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 48824
`lnstitute 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, and
`almost all of this weight is found in the membranes. Although
`each square centimeter of a plant leaf may contain only 0.2
`mg of lipid, this amount can account for -400 cm2 of mem-
`brane, reflecting the fact that membrane lipids are arranged
`in layers just two molecules thick (5 to 8 nm). If a leaf mesophyll
`cell were expanded a million times, the membranes would still
`be less than 1 cm thick. Despite their slight dimensions, how-
`ever, the lipid membranes are the major barriers that delineate
`the cell and its compartments, and they form the sites where
`many essential processes occur, including the light harvest-
`ing and electron transport reactions of photosynthesis.
`Some plant cells produce much more lipid than does a leaf
`mesophyll cell. Lipids are the major form of carbon storage
`in the seeds of many plant species, constituting up to .u6O%
`of the dry weight of such seeds. Epidermal cells produce cutic-
`ular lipids that coat the surface of plants, providing the crucial
`hydrophobic barrier that prevents water loss and also forming
`a protection against pathogens and other environmental
`stresses. In addition to the abundant cellular lipids, minor
`amounts of fatty acids are important as precursors to the hor-
`mone jasmonic acid and in acylation of certain membrane
`proteins.
`Unlike the other major constituents of plants (proteins, car-
`bohydrates, and nucleic acids), lipids are defined on the basis
`of their physical properties rather than their common chemi-
`cal structure. Thus, lipids are often loosely defined as those
`compounds that are insoluble in water and that can be extracted
`from cells by nonpolar organic solvents (such as chloroform).
`As such, this class of compound is extremely diverse in struc-
`ture and actually constitutes the products of severa1 distinct
`biosynthetic pathways. The most abundant types of lipid in most
`cells, however, are those that derive from the fatty acid and
`glycerolipid biosynthetic pathway, and these lipids constitute
`the major subject of this article. Other recent reviews include
`Ohlrogge et al. (1993b), Kinney (1994), Miquel and Browse
`(1994), and Topfer et al. (1995).
`Other classes of lipid include many types of compounds
`derived from the isoprenoid pathway. Over 25,000 different
`isoprenoid-derived compounds have been described in the
`
`To whom correspondence should be addressed.
`
`plant kingdom, making this probably the richest store of chem-
`ical structures in the biosphere. Most of these compounds are
`considered “secondary” metabolites because they are not
`found in all cells and are probably not essential to cell growth.
`However, the sterols, gibberellins, abscisic acid, and the phytol
`side chain of chlorophyll are also derived from this pathway.
`A recent book by Moore (1993) and articles in this issue by
`Bartley and Scolnick (1995) and McGarvey and Croteau (1995)
`provide more detailed information on some of these other lipid
`classes.
`The fatty acid biosynthesis pathway is a primary metabolic
`pathway, because it is found in every cell of the plant and is
`essential to growth. lnhibitors of fatty acid biosynthesis are le-
`thal to cells, and no mutations that block fatty acid synthesis
`have been isolated. The major fatty acids of plants (and most
`other organisms) have a chain length of 16 or 18 carbons and
`contain 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 over
`90% of the acyl chains of the structural glycerolipids of almost
`all plant membranes (Figure 1).
`Fatty acids in cells are almost never found in the form of
`“free” fatty acids. Instead, their carboxyl group is esterified or
`otherwise modified. In membranes, almost all the fatty acids
`are found esterified to glycerol; this class of lipid is termed
`glycerolipids. Membrane glycerolipids have fatty acids attached
`to both the sn-1 and sn-2 positions of the glycerol backbone
`and a polar headgroup attached to the sn-3 position (Figure
`1). The combination of nonpolar fatty acyl chains and polar
`headgroups leads to the amphipathic physical properties of
`glycerolipids, which are essential to formation of membrane
`bilayers. If all three positions on glycerol are esterified with
`fatty acids, a “triacylglycerol” structure results that is not suit-
`able for membranes but instead constitutes the major form
`of lipid storage in seeds. The cuticular lipids, which are found
`on the surface of all terrestrial plants (von Wettstein-Knowles,
`1993), contain cutin, which is a polymer of primarily 16- and
`18-carbon hydroxy fatty acids cross-linked by esterification of
`their carboxyl groups to hydroxyl groups on neighboring acyl
`chains. Wax esters in the cuticular lipids are formed by esteri-
`fication of fatty acids to fatty alcohols. Finally, many fatty acids
`are reduced to aldehydes and alcohols that remain embedded
`in the complex cuticular lipid matrix.
`Although fatty acids are major constituents of every mem-
`brane in a cell and are also found outside cells in the cuticular
`
`CSIRO Exhibit 1014
`
`

`

`958
`
`The Plant Cell
`
`DIACYLGLYCEROL -
`-POLAR
`Y Y Y
`H-C-C-C-HEAD GROUP
`? Y H
`I
`I
` I
`RI Rz
`R, = Fatty Acid:
`
`HO-CH.
`
`ÒH
`
`18:3
`LlNOLENlC
`
` 18:2
`LlNOLElC
`
`O-CH,
`
`H~
`
`H d
`
`GLYCOLIPIDS
`
`LlPlD HEAD GROUPS
`
`PHOSPOLI PI DS
`
`I
`I
`
`-c--w.
`
`CH,
`
`c
`
`H
`
`3
`
`O
`II
`
`O
`11
`-C
`
`V
`O
`
`-‘\CH3 O
`
`-C-
`
`-C-
`
`II
`
`O
`
`O
`
`-C-CH3
`
`16:O
`PALMlTlC
`
`CH,
`
`CH,
`
`16:l-transA3
`
`1 8:O
`STEARIC
`
`HO
`
`MGDG
`monogalactosyl
`diacylglycerol
`
`PC
`OH H H
`I; ;IA
`
`+
`I I
`I
`-o-P-o-c-c-N(cH,), phosphatidyl-
`choline
`
`DGDG
`digalactosyl
`diacylglycerol
`
`?H Y T
`A A
`‘d
`-O-P-O-C-C-NH,
`
`PE
`phosphatidyl-
`ethanolamine
`
`yH ? 7 Y
`PG
`-O-P-O-C-C-C-OH phosphatidyl-
`I;
`glycerol
`A Ò H h
`
`OH OHOH
`
`PI
`
`OH
`
`OH H H
`I
`I
`I
`I
`suphoquinovosyl I -O-!-O-F-F-NH2
`diacylglycerol
`O
`H COOH
`I
`I
`I
`I O\Hp Y q H V
`
`
`
`I -0-P-O-C-F-C-H
`A b
`I !A
`I ? ?
`
`!A
`
`I
`I CHFCHCH~O- P=O
`ÒH
`
`PS
`phosphatidyl-
`serine
`
`CL
`cardiolipin
`
`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 are
`referred 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 this
`regard, the process of lipid biosynthesis in plants is fundamen-
`tally different from that in animals and fungi, which produce
`fatty acids primarily in the cytosol. The plastid localization of
`fatty acid synthesis means that unlike animals and fungi, plants
`must have mechanisms to export fatty acids from the plastid
`to other sites in the cell. Furthermore, there must be mecha-
`nisms by which the rest of the cell controls the production and
`export of fatty acids from the plastid. How the demand for fatty
`acids for assembly of extraplastidial lipids is communicated
`to the plastid is a major unknown in plant lipid metabolism.
`
`SUBSTRATES FOR FATTY AClD SYNTHESIS
`
`All the carbon atoms found in a fatty acid are derived from
`the pool of acetyl-coenzyme A (COA) present in the plastid.
`The concentration of acetyl-COA in chloroplasts is only 30 to
`50 BM (Post-Beittenmiller et al., 1992), which is sufficient to
`supply the needs of fatty acid synthesis for only a few seconds.
`
`Nevertheless, acetyl-CoA pools remain relatively constant, even
`when rates of fatty acid synthesis vary greatly, as in light (when
`synthesis is relatively high) and dark (when synthesis is low).
`Thus, a system must be available that rapidly produces acetyl-
`COA in the plastid for fatty acid production.
`A major unresolved question in plant metabolism is how
`this pool of acetyl-COA is generated. The most straightforward
`pathway would be through the action of plastidial pyruvate de-
`hydrogenase (PDH) acting on pyruvate, either derived from
`the glycolytic pathway or perhaps produced as a side reac-
`tion of ribulose bisphosphate carboxylase activity (Andrews
`and Kane, 1991). However, this route has been questioned on
`severa1 grounds. First, although PDH activities are generally
`high in nongreen plastids, PDH activity in isolated chloroplasts
`of some species is insufficient to account for rates of fatty acid
`synthesis (Lernmark and Gardestrom, 1994, and references
`therein). In addition, chloroplasts contain an extremely active
`acetyl-COA synthetase, and free acetate has been found super-
`ior to pyruvate and other substrates as a precursor of fatty acid
`synthesis by isolated chloroplasts (reviewed by Roughan and
`Slack, 1982). These considerations have led to suggestions
`
`CSIRO Exhibit 1014
`
`

`

`Lipid Biosynthesis
`
`959
`
`potential pathways. Furthermore, essentially all of the sugges-
`tions on the origin of plastidial acetyl-COA are based on in vitro
`analyses of enzyme activities or precursor incorporations. In
`no case do in vivo data address how photosynthate is 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 may
`vary with tissue, developmental stage, IighVdark conditions,
`and species.
`
`STRUCTURE AND ROLE OF ACETYL-COA
`CARBOXYLASE
`
`The enzyme acetyl-COA carboxylase (ACCase) is generally
`considered to catalyze the first reaction of the fatty acid bio-
`synthetic pathway-the formation of malonyl-COA from acetyl-
`COA and C02. This reaction actually takes place in two steps,
`which are catalyzed by a single enzyme complex, as shown
`in Figure 2A. In the first reaction, which is ATP dependent,
`C02 (from HC03-) is transferred by the biotin carboxylase
`portion of ACCase to a nitrogen of a biotin prosthetic group
`attached to the eamino group of a lysine residue. In the second
`reaction, catalyzed by the carboxyltransferase, the activated
`C02 is transferred from biotin to acetyl-COA to form
`malonyl-COA.
`The structure of the plant ACCase has been a subject of
`considerable confusion in the past, but recent evidence from
`several laboratories is starting to provide new insights into the
`organization of this complex key regulatory enzyme (Figure
`26). The confusion arose in large part because plants contain
`different forms of the enzyme, one of which easily loses activ-
`ity during attempts to characterize it.
`It is now understood that there are at least two different types
`of ACCase structures. In one type of organization (frequently
`referred 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 details
`about 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- to
`38-kD protein that is almost certainly also nuclear encoded.
`A gene for a third subunit (-65 to 80 kD) has been identified
`in the plastid genome by its homology to one of the carbox-
`yltransferase subunits of Escherichia coli ACCase. This is the
`only component of plant lipid metabolism known to be encoded
`in the plastid genome. Furthermore, it may be unusual among
`plastome-encoded proteins in that its expression does not seem
`highly regulated by light. Antibodies to this carboxyltransfer-
`ase subunit inhibit ACCase activity and coprecipitate the BCCP
`subunit (Sasaki et al., 1993). This result indicates that, unlike
`in E. coli, the separate subunits associate in acomplex whose
`components can be coprecipitated. At present, it is not clear
`if the three known subunits are sufficient to produce an active
`
`CSIRO Exhibit 1014
`
`B
`
`I
`
`BC
`
`BCCP
`
`-1
`
`CT
`
`MF-ACCase
`
`MS-ACCase
`
`MF-ACCase
`
`GRAMINEAE
`
`MF-ACCase
`
`HF-ACCau
`
`Figure 2. The Acetyl-COA Carboxylase Reaction
`(A) ACCase has three functional regions: biotin carboxylase, which
`activates COz by attaching it to biotin in an ATP-dependent reaction;
`n
`biotin carboxyl carrier protein; and carboxyltransferase, which trans-
`fers activated COz from 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) encoded
`in a single, large (>200 kD) polypeptide. A multisubunit structure (MS-
`ACCase) consists of three or more subunits that form a large com-
`plex. The MF-ACCase is believed to occur in the cytosol of dicots and
`in both the plastid and cytosol of graminaceous plants. The MS-ACCase
`occurs in the plastids of most other plants, including all dicots exam-
`ined so far.
`
`of a number of alternate pathways, including production of
`acetyl-COA by a mitochondrial PDH followed by transport of
`free acetate or acetylcarnitine to the plastid. Free acetate
`entering plastids is activated to acetyl-COA by acetyl-COA syn-
`thetase, an enzyme with 5- to 15-fold higher activity than the in
`vivo rate of fatty acid synthesis (Roughan and Ohlrogge, 1994).
`In addition, cytosolic malate and glucosed-phosphate have
`been proposed as precursors of the plastid acetyl-COA pool
`in 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
`
`

`

`960
`
`The Plant Cell
`
`ACCase complex. Most likely, other components of the ACCase
`complex exist that have yet to be characterized, because even
`dimers of the described subunits do not add up to a 700-kD
`complex. The 700-kD complex remains associated during gel
`filtration experiments, but attempts by several groups to purify
`it to homogeneity have resulted in the loss of activity, presum-
`ably due to dissociation of the subunits. An intensive research
`effort is currently under way to characterize further the struc-
`ture of this complex.
`In the second type of ACCase organization, the three
`components of the reaction are present on a single large mul-
`tifunctional polypeptide. This structure is termed eukaryotic
`because it is similar to that found in the cytosol of yeast and
`animals. Severa1 genes and cDNA clones have been isolated
`for this type of ACCase from plants, animals, and fungi, all
`of which encode proteins with the biotin carboxylase domain
`at the N terminus, the BCCP domain in the middle, and the
`carboxyltransferase at the C terminus.
`The two ACCase isozymes have several important differ-
`ences in their biochemical properties. The multifunctional
`enzyme has a much lower K,,, for acetyl-COA than the mul-
`tisubunit complex and has the ability to carboxylate
`propionyl-CoA at substantial rates, which the multisubunit com-
`plex does not. In addition, the multifunctional enzyme is
`sensitive to several important herbicides of the aryloxyphenoxy-
`propionic acid and cyclohexane-l,9dione classes that have
`no effect on the multisubunit ACCase.
`Which type of ACCase structure is present depends on its
`subcellular localization and the typeof plant. Dicots have both
`types of enzyme. The prokaryotic multisubunit form is found
`in 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 for de novo fatty
`acid synthesis. A second isozyme of ACCase is presumably
`needed in the cytosol to supply malonyl-COA for a variety of
`pathways, including fatty acid elongation for cuticular lipid
`production and flavonoid biosynthesis. 60th pathways are
`found primarily in leaf epidermal cells, and the epidermis is
`indeed the main location of the multifunctional ACCase in
`leaves (Alban et al., 1994). In addition, the elongation of oleic
`acid (C18) to erucic acid (C22) is a major malonyl-COA-depen-
`dent pathway in some oilseeds, such as Brassica napos. This
`elongation OCCUIS outside the plastid and presumably depends
`on the cytosolic ACCase isozyme. Finally, substantial concen-
`trations of malonic acid that may derive from a cytosolic
`ACCase isozyme occur in the leaf and root of soybean (Stumpf
`and Burris, 1981). Although a cytosolic location Seems most
`reasonable for the multifunctional ACCases that have been
`cloned from dicots (Shorrosh et al., 1994), such a location has
`yet to be demonstrated directly.
`Although the evidence is still fragmentary, many monocots
`share with dicots the occurrence and localization of the two
`types of ACCase. However, the Gramineae family of plants is
`different in that both the plastid and cytosolic ACCase isozymes
`are 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 gene
`that encodes the putative carboxyltransferase subunit of the
`prokaryotic-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 only
`the eukaryotic form of the enzyme (Konishi and Sasaki, 1994).
`Although both the cytosolic and plastid eukaryotic ACCases
`are inhibited by these herbicides, the plastid form in the
`Gramineae is much more sensitive than is the cytosolic form,
`and the plastid fatty acid synthesis pathway is more essential
`to growth than are the secondary pathways dependent upon
`the cytosolic ACCase.
`
`REGULATION OF FATTY AClD SYNTHESIS
`
`In animals, yeast, E. coli, and plants, ACCase is a regulatory
`enzyme that controls, at least in part, the rate of fatty acid syn-
`thesis. Light/dark regulation of ACCase activity is responsible
`for the light/dark modulation of fatty acid synthesis rates of
`spinach leaves (Post-Beittenmiller et al., 1991, 1992). In addi-
`tion, fatty acid synthesis in tobacco suspension cells is subject
`to feedback inhibition by lipids provided exogenously in the
`media, and this feedback appears to act at the leve1 of ACCase
`activity (Shintani and Ohlrogge, 1995). Although the regula-
`tory role of ACCase is well established in some tissues, several
`important questions remain about how flux through the fatty
`acid 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 an
`understanding of what factors control ACCase. In animals and
`fungi, ACCase is regulated by several biochemical mecha-
`nisms, including phosphorylation, activation by citrate, and
`feedback inhibition by acyl-COA. None of these mechanisms
`has yet been shown to occur in plants; nevertheless, clearly
`biochemical 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 arise
`from alterations in cofactor supply, ACCase rapidly extracted
`from light-incubated chloroplasts is two- to fourfold more ac-
`tive than that from dark-incubated chloroplasts, even when in
`vitro conditions and cofactors are identical (Ohlrogge et al.,
`1993a). At present, we have no explanation for this difference
`in activity.
`(2) What other enzymes control the flux of fatty acid synthe-
`sis? ACCase may be only one of a number of enzymes that
`can be considered rate limiting. The condensing enzymes (see
`later discussion), in particular, 3-ketoacyl-ACP synthase III
`(KAS III), are also logical control points. In some metabolic path-
`ways, control is spread over several regulatory enzymes, and
`the flux control coefficient of each varies with the conditions.
`Now that clones are available for ACCase and most of the other
`enzymes of fatty acid synthesis, transgenic plant experiments
`
`CSIRO Exhibit 1014
`
`

`

`CH,-CB /
`
`Acetyl:EiA
`
`Lipid Biosynthesis
`
`961
`
`Acetyl-COA
`Carboxylase
`
`Malonvl-ACP
`
`MalonvCCoA
`
`CH3-C- CH2-C -S-ACP
`
`::
`
`3-Ketobutyryl-ACP
`
`CO2
`
`NADPH + H+
`
`Condensation
`
`O
`Ch-CHz- CH2-C-S-ACP
`Butyryl-ACP
`
`Reduction
`of
`3-keto group
`
`R-C- CH2-C-S-ACP
`
`6
`3-Ketoacyl-ACP
`
`O
`CH3-C- CH2-C-S-ACP
`I
`OH
`3-Hydroxybutyryl-ACP
`
`\
`
`Reduction of
`double bond
`
`continues
`
`O
`C%-CH=CH-
`C-S-ACP
`trans-A*-Butenoyl-ACP
`
`Dehydration
`
`Figure 3. The Reactions of Saturated Fatty Acid Biosynthesis.
`Acetyl-COA is the basic building block of the fatty acid chain and enters the pathway both as a substrate for acetyl-COA carboxylase (reaction
`1) and as a primer for the initial condensation reaction (reaction 3). Reaction 2, catalyzed by malonyl-CoA:ACP transacylase, transfers malonyl
`from COA to form malonyl-ACP, which is the carbon donor for all subsequent elongation reactions. After each condensation, the 3-ketoacyl-ACP
`product is reduced (reaction 4), dehydrated (reaction 5), and reduced again (reaction 6), by 3-ketoacylACP reductase, 3-hydroxyacyl-ACP dehy-
`drase, and enoyl-ACP reductase, respectively.
`
`will provide crucial in vivo tests of the role of each enzyme
`in 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 from
`acetyl-COA and malonyl-COA, at least 30 enzymatic reactions
`are required. In animals, fungi, and some bacteria, all of these
`reactions are catalyzed by a multifunctional polypeptide com-
`plex located in the cytosol. In plants, the individual enzymes
`of the pathway are dissociable soluble components located
`in the stroma of plastids. Although the component enzymes
`of plant fatty acid synthesis can be separated easily in vitro,
`an intriguing question is whether they may be organized in
`vivo into a supramolecular complex.
`
`The central carbon donor for fatty acid synthesis is the
`malonyl-COA produced by ACCase. However, before entering
`the fatty acid synthesis pathway, the malonyl group is trans-
`ferred from COA to a protein cofactor, acyl carrier protein (ACP).
`From this point on, all the reactions of the pathway involve ACP
`until the 16- or 18-carbon product is ready for transfer to
`glycerolipids or export from the plastid (Figure 3). ACP is a
`small(9 kD) acidic protein that contains a phosphopantethein
`prosthetic group to which the growing acyl chain is attached
`as a thioester. After transfer to ACP, the malonyl-thioester enters
`into a series of condensation reactions with acyl-ACP (or acetyl-
`COA) acceptors. These reactions result in the formation of a
`carbon-carbon bond and in the release of the COn that was
`added by the ACCase reaction. Removal of this C02 helps
`to drive this reaction forward, making it essentially irreversible.
`At least three separate condensing enzymes (also known
`as 3-ketoacyl-ACP synthases) are required to produce an 18-
`carbon fatty acid. The first condensation of acetyl-COA and
`malonyl-ACP to form a four-carbon product is catalyzed by
`
`CSIRO Exhibit 1014
`
`

`

`962
`
`The Plant Cell
`
`Prokaryotic Pathway - Plastidial compartment
`
`[
`Eukaryotic Pathway - Extraplastidial compartment
`
`["6&2;2,OA 161 -COA
`
`18:1(160)
`18:p;>,'~
`
`Pi
`
`h=dgroup,
`
`activated m,m
`headgroup , lPGl,m
`
`@
`IG3-PI
`
`@
`
`CTP
`
`-1
`
`PPi
`
`@
`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 of glycerol.
`The eukaryotic pathway occurs outside the plastid (primarily at the ER), uses acyl-CoAs as substrates, and positions 18-carbon fatty acids at
`position 2 of glycerol-3-phosphate. The amount of lipid synthesized by the prokaryotic pathways varies in angiosperms from 5 to 40% depending
`on the plant species and the tissue. Reactions 1 and 2 are 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, is believed responsible for producing chain lengths from
`six 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 is
`a 3-ketoacyl-ACP. Three additional reactions occur after each
`condensation to form a saturated fatty acid (Figure 3). The
`3-ketoacyl-ACP is reduced at the carbonyl group by the en-
`zyme 3-ketoacyl-ACP reductase, which uses NADPH as the
`electron donor. The next reaction is dehydration by hydroxyacyl-
`ACP dehydratase. Each round of fatty acid synthesis is then
`completed by the enzyme enoyl-ACP reductase, which uses
`NADH or NADPH to reduce the trans-2 double bond to form
`a saturated fatty acid. The combined action of these four reac-
`tions leads to the lengthening of the precursor fatty acid by
`two carbons while it is still attached to ACP as a thioester.
`The fatty acid biosynthesis pathway produces saturated fatty
`acids, but in most plant tissues, over 75% of the fatty acids
`are unsaturated. The first double bond is introduced by the
`soluble enzyme stearoyl-ACP desaturase. This fatty acid
`desaturase is unique to the plant kingdom in that all other
`known desaturases are integral membrane proteins. The clon-
`ing of this soluble enzyme has recently led to its crystallization
`and the determination of its three-dimensional structure by x-ray
`crystallography (J. Shanklin, personal communication). This
`and other structural studies have led to the first detailed in-
`sights into the mechanism of fatty acid desaturation and the
`nature of the active site. The enzyme is a homodimer in which
`each monomer has an independent active site consisting of
`a diiron-oxo cluster. The two iron atoms are coordinated within
`a central four helix bundle in which the motif (D/E)-E-X-R-H
`is 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 terminated
`when the acyl group is removed from ACP This can happen
`in 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 fatty
`acid from ACP to glycerol-3-phosphate or to monoacylglycerol-
`3-phosphate. The first of these acyltransferases is a soluble
`enzyme that prefers oleoyl-ACP as a substrate. The second
`acyltransferase resides on the inner chloroplast envelope
`membrane and preferentially selects palmitoyl-ACP Whether
`the fatty acid is released from ACP by a thioesterase or an
`acyltransferase determines whether it leaves the plastid. If a
`thioesterase acts on acyl-ACP, then the free fatty acid is able
`to leave the plastid. It is not known how free fatty acids are
`transported out of the plastid, but it may occur by simple diffu-
`sion across the envelope membrane. On the outer membrane
`of the chloroplast envelope, an acyl-COA synthetase is thought
`to assemble an acyl-COA thioester that is then available for
`acyltransferase reactions to form glycerolipids in the endoplas-
`mic reticulum (ER). How the acyl-COA moves from the outer
`chloroplast envelope to the ER is also unknown, but it may
`involve 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 the
`plastid is to form the hydrophobic portion of glycerolipid
`
`CSIRO Exhibit 1014
`
`

`

`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 form
`phosphatidic acid (PA; Figure 4). Diacylglycerol (DAG) is
`produced from PA by a specific phosphatase; alternatively, a
`nucleotide-activated form of DAG (CDP-DAG) is produced from
`the reaction of PA with cytidine 5'4riphosphate (CTP). The
`energy to drive attachment of the polar headgroup during de
`novo glycerolipid synthesis is provided by nucleotide activa-
`tion. When DAG is the lipid substrate, it is the headgroup that
`is activated. Thus, cytidine 5'diphosphate (CDP)choline, CDP-
`ethanolamine, and CDP-methylethanolamine can be substrates
`for phospholipid synthesis, and UDP-galactose and UDP-sulfo-
`quinovose are substrates for monogalactosyldiacylglycerol
`(MGDG) and sulfoquinovosyldiacylglycerol (SL) synthesis,
`respectively. Conversely, when CDP-DAG is the lipid substrate,
`reactions with myo-inositol, serine, and glycerol-3-phosphate
`result in formation of the phospholipids phosphatidylinositol
`(Pl), phosphatidylserine (PS), and phosphatidyl glycerol
`phosphate (the precursor of PG), respectively. Digalactosyl-
`diacylglycerol (DGDG) is synthesized from MGDG (Joyard et
`al., 1993).
`Although the synthetic pathways are presented here as a
`linear series of simple enzymatic steps, the actual biochemis-
`try involved is complicated by the possibilities of headgroup
`modification (for example, the synthesis of phosphatidylcho-
`line [PC] from phosphatidylmethylethanolamine (PE] by two
`rounds of methylation) and headgroup exchange. The details
`of 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).
`
`to the synthesis of plastid lipids (Figure 5). Evidence from
`severa1 Arabidopsis mutants indicates that lipid exchange
`between the ER and the chloroplast is reversible to some ex-
`tent (Miquel and Browse, 1992; Browse et al., 1993) because
`extra chloroplastic membranes in mutants deficient in ER
`desaturases (see later discussion) contain polyunsaturated fatty
`acids derived from the chloroplasts.
`In many species of higher plants, PG is the only product
`of the prokaryotic pathway, and the remaining chloroplast lipids
`are synthesized entirely by the eukaryotic pathway. In other
`species, including Arabidopsis and spinach, both pathways
`contribute about equally to the synthesis of MGDG, DGDG,
`and SL (Browse and Somerville, 1991), and the leaf lipids of
`such plants characteristically contain substantial amounts of
`hexadecatrienoic acid (16:3), which is found only in MGDG
`and DGDG molecules produced by the prokaryotic pathway.
`These plants have been termed 16:3 plants to distinguish them
`from the other angiosperms (18:3 plants), whose galactolipids
`contain predominantly linolenate. The contribution of the eu-
`karyotic pathway to MGDG, DGDG, and SL synthesis is reduced
`in lower plants, and in many green algae the chloroplast is
`almost entirely autonomous with respect to membrane lipid syn-
`thesis. One problem presented by the two-pathway model is
`the need to move hydrophobic lipid molecules from the ER to
`other sites, particularly the chloroplast. Until recently, a class of
`soluble proteins characterized (from in vitro experiments) as
`lipid transfer proteins had been considered to be the intracellular
`transporters. However, biochemical and immunohistochemical
`evidence 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).
`
`Two Pathways for Membrane Lipid Synthesis
`
`Membrane Desaturases
`
`As outlined in Figure 4, higher plants possess two distinct path-
`ways for the synthesis of glycerolipids: the prokaryotic pathway
`o