Fatty acid

m

t

t

a

Metabolic Engineering 4, 12–21 (2002)doi:10.1006/mben.2001.0204, available online at http://www.idealibrary.co

Metabolic Engineering of Fat

Jay J. Thelen and

Department of Plant Biology, Michigan Sta

Received June 28, 2001;

Fatty acids are the most abundant form of reduced carbon chainsavailable from nature and have diverse uses ranging from food toindustrial feedstocks. Plants represent a significant renewable sourceof fatty acids because many species accumulate them in the form oftriacylglycerol as major storage components in seeds. With theadvent of plant transformation technology, metabolic engineering ofoilseed fatty acids has become possible and transgenic plant oilsrepresent some of the first successes in design of modified plant pro-ducts. Directed gene down-regulation strategies have enabled thespecific tailoring of common fatty acids in several oilseed crops. Inaddition, transfer of novel fatty acid biosynthetic genes from non-commercial plants has allowed the production of novel oil composi-tions in oilseed crops. These and future endeavors aim to produceseeds higher in oil content as well as new oils that are more stable, arehealthier for humans, and can serve as a renewable source of indus-trial commodities. Large-scale new industrial uses of engineeredplant oils are on the horizon but will require a better understanding offactors that limit the accumulation of unusual fatty acid structures inseeds. © 2002 Elsevier Science

INTRODUCTION

Plant oils represent a vast renewable resource of highlyreduced carbon. Current world vegetable oil production isestimated at 87 million metric tons with an approximatemarket value of 40 billion U.S. dollars (Gunstone, 2001).Currently, the majority of vegetable oil goes directly tohuman consumption and as much as 25% of human caloricintake in developed countries is derived from plant fattyacids (Broun et al., 1999). In addition to their importance inhuman nutrition, plant fatty acids are also major ingre-dients of nonfood products such as soaps, detergents,lubricants, biofuels, cosmetics, and paints (Ohlrogge, 1994).

121096-7176/02 $35.00© 2002 Elsevier ScienceAll rights reserved.

The demand for vegetable oils has increased steadily inrecent years (Gunstone, 2001) but production capacity tomeet this demand is more than adequate and prices ofvegetable oils have remained below or near $0.6 perkilogram. This low cost of production has stimulatedinterest in use of vegetable oils as renewable alternatives topetroleum-derived chemical feedstocks.

on

y Acid Biosynthesis in Plants

John B. Ohlrogge

e University, East Lansing, Michigan 48824

ccepted August 22, 2001

Fatty acids stored in plant seeds are usually unbranchedcompounds with an even number of carbons ranging from12 to 22 and with 0 to 3 cis double bonds.1 However,

1 Note on lipid nomenclature. A simple shorthand notation based onmolecule length and the number and position of double bonds has beendeveloped to designate fatty acids. For example, the common monounsa-turated fatty acid oleic acid (octadecenoic acid) is designated 18:1. The firstvalue, 18, represents the number of carbons. The second value, 1, indicatesthe number of double bonds. In addition, the position of the double bonds,counting from the carboxyl group is designated by delta (D) and oleic acidcan be more fully designated as 18:1 D9. The double bonds in naturallyoccurring fatty acids are almost exclusively cis isomers, and usually no

numerous variations on this theme exist in nature particu-larly with regard to additional functional groups such ashydroxy, epoxy, cyclopropene, or acetylenic. Plants repre-sent a large reservoir of fatty acid diversity, synthesizing atleast 200 different types of fatty acids (van de Loo et al.,1993). Human use, however, has been predominantlyrestricted to a select few fatty acids that accumulate indomesticated plants. The four most important oilseed cropsare, in descending order, soybean, oil palm, rapeseed, andsunflower, which together account for 65% of currentworldwide vegetable oil production (Gunstone, 2001). Theabundant fatty acids produced in these major commercialoilseeds comprise just 4 of the > 200 possibilities, namelylinoleate, palmitate, laurate, and oleate.

Why Plant Oils Are Attractive Targets for MetabolicEngineering

Metabolic engineering of plant oils has attracted indus-trial and academic researchers for several reasons. First,although the fatty acid content and composition of plantmembranes are highly conserved, seed oils vary greatlyamong plant species. This suggests that the storage form of

⁄

designation for the type of double bond is used unless it is a trans isomer, asin 16:1 D3t. Some authors also designate the positions of the double bondsrelative to the terminal methyl carbon. Thus, an omega-3 fatty acid con-tains a double bond 3 carbons from the methyl end of the fatty acid (e.g.,18:3 D9, 12, 15 is an omega-3 fatty acid). The position at which a fatty acidis esterified to the glycerol backbone of glycerolipids is designated sn-3 (theterminal hydroxyl that is phosphorylated in glycerol 3-phosphate), sn-2(the central hydroxyl), and sn-1 (the terminal hydroxyl that is notphosphorylated).

fatty acids is tolerant to changes in chemical structure and isa good target for genetic manipulations that are unlikely todisturb the physiology of the plant. Second, up to one-thirdof plant oil is already used for nonfood applications and thechemical industry is familiar with fatty acid chemistry andapplications. Third, as noted above, over 200 differentfatty acid structures with attractive functional propertiesoccur in plants. In many cases the pathways that producethese structures have been identified (review: Voelker andKinney, 2001). Finally, rising costs of imported petroleumcoupled with efforts to move toward renewable resourcessuggest good long-term prospects for increased use of plantoils to provide biobased alternatives to petroleum.

Because plant oils have broad uses in both food andnonfood applications the goals of plant oilseed biotech-nologists are diverse. The major goals can be summarizedas:

• Increase content of ‘‘healthy’’ fatty acids and reduce‘‘unhealthy’’ fatty acids.

• Improve oil stability to expand applications andreduce the need for hydrogenation.

• Expand the repertoire of fatty acids available at lowcost and high volume through exploitation of geneticdiversity and enzyme engineering.

• Increase oil content to reduce production costs.

Some success has been achieved in reaching all of thesegoals. In at least two cases, this has led to new commercialcrops and thus oilseed engineering has led the way toward anew generation of agricultural products whose traits havebeen enhanced through metabolic engineering. In othercases attempts to modify plant oils have had disappointingoutcomes that reveal our ignorance of lipid biochemistryand seed metabolism. This review discusses some recentadvances toward the goal of engineering qualitative andquantitative fatty acid traits in plants and some of thechallenges that have emerged.

Overview of Fatty Acid and Triacylglycerol Biosynthesisin Plants

In plants, the reactions for de novo fatty acid synthesis(FAS)2 are located in plastids (Ohlrogge et al., 1979), whichare plant-specific organelles bound by an envelope doublemembrane. Priming and elongation of nascent acyl chains

Fatty Acid Biosynthesis

2 Abbreviations used: ACP, acyl carrier protein; ACCase, acetyl-CoAcarboxylase; BC, biotin carboxylase; CHD, coronary heart disease; ER,endoplasmic reticulum; FAS, fatty acid synthesis; KAS, 3-ketoacyl-ACPsynthase; LPAAT, lysophosphatidic acid acyltransferase; TAG, triacyl-glycerol.

requires acetyl- and malonyl-CoA, respectively, as directprecursors (Fig. 1A). The fatty acid synthase machinery is

13

similar to prokaryotes in that the enzymatic components areseparable polypeptides rather than large multifunctionalpolypeptides as found in animals and fungi. The series ofreactions necessary for de novo synthesis of fatty acids, up to18 C in length, has been elucidated and is discussed indetail elsewhere (Schultz and Ohlrogge, 2001; Voelker andKinney, 2001). The first desaturation step for fatty acids iscatalyzed by a plastidial stearoyl-acyl carrier protein (ACP)desaturase. Termination of plastidial fatty acid chainelongation is catalyzed by acyl-ACP thioesterases, whichhydrolyze acyl chains from ACP. After termination, freefatty acids are activated to CoA esters, exported from theplastid, and assembled into glycerolipids at the endoplasmicreticulum (ER). In addition, further modifications (desatu-ration, hydroxylation, elongation, etc.) occur in the ERwhile acyl chains are esterified to glycerolipids or CoA(Fig. 1B). In developing seeds, the flux of acyl chains in theER eventually leads to esterification on all three positions ofglycerol to form triacylglycerol (TAG). The low polarity ofTAG is believed to result in the accumulation of this lipidbetween bilayer leaflets leading to the budding of storageorganelles termed oil bodies.

TAILORING OF COMMON FATTY ACIDSIN OILSEED CROPS

Modification of naturally occurring common fatty acidsfound in oilseed crops has led to major technical achieve-ments and a commercial product in transgenic high-oleicsoybean oil. Simply by overexpressing or suppressing singlegenes it has been possible to make large compositionalchanges (Table 1). Because seed-specific promoters are usedthe changes have been restricted to the storage oils of seeds,which appear to tolerate a wide range of oil physicalproperties.

Current medical understanding indicates a strong impactof dietary fatty acids on cardiovascular disease andhuman health (Hu et al., 2001). Consequently, there ismuch interest in tailor-producing healthier vegetable oilsand such products may help to balance consumer opposi-tion to ‘‘GMO’’ foods. One health concern regardingvegetable oil-derived food is the presence of trans-unsa-turated fatty acids. Most vegetable oil used for foodapplications is partially or fully hydrogenated during pro-

Metabolic Engineering 4, 12–21 (2002)doi:10.1006/mben.2001.0204

cessing to make the oil semisolid for spreads and alsoto increase oxidative stability during storing or frying(Kinney, 1996). Industrial hydrogenation increases satu-rated fatty acid content and also results in production oftrans-isomers of unsaturated fatty acids that are normallynot found in vegetable oils and have been associated withcoronary heart disease (Broun et al., 1999). For many

lrars

a

FIG. 1. Fatty acid synthesis, modification, and assembly into triacylgtransgenic plants and are described in Table 1. (A) Simplified scheme ofterminated by acyl-ACP thioesterases (FatA and FatB classes), which reletion to CoA at the plastid envelope. (B) Simplified scheme of reactions focerols. After activation to CoA, fatty acids formed in the plastid can belysophosphatidic acid (LPA), phosphatidic acid (PA), diacylglycerol, andinvolves movement through phosphatidylcholine (PC) pools where modificor plants transformed with jojoba genes are wax esters formed in seeds.

food applications, vegetable oils with a reduced amount oftrans-unsaturated fatty acids are desirable to improvehuman health. This has been achieved using strategiessuch as cosuppression, antisense, and RNA interference todown-regulate endogenous stearoyl-ACP desaturase genesin soybean, cotton, and Brassica oilseeds (Table 1). Inthese plants, levels of stearic were increased up to 40% toprovide a semisolid margarine feedstock without the needfor hydrogenation.

An oxidatively stable liquid oil low in saturated fattyacids was also produced in soybean by suppression of theoleoyl desaturase (Kinney, 1996). Oleic acid content wasincreased up to 86%, 18:2 content was reduced from 55%to less than 1%, and saturated fatty acids were reduced to10%. This oil has been produced commercially and isextremely stable for high-temperature frying applications.


In addition, its stability matches that of mineral oil-derived lubricants and therefore nonfood uses as bio-degradable lubricants are under way. An added benefit toconsumers from future use of engineered high-oleic oils infoods may be a reduction in coronary heart disease (CHD)associated with high omega-6 fatty acid consumption. Inrecent years evidence has accumulated that the balance of

14

ycerols in plants. Numbers refer to reactions that have been modified ineactions of plastid fatty acid synthesis. In oilseeds, fatty acid synthesis isse free fatty acids, allowing their export from the plastid and reesterifica-modification of fatty acids in oilseeds and their assembly into triacylgly-

equentially esterified directly to glycerol 3-phosphate (G-3-P) to producetriacylglycerol. However, in most oilseeds the major flux of acyl chainstions such as further desaturation and hydroxylation occur. Only in jojoba

omega-3 and omega-6 unsaturated fatty acids in dietsinfluences the risk of CHD (Hu et al., 2001). The domi-nance of plant oils with high omega-6 18:2 in many dietshas led to omega-6/omega-3 consumption ratios near10:1, whereas populations that consume ratios near 1:1(e.g., Greenland, Japan) have strikingly lower incidence ofCHD.

ENGINEERING OF UNUSUAL FATTY ACIDSIN OILSEED CROPS

Among the approximately 200 fatty acid structuresproduced by plants are several that might find wide use ifavailable in high quantity and at low cost. Included in thislist are hydroxy, epoxy, conjugated, and acetylenic fattyacids, all of which result from the action of enzymes

Thelen and Ohlrogge

closely related to the ubiquitous oleoyl desaturase (Brounet al., 1998). These fatty acids have interest because theyprovide a second reactive functional group to a hydro-carbon chain and offer opportunities for polymerizationsor other chemical modifications. Therefore, considerableinterest has developed in engineering high-yielding oilseedsto produce these or other specialty fatty acids and in a few

L

n

e

st

st

st

st

st

-9

s

-9

-9

st

D

u

x

x

)

a

(

i

TAB

Selected Examples of Fatty Acid

E

Engineered Transgenic Max. produced

fatty acid plant (mol %) Reaction (numb

Caprylic, capric Brassica napus 38 Acyl-ACP thioe

Lauric Brassica napus 58 Acyl-ACP thioe

Lauric Arabidopsis 24 Acyl-ACP thioe

Palmitic Arabidopsis 39 Acyl-ACP thioe

Palmitic Brassica napus 34 Acyl-ACP thioe

Stearic Soybean 30 Stearoyl-ACP D

oleoyl-D-12 de

Stearic Brassica napus 40 Stearoyl-ACP D

Stearic Cotton 38 Stearoyl-ACP D

Stearic Brassica napus 22 Acyl-ACP thioe

Petroselinic Tobacco 4 Palmitoyl-ACP

Oleic Soybean 86 Oleoyl-D-12 desa

Oleic Brassica napus 89 Oleoyl-D-12 desa

Oleic Cotton 77 Oleoyl-D-12 desa

Oleic Brassica juncea 73 Oleoyl-D-12 desa

Oleic Arabidopsis 54 Oleoyl-D-12 desa

c-Linolenic (18:3 w-6) Brassica napus 47 Oleoyl-D-6 and D

c-Linolenic acid Tobacco 1 Oleoyl-D-6 desat

Eleostearic, parinaric Soybean 17 Conjugase (11)

D-5 Eicosenoic Soybean 18 b-Ketoacyl-CoA

acyl-CoA desa

Hydroxy fatty acids Arabidopsis 30 Oleate-12-hydro

Ricinoleic Arabidopsis 17 Oleate-12-hydro

Acetylenic Arabidopsis 25 Acetylenase (11)

12, 13-Epoxy-cis-9-oleic Arabidopsis 15 Epoxygenase (11

Wax esters Arabidopsis 70 b-ketoacyl synth

acyl-CoA redu

wax synthase

Note. The numbers after the names of engineered reactions refer to Foccur at the ER or other nonplastidial membrane.

cases such engineering efforts have been successful (sum-marized in Table 1). In the following we discuss selectedexamples of the engineering of novel fatty acids in plants.


Engineering of Fatty Acid Chain Length

Plants that accumulate short- to medium-chain (C8 toC14) fatty acids in seed triacylglycerol have seed-specificacyl-ACP thioesterase activities toward the correspondingacyl-ACPs (Pollard et al., 1991; Davies, 1993). For example,California bay and Cuphea seeds accumulate up to 90%

15

E 1

Engineering in Transgenic Plants

gineered reaction(s)

Gene

r) Gene source regulation Reference

erase (1) Cuphea Up Dehesh et al., 1996

erase (2) California bay Up Voelker et al., 1996

erase (2) California bay Up Voelker et al., 1992

erase (3) Arabidopsis Up Dormann et al., 2000

erase (3) Cuphea Up Jones et al., 1995

(5) and Soybean Down Kinney, 1996

aturase (7)

desat (5) Brassica Down Knutzon et al., 1992

desat (5) Cotton Down Liu et al., 2000

erase (4) Mangosteen Up Hawkins and Kridl, 1998

-4 desat (6) Coriander Up Cahoon et al., 1992

turase (7) Soybean Down Kinney, 1996

turase (7) Brassica Down Stoutjesdijk et al., 2000

turase (7) Cotton Down Liu et al., 2000

turase (7) Brassica Down Stoutjesdijk et al., 2000

turase (7) Arabidopsis Down Okuley et al., 1994

-12 desat (7) Mortierella apina Up Ursin et al., 2000

rase (7) Cyanobacteria Up Reddy and Thomas, 1996

Momordica Up Cahoon et al., 1999

synthase (8), Meadowfoam Up Cahoon et al., 2000

turase (9)

ylase (10) Castor, Lesquerella Up Smith et al., 2000

ylase (10) Castor Up Brown and Somerville, 1997

Crepis Up Lee et al., 1998

Crepis Up Singh et al., 2000

se (12), Jojoba Up Lardizabal et al., 2000

ctase (13),

14)

g. 1A and 1B. Reactions 1 to 6 occur in the plastid and reactions 7 to 14

short chain saturated fatty acids in triacylglycerol. Ingroundbreaking studies, expression of a California baythioesterase in the seeds of non-laurate (12:0)-accumu-lating plants, Arabidopsis and Brassica napus (rapeseed),


resulted in the ‘‘short-circuiting’’ of acyl chain elongationto produce laurate up to 24 and 58% of total seed fattyacids, respectively (Table 1; Voelker et al., 1992, 1996).Position analysis of TAG revealed that laurate was presentat both sn-1 and sn-3 positions but not the sn-2 position.Lack of laurate at the sn-2 position was attributed to thehigh specificity of lysophosphatidic acid acyltransferase

(LPAAT). Further increases in laurate yield seemed pos-sible if all three positions of TAG were acylated withlaurate. The introduction of a laurate-specific coconutLPAAT into rapeseed containing the California baythioesterase resulted in further increases in laurate levels,up to 67% total fatty acid, by catalyzing laurate acylationat the sn-2 position of TAG (Knutzon et al., 1999). Thislevel of laurate is higher than observed in palm kernel, acommercial source of laurate. Applications of high-lauraterapeseed oil include detergents and soaps, a large marketthat is currently met by imported palm kernel and coconutoils.

The previous example demonstrates that the transfer ofa single gene into rapeseed confers laurate accumulation atlevels very similar to California bay seed. However, suchsuccess with a single gene may be the exception ratherthan the rule. For example, when a medium-chain thio-esterase from Cuphea hookeriana was introduced intorapeseed, caprylate (8:0) accumulated to only 12% intransgenic rapeseed, while Cuphea contains 50% caprylate(Dehesh et al., 1996). In another investigation, expressionof elm or nutmeg FatB thioesterases in rapeseed did notresult in seed containing 65% caprate (10:0) or 80%laurate as observed in these two respective plants butrather 4% caprate and 20% laurate, respectively (Voelkeret al., 1997). In these examples short-chain fatty acids weresignificantly lower in transgenic hosts compared to donorspecies. One explanation for these differences is the lowavailability of short-chain acyl-ACP pools for thioesterasetermination in non-short-chain-accumulating plants. Thiswas addressed by crossing plants expressing condensingenzymes (3-ketoacyl-ACP synthase, KAS) from Cupheathat have unique specificity for 6:0-(caproic) and 8:0-acyl-ACPs with lines carrying Cuphea FatB thioesterases(Leonard et al., 1998; Dehesh et al., 1998). All lines carry-ing both a Cuphea KAS and a Cuphea thioesterase hadhigher levels of short-chain fatty acids than the single-transgene parents. Enhancement of short-chain fatty acidaccumulation was attributed to the short-chain specificityof the Cuphea KAS, effectively increasing 10:0- and 12:0-acyl-ACP pool sizes for short-chain thioesterase cleavage.Thus obtaining significant amounts of short-chain fattyacids in TAG may require multiple genes which increasethe substrate pools for the thioesterase as well as short-chain-specific acyltransferases which can assemble the


novel fatty acids into TAG.

Plants Sometimes Fight Back against MetabolicEngineering Schemes

An unexpected lesson learned from the study of laurate-producing transgenic plants described above was that high-

16

level production of novel fatty acids can induce a futilecycle of fatty acid synthesis and degradation (Fig. 2).By analyzing hundreds of independent transgenic lines,Voelker et al., (1996) found that laurate production incanola seeds increased linearly up to about 35 mol% withincreased lauroyl-ACP thioesterase expression. However,to achieve 58 mol% laurate required 10-fold higherlevels of the introduced enzyme, raising the question ofwhat limits higher laurate accumulation. Eccleston andOhlrogge (1998) examined these high-laurate canola seedsand found that enzymes for medium-chain fatty acidb-oxidation were increased severalfold, as were malatedehydrogenase and isocitrate lyase, which participate inthe glyoxylate cycle for fatty acid carbon reutilization.These and other results led to the conclusion that highproduction of unusual fatty acids in transgenic hosts caninduce pathways for their breakdown. Surprisingly, seedoil yield was not reduced, which led to the additionaldiscovery that the FAS pathway was also induced,presumably to compensate for the loss by oxidation ofmedium-chain fatty acids.

Production of Waxes

Long-chain wax esters were once harvested from spermwhales and were a major ingredient of industrial lubri-cants and transmission fluids. Banning of whale harvestsled to searches for alternative biological sources of suchstructures. Jojoba, a desert shrub found in the Americansouthwest, is the only plant species known to accumulatewaxes (up to 60% dry weight) rather than TAG as a seed

FIG. 2. Scheme for a futile cycle of production and oxidation of

Thelen and Ohlrogge

lauric acid in transgenic canola, based on results of Eccleston andOhlrogge (1998). Transgenic seeds that produce 58 mol% lauric acidwere found to have increased activity of lauric acid b-oxidation,isocitrate lyase, and malate dehydrogenase. In addition, up to 50% of[14C]acetate added to seeds was recovered in malate, sucrose, and otherwater-soluble metabolites. These results suggest that up to half the lauricacid produced is degraded and returned to intermediate pools in a futilecycle of fatty acid synthesis and turnover.

storage reserve. These waxes are mostly derived fromC20–C24 monounsaturated fatty acids and alcohols andare synthesized by the elongation of oleate followed byreduction to alcohols by a fatty acid reductase (Metzet al., 2000). The wax storage lipid is formed by a fatty acyl-CoA:fatty alcohol acyltransferase, also referred to as waxsynthase. The reductase and acyltransferase were purifiedfrom jojoba and the corresponding cDNAs cloned (Metz etal., 2000; Lardizabal et al., 2000). Coordinated expressionof three genes—a Lunaria annua long-chain acyl-CoAelongase and the jojoba reductase and acyltransferase—inArabidopsis resulted in wax production at up to 70% of theoil present in mature seeds (Lardizabal et al., 2000).The high levels of accumulation indicated that all the genesnecessary for this trait were identified. If this trait can besuccessfully transferred to commercial crops this wouldrepresent a large potential source of waxes for a variety ofapplications, including cosmetics and industrial lubricants.

Production of Novel Monoenoic Fatty Acids

Introduction of the first double bond in fatty acids occursin plastids by a soluble desaturase specific for acyl-ACPsubstrates. The location of this double bond can varydepending upon specificity of the plastidial acyl-ACP desa-turase. Typically the double bond is inserted betweencarbons 9 and 10 of a stearoyl-ACP substrate. However,seed-specific plastidial acyl-ACP desaturases that intro-duce double bonds at the D4, D6, or D9 position ofpalmitoyl-ACP have been identified from coriander,black-eyed Susan vine (Thunbergia alata), and cat’s claw,respectively, which accumulate these unusual monoenesup to 80% in seed oil (Cahoon et al., 1992, 1994a, 1998).Double-bond position on palmitate and stearate alters thephysical properties such that unusual monoenes havepotentially different commercial uses including monomerfeedstocks for specific nylon polymer applications or ashigher melting unsaturated fatty acids for margarines.Since monomers for most nylons are derived fromthe petrochemical industry there is interest in plants asrenewable sources for these precursors. To achieve wide useof such fatty acids it will be essential to move the unusualmonoene trait into high-yielding oilseed crops fromwhich the oil can be produced at low cost. However, intro-


duction of a coriander D4 16:0-ACP desaturase or aThunbergia D6 16:0-ACP desaturase into tobacco callusand Arabidopsis seed, respectively, resulted in less than10% accumulation of these unusual fatty acids (Cahoonet al., 1992; Schultz and Ohlrogge, 2001). The reason for thelow levels of unusual monoene production in non-nativeplants remains unknown and represents a major challenge

17

in our understanding of plant lipid synthesis. Some evidencesuggests specific isoforms of the cofactors, ferredoxin andACP, may be important for production of unusual monoenes(Suh et al., 1999; Schultz et al., 2000). In addition,coriander and Thunbergia unusual monoenes are incor-porated into phosphatidylcholine pools prior to accu-mulation into TAG (Cahoon et al., 1994b; Schultz andOhlrogge, 2000). Coriander also expresses KAS (Mekhedovet al., 2001), thioesterase (Dörmann et al., 1994), andacyltransferase (Dutta et al., 1992) activities specific forthese unusual fatty acids, which are likely important fortheir accumulation in TAG.

In a recent investigation, transgenic expression of anengineered castor D9 18:0-ACP desaturase (with improvedspecificity toward 16:0-ACP) in Arabidopsis seed resultedin 13% of total seed fatty acids as 16:1D9 and elonga-tion products 18:1D11 and 20:1D13 (Cahoon and Shanklin,2000). Expression of this same desaturase in fab1 Arabi-dopsis mutants containing a lesion in KAS II, which cata-lyzes the elongation of 16:0-ACP to 18:0-ACP, resulted inup to 30% accumulation of the same three fatty acids.Thus availability of 16:0 ACP substrate is likely one limi-tation for unusual monoene production. In addition, thisstudy suggests that novel acyl-ACP desaturases producedby protein engineering strategies may be more effectivethan enzymes derived from wild species.

Product Yield: The New Challenge in OilseedMetabolic Engineering

Identification of key genes as described earlier and theirtransfer into transgenic crops have occupied many aca-demic and industrial laboratories for the past 10–15 years.However, in many cases this is not the central problemin oilseed modification. For a new oil to be economic,the desired fatty acid almost always must be the majorconstituent to avoid expensive purification costs. Despiteimpressive successes with medium-chain fatty acids andwax esters, in most cases in which a newly identified genehas been transferred into another oilseed, the proportionof the desired product in the transgenic host has beenconsiderably lower than in the wild species from whichthe gene was obtained. The activity of the introducedenzyme has generally not been limiting, so it is necessary to


determine what other factors limit product accumulation.Accumulation of unusual fatty acids to levels found

naturally will likely require introduction of activities inaddition to those directly responsible for synthesizing theunusual fatty acid. One possible explanation for this is thepresence of a redundant set of biosynthetic enzymes fornovel fatty acids in seeds. Such a scenario would explain

differences in substrate specificity between seed-specificlipid biosynthetic enzymes and those involved in generalcell lipid synthesis. Presumably this is because mostunusual fatty acids possess physical properties distinctlydifferent from fatty acids commonly found in membranes,and thus plants must possess ‘‘editing’’ or exclusionmechanisms to prevent the accumulation of these fattyacids in lipid bilayers (reviewed in Volker and Kinney,2001). Addressing these issues will require more knowl-edge of the cellular biochemistry in oil-accumulatingtissues than is currently available.

PROGRESS TOWARD INCREASING SEEDOIL CONTENT

For both edible and industrial uses, an increase in seedoil content is desirable and has been a major goal ofoilseed engineering. However, to be economically useful,such a change must not come at the expense of overallseed yield or at the loss of other high-value components.For example, soybean is the largest source of vegetable oil,comprising 30% of the world market, and now consti-tutes over 80% of all dietary vegetable oils in the UnitedStates. Although termed an oilseed, soybean contains only18–22% oil on a seed dry-weight basis and is grown prin-cipally as a high-protein meal for animal feeds. Thus,increasing oil in soybean will in most cases not be useful ifit comes at the expense of high-value soy protein thatdrives the crop’s economics. By comparison, other oilseedcrops (except cotton) are grown primarily for their oil andproduce seeds with 40–60% oil. The wide range of seed oilpercentage observed in nature suggests that this pathwaymight be amenable to metabolic engineering, particularlyin ‘‘low-oil’’ oilseeds, provided the key mechanisms whichcontrol oil content are identified.

Production of Malonyl-CoA by Acetyl-CoA CarboxylaseIs a Key Regulatory Step

The committed step for de novo FAS is the productionof malonyl-CoA catalyzed by acetyl-CoA carboxylase(ACCase) (Fig. 1). Malonyl-CoA production appears to bea potential control point for this pathway, based uponanalysis of acyl-CoA and acyl-ACP pool sizes (Post-Beittenmiller et al., 1991, 1992; Roughan, 1997). Since


malonyl-CoA levels in plastids are very low (less than10%) compared to acetyl-CoA, it seemed likely that up-regulating ACCase activity would increase flux to fattyacids. This has been clearly shown to be the case inEscherichia coli (Davis et al., 2000). The plastidial ACCasefrom most plants is a complex comprising four differentsubunits. One early effort to increase ACCase was to

18

overexpress the biotin carboxylase (BC) subunit using aCaMV 35S promoter in tobacco. Although BC proteinincreased threefold in leaves, there was no accompanyingincrease in the amount of other ACCase subunits(Shintani et al., 1997) and no effect on fatty acid contentor composition. Thus, for ACCase—unlike some other multi-enzyme complexes—overexpressing just one subunit doesnot increase the amount of the remaining subunits.

Evidence that increased malonyl-CoA pools couldincrease fatty acid production was obtained by targeting ahomomeric ACCase to rapeseed plastids (Roesler et al.,1997). Under the control of a seed-specific promoter thischimeric protein resulted in higher ACCase activities andincreased oil yield by 3–5% on a seed dry-weight basis.These data provided the first evidence that seed oil couldbe quantitatively enhanced by increasing the pool size ofmalonyl-CoA precursor. However, the small increasepointed toward additional control points for FAS.

Overexpression of Several Individual Fatty Acid SynthaseEnzymes Does Not Increase Flux through Fatty AcidBiosynthesis

Increasing malonyl-CoA precursor pools for FASresulted in only slight increases in seed oil yield. Such amodest improvement would suggest that another step(s)might be limiting. Could fatty acid synthase activities alsobe limiting FAS? Several labs have addressed this questionby overexpressing enzymes downstream of malonyl-CoAproduction. The conclusion from these investigations isthat up-regulation of any one enzyme does not increaseflux through FAS. Indeed, overexpression of some activi-ties actually decreased FAS and fatty acid content asobserved with the overexpression of a condensing enzyme.

Condensation of acetyl-CoA with malonyl-ACP iscatalyzed by KAS III. Recently, a spinach KAS IIIwas expressed in tobacco and resulted in approximately50-fold increases in activity above control levels. Ratherthan an increase in fatty acid content a 5–10% decreasewas observed (Dehesh et al., 2001). In the same report, aCuphea KAS III expressed in rapeseed seed embryosresulted in a 9% decrease in fatty acid content. Aninteresting and unexpected consequence of KAS IIIoverexpression was an increase in ACP protein levels in

Thelen and Ohlrogge

tobacco leaves, although other fatty acid synthase activi-ties were unaffected. Decreases in fatty acid content as aresult of KAS III overexpression were attributed todecreased rates of de novo FAS most likely by reducingmalonyl-CoA pools for subsequent KAS condensationreactions. In a related study, targeting of an E. colimalonyl-CoA:ACP trans-acylase to rapeseed leucoplasts

increased this plastid activity up to 45-fold but did notincrease fatty acid content (Verwoert et al., 1994).

Based upon the aforementioned and other studies itseems unlikely that the up-regulation of any single fattyacid synthase enzyme will have a major positive effect onFAS flux. Although not all fatty acid synthase enzymeshave been overexpressed to determine the effect on FAS,substantial increases in flux will likely require up-regula-tion of multiple activities. This conclusion has stimulatedmore comprehensive efforts to identify transcriptional,protein kinase, or other regulatory factors that mightup-regulate the entire pathway (Girke et al., 2000).

Preliminary studies suggest that reactions late in theTAG biosynthetic pathway may provide increased sinkstrength that could stimulate increased fatty acid produc-tion. Overexpression of a yeast long-chain sn-2 acyltrans-ferase resulted in > 50% (dry mass/seed) increases in seedoil content of Arabidopsis and rapeseed (Zou et al.,1997). Field trials of the transgenic rapeseed gave increases of8.1–13.5% (Katavic et al., 2000). Recently, Jako et al.(2001) reported that overexpression of an Arabidopsisdiacylglycerol acyltransferase in Arabidopsis seeds can alsoincrease seed oil content as well as seed weight. Together,these studies suggest that increased flux into oil may bemore easily achieved by strategies targeted at the latersteps in the pathway. It is important to note that despiteintense efforts in this area, commercial varieties with con-sistently increased oil yield per hectare have not beenachieved through transgenic means.

CONCLUSIONS

Engineering of FAS has progressed rapidly in the past 5years and has led to the commercialization or field trial ofseveral modified oilseed crops. Although the engineeringof fatty acid chain length and degree and location of fattyacid desaturation has at least been demonstrated in prin-ciple, engineering plants with increased flux through FAShas been difficult. This is likely due to the complexityassociated with the engineering of primary carbon meta-bolism and an unclear picture of how this pathway isregulated in vivo. One of the challenges that lie ahead is tounderstand the mechanism for feedback inhibition of fattyacid production in vivo (Shintani and Ohlrogge, 1995).

Although plants with increased seed oil and those con-


taining nutritional supplements may have an immediatemarket niche, plants engineered to accumulate industrial‘‘specialty oils’’ may encounter problems and will need tobe cost-evaluated on an individual basis (Hitz, 1999).Some of these potential problems include expensive pro-cessing costs, loss of value associated with toxicity of themeal by-product (a particular problem with high-value

19

soybean meal), and occasional undesirable side-effectsresulting from major alterations in fatty acid profiles(Knutzen et al., 1992; Miquel et al., 1993; Miquel andBrowse, 1994). Nevertheless, the long-term forecast ofvegetable oils as an alternative to petroleum for chemicalfeedstocks is not fanciful. With rapid advances occurringin plant lipid biotechnology and the increasing cost ofpetroleum, vegetable oils will eventually provide newcost-effective raw materials.

REFERENCES

Broun, P., Gettner, S., and Somerville, C. (1999). Genetic engineering ofplant lipids. Annu. Rev. Nutr. 19, 197–216.

Broun, P., Shanklin, J., Whittle, E., and Somerville, C. (1998). Catalyticplasticity of fatty acid modification enzymes underlying chemicaldiversity of plant lipids. Science 282, 1315–1317.

Broun, P., and Somerville, C. (1997). Accumulation of ricinoleic,lesquerolic, and denispolic acids in seeds of transgenic Arabidopsisplants that express a fatty acid hydroxylase cDNA from castor bean.Plant Physiol. 113, 933–942.

Cahoon, E. B., Carlson, T. J., Ripp, K. G., Schweiger, B. J., Cook, G. A.,Hall, S. E., and Kinney, A. J. (1999). Biosynthetic origin ofconjugated double bonds: Production of fatty acid components ofhigh-value drying oils in transgenic soybean embryos. Proc. Natl.Acad. Sci. USA 96, 12935–12940.

Cahoon, E. B., Cranmer, A. M., Shanklin, J., and Ohlrogge, J. B.(1994a). Delta-6-hexadecenoic acid is synthesized by the activity of asoluble delta 6 palmitoyl-acyl carrier protein desaturase in Thunbergiaalata endosperm. J. Biol. Chem. 269, 27519–27526.

Cahoon, E. B., Marillia, E.-F., Stecca, K. L., Hall, S. E., Taylor, D. C.,and Kinney, A. J. (2000). Production of fatty acid components ofmeadowfoam oil in somatic soybean embryos. Plant Physiol. 124,243–251.

Cahoon, E. B., and Ohlrogge, J. B. (1994b). Apparent role of phospha-tidylcholine in the metabolism of petroselinic acid in developingUmbelliferae endosperm. Plant Physiol. 104, 845–855.

Cahoon, E. B., Shah, S., Shanklin, J., and Browse, J. (1998). A deter-minant of substrate specificity predicted from the acyl-acyl carrierprotein desaturase of developing cat’s claw seed. Plant Physiol. 117,593–598.

Cahoon, E. B., and Shanklin, J. (2000). Substrate-dependent mutantcomplementation to select fatty acid desaturase variants for meta-bolic engineering of plant seed oils. Proc. Natl. Acad. Sci. USA 97,12350–12355.

Cahoon, E. B., Shanklin, J., and Ohlrogge, J. B. (1992). Expressionof a coriander desaturase results in petroselinic acid production intransgenic tobacco. Proc. Natl. Acad. Sci. USA 89, 11184–11188.

Davies, H. M. (1993). Medium chain acyl-ACP hydrolysis activities ofdeveloping oilseeds. Phytochemistry 33, 1353–1356.


Davis, M. S., Solbiati, J., and Cronan, J. E. (2000). Overproduction ofacetyl-CoA carboxylase activity increases the rate of fatty acidbiosynthesis in Escherichia coli. J. Biol. Chem. 15, 28593–28598.

Dehesh, K., Edwards, P., Fillatti, J., Slabaugh, M., and Byrne, J. (1998).KAS IV: A ketoacyl-ACP synthase from Cuphea sp. is a mediumchain specific condensing enzyme. Plant J. 15, 383–390.

Dehesh, K., Jones, A., Knutzon, D. S., and Voelker, T. A. (1996).Production of high levels of 8:0 and 10:0 fatty acids in transgenic

canola by overexpression of Ch FatB2, a thioesterase cDNA fromCuphea hookeriana. Plant J. 9, 167–172.

Dehesh, K., Tai, H., Edwards, P., Byrne, J., and Jaworski, J. G. (2001).Overexpression of 3-ketoacyl acyl carrier protein synthase IIIs inplants reduces the rate of lipid synthesis. Plant Physiol. 125,1103–1114.

Dörmann, P., Frentzen, M., and Ohlrogge, J. B. (1994). Specificities ofthe acyl-acyl carrier protein (ACP) thioesterase and glycerol-3-phosphate acyltransferase for octadecenoyl-ACP isomers. PlantPhysiol. 104, 839–844.

Dörmann, P., Voelker, T. A., and Ohlrogge, J. B. (2000). Accumulationof palmitate in Arabidopsis mediated by the acyl-acyl carrier proteinthioesterase FATB1. Plant Physiol. 123, 637–643.

Dutta, P. C., Appelqvist, L.-A., and Stymne, S. (1992). Utilization ofpetroselinate (C 18:1 delta 6) by glycerol acylation enzymes inmicrosomal preparations of developing embryos of carrot (Daucuscarota L.), safflower (Carthamus tinctorius L.), and oil rape (Brassicanapus L.). Plant Sci. 81, 57–64.

Eccleston, V., and Ohlrogge, J. B. (1998). Expression of lauroyl-acylcarrier protein thioesterase in Brassica napus seeds induces pathwaysfor both fatty acid oxidation and biosynthesis and implies a set-pointfor triacylglycerol accumulation. Plant Cell 10, 613–621.

Girke, T., Todd, J., Ruuska, S., White, J., Benning, C., and Ohlrogge, J. B.(2000). Microarray analysis of developing Arabidopsis seeds. PlantPhysiol. 124, 1570–1581.

Gunstone, F. D. (2001). Soybeans pace boost in oilseed production.Inform 11, 1287–1289.

Hawkins, D. J., and Kridl, J. C. (1998). Characterization of acyl-ACPthioesterases of mangosteen (Garcinia mangostana) seed and high levelsof stearate production in transgenic canola. Plant J. 13, 743–752.

Hitz, B. (1999). Economic aspects of transgenic crops which producenovel products. Curr. Opin. Plant Biol. 2, 135–138.

Hu, F. B., Manson, J. E., and Willett, W. C. (2001). Types of dietary fatand risk of coronary heart disease: A critical review. J. Am. Coll.Nutr. 20, 5–19.

Jako, C., Kumar, A., Wei, Y., Zou, J., Barton, D. L., Giblin, M.,Covello, P. S., and Taylor, D. C. (2001). Seed-specific over-expressionof an Arabidopsis cDNA encoding a diacylglycerol acyltransferaseenhances seed oil content and seed weight. Plant Physiol. 126,861–874.

Jones, A., Davies, H. M., and Voelker, T. A. (1995). Palmitoyl-acylcarrier protein (ACP) thioesterase and the evolutionary origin of plantacyl-ACP thioesterases. Plant Cell 7, 359–371.

Katavic, V., Friesen, W., Barton, D. L., Gossen, K. K., Giblin, E. M.,Luciw, T., An, J., Zou, J., MacKenzie, S. L., Keller, W. A., Males, D.,and Taylor, D. C. (2000). Utility of the Arabidopsis FAE1 and yeastSLC1–1 genes for improvements in erucic acid and oil content inrapeseed. Biochem. Soc. Trans. 28, 935–937.

Kinney, A. J. (1996). Development of genetically engineered soybean oilsfor food applications J. Food Lipids 3, 273–292.

Knutzon, D. S., Hayes, T. R., Wyrick, A., Xiong, H., Davies, H. M., andVoelker, T. A. (1999). Lysophosphatidic acid acyltransferase fromcoconut endosperm mediates the insertion of laurate at the sn-2 posi-tion of triacylglycerols in lauric rapeseed oil and can increase total


laurate levels. Plant Physiol. 120, 739–746.Knutzon, D. S., Thompson, G. A., Radke, S. E., Johnson, W. B., Knauf,

V. C., and Kridl, J. C. (1992). Modification of Brassica seed oil byantisense expression of a stearoyl-acyl carrier protein desaturase gene.Proc. Natl. Acad. Sci. USA 89, 2624–2628.

Lardizabal, K. D., Metz, J. G., Sakamoto, T., Hutton, W. C., Pollard,M. R., and Lassner, M. W. (2000). Purification of a jojoba embryowax synthase, cloning of its cDNA, and production of high levels

20

of wax in seeds of transgenic Arabidopsis. Plant Physiol. 122,645–655.

Lee, M., Lenman, M., Banas, A., Bafor, M., Singh, S., Schweizer, M.,Nilsson, R., Liljenberg, C., Dahlqvist, A., Gummeson, P. O., Sjodahl,S., Green, A., and Stymne, S. (1998). Identification of non-hemediiron proteins that catalyze triple bond and epoxy group formation.Science 280, 915–918.

Leonard, J. M., Knapp, S. J., and Slabaugh, M. B. (1998). A Cupheabeta-ketoacyl-ACP synthase shifts the synthesis of fatty acidstowards shorter chains in Arabidopsis seeds expressing Cuphea FatBthioesterases. Plant J. 13, 621–628.

Liu, Q., Singh, S., and Green, A. (2000). Genetic modification of cottonseed oil using inverted-repeat gene-silencing techniques. Biochem. Soc.Trans. 28, 927–929.

Mekhedov, S., Cahoon, E., and Ohlrogge, J. (2001). An unusual seedspecific 3-ketoacyl-ACP synthase associated with the biosynthesis ofpetroselinic acid in coriander. Plant Mol. Biol., in press.

Metz, J. G., Pollard, M. R., Anderson, L., Hayes, T. R., and Lassner,M. W. (2000). Purification of a jojoba embryo fatty acyl-coenzyme Areductase and expression of its cDNA in high erucic acid rapeseed.Plant Physiol. 122, 635–644.

Miquel, M. F., and Browse, J. A. (1994). High-oleate oilseeds fail todevelop at low temperature. Plant Physiol. 106, 421–427.

Miquel, M., James, D., Donner, H., and Browse, J. (1993). Arabidopsisrequires polyunsaturated lipids for low-temperature survival. Proc.Natl. Acad. Sci. USA 90, 6208–6212.

Ohlrogge, J. B. (1994). Design of new plant products: Engineering offatty acid metabolism. Plant Physiol. 104, 821–826.

Ohlrogge, J. B., Kuhn, D. N., and Stumpf, P. K. (1979). Subcellularlocalization of acyl carrier protein in leaf protoplasts of Spinaciaoleracea. Proc. Natl. Acad. Sci. USA 76, 1194–1195.

Okuley, J., Lightner, J., Feldmann, K., Yadav, N., Lark, E., andBrowse, J. (1994). Arabidopsis FAD2 gene encodes the enzymethat is essential for polyunsaturated lipid synthesis. Plant Cell 6,147–158.

Pollard, M. R., Anderson, L., Fan, C., Hawkins, D. J., and Davies,H. M. A. (1991). Specific acyl-ACP thioesterase implicated in medium-chain fatty acid production in immature cotyledons of Umbellulariacalifornica. Arch. Biochem. Biophys. 284, 2306–2312.

Post-Beittenmiller, M. A., Jaworski, J. G., and Ohlrogge, J. B. (1991). Invivo pools of free and acylated acyl carrier proteins in spinach:Evidence for sites of regulation of fatty acid biosynthesis. J. Biol.Chem. 266, 1858–1865.

Post-Beittenmiller, M. A., Roughan, G., and Ohlrogge, J. B. (1992).Regulation of plant fatty acid biosynthesis: Analysis of acyl-CoAand acyl-acyl carrier protein substrate pools in spinach and peachloroplast. Plant Physiol. 100, 923–930.

Reddy, A. S., and Thomas, T. L. (1996). Expression of a cyanobacterialdelta-6 desaturase gene results in gamma-linolenic acid production intransgenic plants. Nat. Biotechnol. 14, 639–642.

Roesler, K., Shintani, D., Savage, L., Boddupalli, S., and Ohlrogge, J.(1997). Targeting of the Arabidopsis homomeric acetyl-coenzyme Acarboxylase to plastids of rapeseeds. Plant Physiol. 113, 75–81.

Roughan, P. G. (1997). Stromal concentrations of coenzyme A and its

Thelen and Ohlrogge

esters are insufficient to account for rates of chloroplast fatty acidsynthesis: Evidence for substrate channelling within the chloroplastfatty acid synthase. Biochem. J. 327, 267–273.

Schultz, D. J., and Ohlrogge, J. B. (2000). Biosynthesis of triacylglycerolin Thunbergia alata: Additional evidence for involvement of phospha-tidylcholine in unusual monoenoic oil production. Plant Physiol.Biochem. 38, 169–175.

Schultz, D. J., and Ohlrogge, J. B. (2001). Metabolic engineering of fattyacid biosynthesis. In ‘‘Lipid Biotechnology,’’ in press.

Schultz, D. J., Suh, M. C., and Ohlrogge, J. B. (2000). Stearoyl-acylcarrier protein and unusual acyl-acyl carrier protein desaturaseactivities are differentially influenced by ferredoxin. Plant Physiol.124, 681–692.

Shintani, D. K., and Ohlrogge, J. B. (1995). Feedback inhibition of fattyacid synthesis in tobacco suspension cells. Plant J. 7, 577–587.

Shintani, D. K., Roesler, K. R., Shorrosh, B. S., Savage, L. J., andOhlrogge, J. B. (1997). Antisense expression and overexpression ofbiotin carboxylase in tobacco leaves. Plant Physiol. 114, 881–886.

Singh, S., Thomaeus, S., Lee, M., Green, A., and Stymne, S. (2000).Inhibition of polyunsaturated fatty acid accumulation in plantsexpressing a fatty acid epoxygenase. Biochem. Soc. Trans. 28,940–942.

Smith, R. G., Gauthier, D. A., Dennis, D. T., and Turpin, D. H.(1992). Malate- and pyruvate-dependent fatty acid synthesis inleucoplasts from developing castor endosperm. Plant Physiol. 98,1233–1238.

Smith, M., Moon, H., and Kunst, L. (2000). Production of hydroxy fattyacids in the seeds of Arabidopsis thaliana. Biochem. Soc. Trans. 28,947–950.

Stoutjesdijk, P. A., Hurlestone, C., Singh, S. P., and Green, A. G. (2000).High-oleic acid Australian Brassica napus and B. juncea varietiesproduced by co-suppression of endogenous delta-12 desaturases.Biochem. Soc. Trans. 28, 938–940.

Suh, M. C., Schultz, D. J., and Ohlrogge, J. B. (1999). Isoforms of acylcarrier protein involved in seed-specific fatty acid synthesis. Plant J.17, 679–688.

Ursin, V., Knutzon, D., Radke, S., Thornton, J., and Knauf, V. (2000).Production of beneficial dietary omega-3 and omega-6 fatty acids in


21

transgenic canola. In ‘‘Abstracts of the 14th International Symposiumon Plant Lipids,’’ p. 13.

van de Loo, F. J., Fox, B. G., and Somerville, C. (1993). Unusual fattyacids. In ‘‘Plant Lipids’’ (T. Moore, Ed.), pp. 91–126, CRC, BocaRaton, FL.

Verwoert, I. I. G. S., van der Linden, K. H., Nijkamp, H. J. J., andStuitje, A. R. (1994). Developmental specific expression and organelletargeting of the Escherichia coli fabD gene, encoding malonylcoenzyme A-acyl carrier protein transacylase in transgenic rape andtobacco seeds. Plant Mol. Biol. 26, 189–202.

Voelker, T. A., Hayes, T. R., Cranmer, A. M., Turner, J. C., and Davies,H. M. (1996). Genetic engineering of a quantitative trait: Metabolicand genetic parameters influencing the accumulation of laurate inrapeseed. Plant J. 9, 229–241.

Voelker, T. A., Jones, A., Cranmer, A. M., Davies, H. M., and Knutzon,D. S. (1997). Broad-range and binary-range acyl-acyl carrier proteinthioesterases suggest an alternative mechanism for medium-chainproduction in seeds. Plant Physiol. 114, 669–677.

Voelker, T. A., and Kinney, A. J. (2001). Variations in the biosynthesisof seed storage lipids. Annu. Rev. Plant Physiol. Mol. Biol. 52,335–361.

Voelker, T. A., Worrell, A. C., Anderson, L., Bleibaum, J., Fan, C.,Hawkins, D. J., Radke, S. E., and Davies, H. M. (1992). Fatty acidbiosynthesis redirected to medium chains in transgenic oilseed plants.Science 257, 72–74.

Zou, J. T., Katavic, V., Giblin, E. M., Barton, D. L., Mackenzie, S. L.,Keller, W. A., Hu, X., and Taylor, D. C. (1997). Modification of seedoil content and acyl composition in the Brassicaceae by expression ofa yeast sn-2-acyltransferase gene. Plant Cell 9, 909–923.


Fatty acid

Food

plant fatty acids

common fatty acids

plant seeds

accfatty acids

new oils

use of vegetable oils

transgenic plant oilspresent

plant fattycids broun