et al., 1993). Theirstructures may vary in chain length from as few as eight car-bons to >22 carbons. In addition, the position and number ofdouble bonds may be unusual, and various functional groupssuch as hydroxy or epoxy may be added to the acyl chain.The reason for this great diversity in plant storage oils is un-known, but the special physical or chemical properties of someof these fatty acids may explain why (if not how) these unusualfatty acids are excluded from membranes. If they were to be-

come major components of the membranes, then the fluidityor other physical properties of the membrane might bedisrupted.

SUMMARY AND PERSPECTIVES FOR FUTURERESEARCH

The basic metabolic pathways that lead to the synthesis of themajor plant glycerolipids are now mostly understood. Much


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968 The Plant Cell

of the research effort in plant lipid biosynthesis over the pastfive years has been directed toward obtaining clones for en-zymes in the pathway. A number of industrial laboratories haveparticipated in this effort because of the potential economicvalue that might derive from genetic engineering of vegetable

oils. Plant TAGs are a major agricultura1 commodity, with avalueof ~ $ 2 5 illion annually. Recently, severa1 successes havebeen achieved in genetically engineering oilseed fatty acidcomposition to produce new or improved vegetable oils(reviewed by Kinney, 1994; Ohlrogge, 1994; Topfer et al., 1995).Almost all of the many soluble enzymes involved inproducingoleic acid rom acetyl-COA have now been cloned (Topfer andMartini, 1994). Severa1 of the membrane-bound enzymes thatwere more difficult to obtain using biochemical approacheshave now yielded to alternative strategies, such as map-basedcloning (Arondel et al., 1992), T-DNA or transposon tagging(Okuley et al., 1994; James et al., 1995), and complementa-tion of mutants (Brown et al., 1994; Dewey et al., 1994). Mostof the already cloned genes have significant homologyto theirmicrobialor animalcounterparts. Within the near future, mostof the expressed genes of Arabidopsis and rice will be se-quenced (Newman et al., 1994; Sasaki et al., 1994), makingit likely that clones for many more of the enzymes in lipid me-tabolism can be identified by appropriate analysis of thesequence databases.

Although we now understand the basic pathways ofglycerolipid synthesis, a number of major questions remainto be answered, among them: (1) What is he source ofplastidialacetyl-COA? Does the pathway or acetyl-COA production varywith different tissues and during tissue development? 2) Whatdetermines how much lipid is produced in a cell? 1s controlexerted primarily through the leve1 of gen e expression or

through metabolic controls over enzyme activity? (3) How isthe production of fatty acids in the plastid coordinated withtheir utilization outside the plastid? Are there signals that com-municate between the ER and the plastid fatty acid synthesispathway, and, if so, what are these signals? (4) How and inwhat form do lipids produced in the ER return for incorpora-tion in plastidial membranes? (5) How do cells determine thesubcellular distribution of different lipid classes and the differ-ent fatty acid compositions associated with different polarheadgroups? Are there specific carrier proteins hat allow lipidsto move between organelles? (6) ow are lipid biosynthesisgenes regulated? Are there globaltrans-acting factors thatsimultaneously control the expression of many genes? (7)Are

membrane ipid desaturases controlled by signals that respondto membrane fluidity? (8) What roles do phospholipases andlipoxygenases play in lipid metabolism? How is the synthesisof jasmonic acid controlled? (9) Are the precursors for mem-brane lipids and storage TAGs synthesized by the sameenzymes and in the same subcellular location?

Many of these questions have been very difficult to approachin the past. The recent availability of a large number of clonesfor enzymes of lipid metabolism, together with the ability tomanipulate their expression in transgenic plants, now offersplant lipid scientists the potential to design robust new experi-ments to address these topics.

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