Acyl Carriers Used as Substrates by the Desaturases and Elongases Involved in Very Long-chain Polyunsaturated Fatty Acids Biosynthesis Reconstituted in Yeast
2003; Elsevier BV; Volume: 278; Issue: 37 Linguagem: Inglês
10.1074/jbc.m305990200
ISSN1083-351X
AutoresFrédéric Domergue, Amine Abbadi, Claudia Ott, T. Zank, Ulrich Zähringer, Ernst Heinz,
Tópico(s)Plant biochemistry and biosynthesis
ResumoThe health benefits attributed to very long-chain polyunsaturated fatty acids and the long term goal to produce them in transgenic oilseed crops have led to the cloning of all the genes coding for the desaturases and elongases involved in their biosynthesis. The encoded activities have been confirmed in vivo by heterologous expression, but very little is known about the actual acyl substrates involved in these pathways. Using a Δ6-elongase and front-end desaturases from different organisms, we have reconstituted in Saccharomyces cerevisiae the biosynthesis of arachidonic acid from exogenously supplied linoleic acid in order to identify these acyl carriers. Acyl-CoA measurements strongly suggest that the elongation step involved in polyunsaturated fatty acids biosynthesis is taking place within the acyl-CoA pool. In contrast, detailed analyses of lipids revealed that the two desaturation steps (Δ5 and Δ6) occur predominantly at the sn-2 position of phosphatidylcholine when using Δ5- and Δ6-desaturases from lower plants, fungi, worms, and algae. The specificity of these Δ6-desaturases for the fatty acid acylated at this particular position as well as a limiting re-equilibration with the acyl-CoA pool result in the accumulation of γ-linolenic acid at the sn-2 position of phosphatidylcholine and prevent efficient arachidonic acid biosynthesis in yeast. We confirm by using a similar experimental approach that, in contrast, the human Δ6-desaturase uses linoleoyl-CoA as substrate, which results in high efficiency of the subsequent elongation step. In addition, we report that Δ12-desaturases have no specificity toward the lipid polar headgroup or the sn-position. The health benefits attributed to very long-chain polyunsaturated fatty acids and the long term goal to produce them in transgenic oilseed crops have led to the cloning of all the genes coding for the desaturases and elongases involved in their biosynthesis. The encoded activities have been confirmed in vivo by heterologous expression, but very little is known about the actual acyl substrates involved in these pathways. Using a Δ6-elongase and front-end desaturases from different organisms, we have reconstituted in Saccharomyces cerevisiae the biosynthesis of arachidonic acid from exogenously supplied linoleic acid in order to identify these acyl carriers. Acyl-CoA measurements strongly suggest that the elongation step involved in polyunsaturated fatty acids biosynthesis is taking place within the acyl-CoA pool. In contrast, detailed analyses of lipids revealed that the two desaturation steps (Δ5 and Δ6) occur predominantly at the sn-2 position of phosphatidylcholine when using Δ5- and Δ6-desaturases from lower plants, fungi, worms, and algae. The specificity of these Δ6-desaturases for the fatty acid acylated at this particular position as well as a limiting re-equilibration with the acyl-CoA pool result in the accumulation of γ-linolenic acid at the sn-2 position of phosphatidylcholine and prevent efficient arachidonic acid biosynthesis in yeast. We confirm by using a similar experimental approach that, in contrast, the human Δ6-desaturase uses linoleoyl-CoA as substrate, which results in high efficiency of the subsequent elongation step. In addition, we report that Δ12-desaturases have no specificity toward the lipid polar headgroup or the sn-position. Very long-chain polyunsaturated fatty acids (VLC-PUFAs) 1The abbreviations used are: VLC-PUFAs, very long-chain polyunsaturated fatty acids; LA, linoleic acid (18:2Δ9,12); GLA, γ-linolenic acid (18:3Δ6,9,12); ARA, arachidonic acid (20:4Δ5,8,11,14); FAMEs, fatty acid methyl esters; PC, phosphatidylcholine; PI, phosphatidylinositol; PS, phosphatidylserine; PE, phosphatidylethanolamine; DGD, diglucosyl-diacylglycerol; FID, flame-ionization detector; KCS, β-ketoacyl-CoA synthase; FAE, fatty acid elongation; MES, 4-morpholineethanesulfonic acid.1The abbreviations used are: VLC-PUFAs, very long-chain polyunsaturated fatty acids; LA, linoleic acid (18:2Δ9,12); GLA, γ-linolenic acid (18:3Δ6,9,12); ARA, arachidonic acid (20:4Δ5,8,11,14); FAMEs, fatty acid methyl esters; PC, phosphatidylcholine; PI, phosphatidylinositol; PS, phosphatidylserine; PE, phosphatidylethanolamine; DGD, diglucosyl-diacylglycerol; FID, flame-ionization detector; KCS, β-ketoacyl-CoA synthase; FAE, fatty acid elongation; MES, 4-morpholineethanesulfonic acid. such as arachidonic acid (ARA, 20:4Δ5,8,11,14), eicosapentaenoic acid (20:5Δ5,8,11,14,17), and docosahexaenoic acid (22:6Δ4,7,10,13,16,19) are important constituents of membranes (particularly in the retina and the central nervous system) as well as precursors of several biologically active eicosanoids (1Kinsella J.E. Lokesh B. Broughton S. Whelan J. Nutrition. 1990; 6: 24-44PubMed Google Scholar). The presence of VLC-PUFAs in the human diet affects diverse physiological processes involved in cardiovascular, immune, neuronal, and visual functions (2Spector A.A. Lipids. 1999; 34: S1-S3Crossref PubMed Google Scholar). Many clinical studies have linked PUFA intake with normal health and development, particularly in the case of newborns and infants (3Simopoulos A.P. Prostaglandins Leuk. Essent. Fatty Acids. 1999; 60: 421-429Abstract Full Text PDF PubMed Scopus (169) Google Scholar). VLC-PUFAs are mainly found in fish, in some fungi and lower plants, as well as in a variety of microorganisms of the phytoplankton. With the exception of the anaerobically operating polyketide synthase-like systems found in some marine bacteria and primitive eukaryotes (4Metz J.G. Roessler P. Facciotti D. Levering C. Dittrich F. Lassner M. Valentine R. Lardizabal K. Domergue F. Yamada A. Yazawa K. Knauf V. Browse J. Science. 2001; 293: 290-293Crossref PubMed Scopus (544) Google Scholar), VLC-PUFAs are synthesized by elongation and desaturation of linoleic acid (LA, 18:2Δ9,12)or α-linolenic acid (ALA, 18:3Δ9,12,15) in the endoplasmic reticulum. Most algae, fungi, and lower plants producing VLC-PUFAs possess the entire biosynthetic pathway to synthesize these fatty acids from acetate, whereas mammalia, which lack Δ12- and Δ15-desaturases, use as precursors LA and ALA that have to be supplied in their diet and thus are essential fatty acids. The numerous health benefits attributed to VLC-PUFAs as well as the absence of sustainable and low cost sources has led to the long-term goal of producing such fatty acids in transgenic oilseed crops (5Abbadi A. Domergue F. Meyer A. Riedel K. Sperling P. Zank T.K. Heinz E. Eur. J. Lipid Sci. Technol. 2001; 103: 106-113Crossref Scopus (29) Google Scholar). Using organisms producing VLC-PUFAs such as the fungus Mortierella alpina, the moss Physcomitrella patens, the worm Caenorhaditis elegans, and the diatom Phaeo-dactylum tricornutum as gene sources, a large collection of sequences coding for elongases and desaturases was created in the last 10 years (reviewed in Ref. 6Drexler H. Spiekermann P. Meyer A. Domergue F. Zank T.K. Sperling P. Abbadi A. Heinz E. J. Plant Physiol. 2002; 160: 779-802Crossref Scopus (81) Google Scholar). Each coding sequence was separately expressed in yeast or plant and the substrate specificity of the encoded enzyme verified so that cDNAs encoding all the enzymatic activities required for DHA synthesis are available. The fatty acid desaturases involved in VLC-PUFA biosynthesis can be divided into two groups, the ω6-/ω3-desaturases and the so-called front-end desaturases (7Aitzetmüller K. Tsevegsüren N. J. Plant. Physiol. 1994; 143: 538-543Crossref Scopus (43) Google Scholar), which contain a cytochrome b 5-domain fused to their N terminus (8Sperling P. Heinz E. Eur. J. Lipid Sci. Technol. 2001; 103: 158-180Crossref Scopus (59) Google Scholar). Whereas the latter group of desaturases inserts the new double bond between the fatty acid carboxyl group and a pre-existing double bond, the ω6-/ω3-desaturases insert it between a pre-existing double bond and the fatty acid methyl end. Using alkenylether glycerolipids and tomato cell cultures, it was unambiguously proven that plant ω6- and ω3-desaturases are acting on lipid-linked substrates (9Sperling P. Linscheid M. Stocker S. Muhlbach H.P. Heinz E. J. Biol. Chem. 1993; 268: 26935-26940Abstract Full Text PDF PubMed Google Scholar). In addition, biochemical studies with plants and fungi strongly suggest that ω6-desaturases are acting on both positions (sn-1 and sn-2) of phosphatidylcholine (PC), whereas Δ6-desaturases are confined to the sn-2 position of PC (10Galle-Le Bastard A.M. Demandre C. Oursel A. Joseph M. Mazliak P. Kader J.C. Physiol. Plant. 2000; 108: 118-124Crossref Scopus (8) Google Scholar, 11Griffiths G. Stobart A.K. Stymne S. Biochem. J. 1988; 252: 641-647Crossref PubMed Scopus (64) Google Scholar, 12Jackson F.M. Fraser T.C. Smith M.A. Lazarus C. Stobart A.K. Griffiths G. Eur. J. Biochem. 1998; 252: 513-519Crossref PubMed Scopus (40) Google Scholar). On the other hand, fatty acid desaturases from vertebrates are referred to as acyl-CoA desaturases (13Sprecher H. Curr. Opin. Clin. Nutr. Metab. Care. 1999; 2: 135-138Crossref PubMed Scopus (46) Google Scholar, 14Okayasu T. Nagao M. Ishibashi T. Imai Y. Arch Biochem. Biophys. 1981; 206: 21-28Crossref PubMed Scopus (108) Google Scholar). In contrast to the fatty acid desaturases involved in VLC-PUFA biosynthesis, the elongation activities remain to be biochemically characterized. Elongation of PUFAs has been linked to ELO sequences. ELO-type proteins were first characterized in yeast, where ELO1 is involved in the elongation of medium-chain saturated and monounsaturated fatty acids, whereas ELO2 and ELO3 catalyze the subsequent elongation yielding the C24–26 saturated fatty acids present in sphingolipids (15Oh C.S. Toke D.A. Mandala S. Martin C.E. J. Biol. Chem. 1997; 272: 17376-17384Abstract Full Text Full Text PDF PubMed Scopus (393) Google Scholar, 16Toke D.A. Martin C.E. J. Biol. Chem. 1996; 271: 18413-18422Abstract Full Text Full Text PDF PubMed Scopus (157) Google Scholar). Related sequences were then identified in M. alpina (17Parker-Barnes J.M. Das T. Bobik E. Leonard A.E. Thurmond J.M. Chaung L.T. Huang Y.S. Mukerji P. Proc. Natl. Acad. Sci. U. S. A. 2000; 97: 8284-8289Crossref PubMed Scopus (135) Google Scholar), C. elegans (18Beaudoin F. Michaelson L.V. Hey S.J. Lewis M.J. Shewry P.R. Sayanova O. Napier J.A. Proc. Natl. Acad. Sci. U. S. A. 2000; 97: 6421-6426Crossref PubMed Scopus (133) Google Scholar), man (19Leonard A.E. Bobik E.G. Dorado J. Kroeger P.E. Chuang L.T. Thurmond J.M. Parker-Barnes J.M. Das T. Huang Y.S. Mukerji P. Biochem. J. 2000; 350: 765-770Crossref PubMed Scopus (183) Google Scholar), and P. patens (20Zank T.K. Zähringer U. Beckmann C. Pohnert G. Boland W. Holtorf H. Reski R. Lerchl J. Heinz E. Plant J. 2002; 31: 255-268Crossref PubMed Scopus (90) Google Scholar), and their involvement in the elongation of PUFAs was demonstrated by expression in yeast. Nevertheless, so far there is no unequivocal evidence that any polypeptide encoded by an ELO sequence catalyzes the actual condensation reaction involved in fatty acid elongation. Compared with the poorly described elongation of PUFAs, the elongation of saturated and monounsaturated fatty acids has been extensively characterized biochemically in both plant and rat liver microsomes. Each C2 elongation is a four-step process (condensation-reduction-dehydration-reduction), which involves four different enzymes, most probably organized in a multifunctional complex. The rate-limiting step of this process is the condensation of the acyl primer with malonyl-CoA catalyzed by the β-ketoacyl-CoA synthase (KCS or FAE for fatty acid elongation), which also confers substrate specificity to the whole elongase complex. Expression of the KCS/FAE condensing enzyme or of an ELO-type protein alone is sufficient to restore elongation activity in yeast, suggesting that the three other enzymes are ubiquitously expressed. Recently, using an Arabidopsis thaliana KCS/FAE condensing enzyme purified nearly to homogeneity, Ghanevati and Jaworski (21Ghanevati M. Jaworski J.G. Eur. J. Biochem. 2002; 269: 3531-3539Crossref PubMed Scopus (57) Google Scholar) measured high in vitro activities with various acyl-CoAs, indicating that acyl-CoAs are most probably the substrates of this type of elongase. Since the elongation process condenses an acyl-CoA primer with malonyl-CoA and finally results in the production of a C2-elongated acyl-CoA, the entire elongation process most probably takes place within the acyl-CoA pool. When the biosynthesis of VLC-PUFAs was reconstituted in yeast by co-expressing the Δ6-desaturase from Borago officinalis, the Δ6-elongase from C. elegans and the Δ5-desaturase from M. alpina in the presence of LA or ALA, small but significant amounts of ARA and EPA, respectively, were detected (18Beaudoin F. Michaelson L.V. Hey S.J. Lewis M.J. Shewry P.R. Sayanova O. Napier J.A. Proc. Natl. Acad. Sci. U. S. A. 2000; 97: 6421-6426Crossref PubMed Scopus (133) Google Scholar). We obtained similar results by co-expressing the Δ5- and Δ6-desaturases from P. tricornutum together with the Δ6-elongase from P. patens (22Bröker M. Bauml O. Gottig A. Ochs J. Bodenbenner M. Amann E. Appl. Microbiol. Biotechnol. 1991; 34: 756-764Crossref PubMed Scopus (33) Google Scholar). Despite this success, these reconstitution experiments were rather inefficient compared with the situation in the genuine organisms and regarding the relatively high activities measured with the separately expressed enzymes. A closer look at the activities of the different enzymes expressed to reconstitute VLC-PUFAs biosynthesis in yeast showed that the elongation of endogenously produced Δ6-fatty acids was less than half that observed with exogenously supplied Δ6-fatty acids. These results suggest a great difference in the availability of exogenously added or in situ produced Δ6-fatty acids for elongation: in contrast to exogenously supplied fatty acids, those produced endogenously by Δ6-desaturation may remain in a pool that is not available for elongation, which consequently limits VLC-PUFAs biosynthesis. In the present work we sought to identify which acyl carriers could be used as substrate by the different enzymes involved in the biosynthesis of ARA reconstituted in S. cerevisiae. In view of the data presented above, special attention was paid to phosphatidylcholine and the acyl-CoA pool. Using cDNAs from various organisms, the four activities leading to the synthesis of arachidonic acid from oleic acid (Δ12-desaturase, Δ6-desaturase, Δ6-elongase, and Δ5-desaturase) were expressed in yeast separately or in combination. After short or long incubation times, the fatty acid profiles of various lipid pools were determined in order to evaluate which acyl carriers are preferentially used by the different desaturases and the elongase. Materials—Restriction enzymes, polymerases, and DNA-modifying enzymes were obtained from New England Bioloabs (Frankfurt, Germany) unless indicated otherwise. All other chemicals were from Sigma. Construction of Vectors—The different fatty acid desaturases and yeast expression constructs used in this study are listed in Table I. Usually, complete open reading frames (ORF) were modified by PCR to create appropriate restriction sites adjacent to the start and stop codons. The amplified DNAs were cloned into the pGEM-T vector (Promega, Madison, WI) before being released and cloned into a yeast expression vector (pVT102-U, pYES2 or pESC-LEU) using the restriction sites inserted by PCR.Table IFatty acid desaturases used in this studySource organismRegio-specificityGenBank™ Accession No.Yeast expression vectorRestriction sites usedSource Ref.Phaeodactylum tricornutumΔ5AY082392pESC-LEU-PSEISpeI/SacI(22Bröker M. Bauml O. Gottig A. Ochs J. Bodenbenner M. Amann E. Appl. Microbiol. Biotechnol. 1991; 34: 756-764Crossref PubMed Scopus (33) Google Scholar)Mortierella alpinaΔ5AF172755pYES2EcoRI/XhoI(46Knutzon D.S. Thurmond J.M. Huang Y.S. Chaudhary S. Bobik Jr., E.G. Chan G.M. Kirchner S.J. Mukerji P. J. Biol. Chem. 1998; 273: 29360-29366Abstract Full Text Full Text PDF PubMed Scopus (131) Google Scholar)Physcomitrella patensΔ5pYES2KpnI/XhoI(47Sperling P. Lucht J.M. Egener T. Reski R. Cirpus P. Heinz E. Murata M.Y.M. Nishida I. Okuyama H. Sekiya J. Wada H. Advanced Research on Plant Lipids. Kluwer Academic Publishers, Dordrecht2003: 113-116Crossref Google Scholar)Phytophthora megaspemaΔ5CAD5323pYES2BamHI/SphIaI. Feussner (unpublished); Patent WO 03012092-A6.Caenorhabditis elegans bExpressed in the yeast strain INVSc1 (Invitrogen).Δ5AY078796pESC-TRPHindIII/BamHI(48Michaelson L.V. Napier J.A. Lewis M. Griffiths G. Lazarus C.M. Stobart A.K. FEBS Lett. 1998; 439: 215-218Crossref PubMed Scopus (77) Google Scholar)Phaeodactylum tricornutumΔ6AY082393pVT102-UBamHI/XhoI(22Bröker M. Bauml O. Gottig A. Ochs J. Bodenbenner M. Amann E. Appl. Microbiol. Biotechnol. 1991; 34: 756-764Crossref PubMed Scopus (33) Google Scholar)Physcomitrella patensΔ6CAA11033pVT102-UBamHI/XhoI(49Girke T. Schmidt H. Zähringer U. Reski R. Heinz E. Plant J. 1998; 15: 39-48Crossref PubMed Scopus (169) Google Scholar)Ceratodon purpureusΔ6CAB94993pYES2KpnI/EcoRI(50Sperling P. Lee M. Girke T. Zähringer U. Stymne S. Heinz E. Eur. J. Biochem. 2000; 267: 3801-3811Crossref PubMed Scopus (72) Google Scholar)Homo sapiensΔ6NP-004256pYES2HindIII/XhoI(51Marquardt A. Stohr H. White K. Weber B.H. Genomics. 2000; 66: 175-183Crossref PubMed Scopus (227) Google Scholar)Borago officinalisΔ6AAC49700pYES2BamHI/XhoI(18Beaudoin F. Michaelson L.V. Hey S.J. Lewis M.J. Shewry P.R. Sayanova O. Napier J.A. Proc. Natl. Acad. Sci. U. S. A. 2000; 97: 6421-6426Crossref PubMed Scopus (133) Google Scholar)Mortierella alpinaΔ6AF1101510pYES2HindIII/EcoRI(52Huang Y.S. Chaudhary S. Thurmond J.M. Bobik Jr., E.G. Yuan L. Chan G.M. Kirchner S.J. Mukerji P. Knutzon D.S. Lipids. 1999; 34: 649-659Crossref PubMed Scopus (127) Google Scholar)Helianthus annuusΔ12AF251842pYES2BamHI/BamHI(45Martinez-Rivas J.M. Sperling P. Lühs W. Heinz E. Mol. Breeding. 2001; 8: 159-168Crossref Scopus (107) Google Scholar)Phaeodactylum tricornutumΔ12AY165023pYES2BamHI/XhoI(31Domergue F. Spiekermann P. Lerchl J. Beckmann C. Kilian O. Kroth P.G. Boland W. Zähringer U. Heinz E. Plant Physiol. 2003; 131: 1648-1660Crossref PubMed Scopus (118) Google Scholar)a I. Feussner (unpublished); Patent WO 03012092-A6.b Expressed in the yeast strain INVSc1 (Invitrogen). Open table in a new tab Expression in S. cerevisiae—The S. cerevisiae strain C13ABYS86 (leu2, ura3, his, pra1, prb1, prc1, cps) (22Bröker M. Bauml O. Gottig A. Ochs J. Bodenbenner M. Amann E. Appl. Microbiol. Biotechnol. 1991; 34: 756-764Crossref PubMed Scopus (33) Google Scholar) was used in all the expressions described in this study. Transformation, selection and growth of the transgenic yeast cells have already been described (23Domergue F. Lerchl J. Zähringer U. Heinz E. Eur. J. Biochem. 2002; 269: 4105-4113Crossref PubMed Scopus (135) Google Scholar). When the cultures had reached an OD600 of about 0.2, expressions were induced by supplementing galactose (2%, w/v) and the appropriate fatty acids to a final concentration of 500 μm. All cultures were then grown for another 24 or 48 h at 20 or 30 °C, as indicated, and harvested by centrifugation. For short time pulses, cultures were grown for 24 h at 30 °C, reaching an OD600 of about 1.5, before the exogenous fatty acid was added. After 1 min, 2.2-ml aliquots were harvested and the cells sedimented by short centrifugation (20 s). After removal of the supernatant, the cell pellets were frozen in liquid nitrogen and stored at –80 °C until needed. Lipid Analysis—Lipid analysis of transgenic yeast cells were made from 150-ml cultures grown for 24 h at 20 °C unless otherwise indicated. Cells were harvested by centrifugation, washed with 30 ml of 0.1 m NaHCO3 and the lipids were extracted on a shaker for 4 h with 15 ml of chloroform/methanol (1:1) and then for 20 h with 15 ml of chloroform/methanol (2:1). The resulting organic phase was extracted with 9 ml of 0.45% NaCl, dried with Na2SO4, and evaporated under vacuum. The residue was dissolved in 2 ml of chloroform and corresponded to the total lipid extract. The major lipid classes PC (phosphatidylcholine), PI+PS (phosphatidylinositol and -serine), PE (phosphatidylethanolamine) and NL (neutral lipids) were purified by thin layer chromatography using chloroform/methanol/acetic acid (65:35:8; v/v/v) as solvent mixture. The different spots were scraped off the plate and the lipids were extracted from silica by adding successively 400 μl of water, 2 ml of methanol, and 2 ml of chloroform and vigorously shaking. After adding 2 ml of 0.2 m H3PO4/1 m KCl, the organic phase was extracted and the resulting aqueous phase re-extracted with 2 ml of chloroform. Both organic phases were combined, dried with Na2SO4 and evaporated under argon. The residues dissolved in 2 ml of chloroform corresponded to the different lipid fractions. For quantification, an aliquot of the total lipid extract was resolved by thin layer chromatography using chloroform/methanol/acetic acid (65:35:8; v/v/v) as solvent mixture. After drying, the plate was dipped in 10% CuSO4 in 8% (v/v) phosphoric acid, dried at 100 °C and heated to 178 °C until the spots appeared (24Weerheim A.M. Kolb A.M. Sturk A. Nieuwland R. Anal. Biochem. 2002; 302: 191-198Crossref PubMed Scopus (120) Google Scholar). The different spots were then quantified by scanning densitometry using a CAMAG TLC Scanner 3 (CAMAG, Muttenz, Switzerland) as already described (25Grether-Beck S. Bonizzi G. Schmitt-Brenden H. Felsner I. Timmer A. Sies H. Johnson J.P. Piette J. Krutmann J. EMBO J. 2000; 19: 5793-5800Crossref PubMed Scopus (96) Google Scholar). Positional Analysis—Positional analyses were conducted using the Rhizopus arrhizus lipase (26Fischer W. Heinz E. Zeus M. Hoppe Seylers Z Physiol. Chem. 1973; 354: 1115-1123Crossref PubMed Scopus (145) Google Scholar) from Sigma. Samples containing PC or PE in chloroform were dried under argon and resuspended in 1 ml of 0.03% Triton X-100, 2 mm CaCl2, 50 mm HEPES, pH 7.2 by sonication. 10,000 units of lipase were added and the digestions were conducted for 2 h at 37 °C. The aqueous phase was extracted with 2.5 ml of chloroform/methanol (2:1) and then 2 ml of chloroform. The resulting organic phase was dried under argon and separated by thin layer chromatography using chloroform/methanol/aqueous ammonia (65:25:0.7; v/v/v) as solvent mix. The spots corresponding to free fatty acids and lysophospholipids were scraped and directly transmethylated for gas-liquid chromatography analysis. Total and Esterified Fatty Acid Analysis—For the analysis of total fatty acids, fatty acid methyl esters (FAMEs) were prepared from sedimented cell pellets or lipid extracts by direct transmethylation with 0.5 m sulfuric acid in methanol containing 2% (v/v) dimethoxypropane. After 1 h at 80 °C, 0.2 ml of 5 m NaCl were added and FAMEs were extracted with 1 ml of petroleum-ether. For the analysis of esterified fatty acids, 1.35 ml of toluene/methanol (1:2, v/v), and 0.5 ml of 0.5 m NaOCH3 in methanol were successively added to sedimented cell pellets. After homogenization, samples were shaken for 1 h at room temperature before adding 2 ml of petroleum-ether and 0.4 ml of 5 m NaCl and extracting FAMEs. FAMEs were then analyzed by gas-liquid chromatography as previously described (23Domergue F. Lerchl J. Zähringer U. Heinz E. Eur. J. Biochem. 2002; 269: 4105-4113Crossref PubMed Scopus (135) Google Scholar). Acyl-CoA Analysis—For acyl-CoA analysis, the highly sensitive method relying on the fluorescence of etheno derivatives developed by Larson and Graham for plant tissues (27Larson T.R. Graham I.A. Plant J. 2001; 25: 115-125Crossref PubMed Google Scholar) was used. Frozen yeast cell pellets equivalent to 2.2 ml of yeast culture (OD600 of 1.5) were used as starting material and the extraction of acyl-CoAs was performed exactly as described (27Larson T.R. Graham I.A. Plant J. 2001; 25: 115-125Crossref PubMed Google Scholar). After drying the extracted acyl-CoAs under argon at 50 °C, derivatization was carried out in 300 μl of chloracetaldehyde derivatization reagent (27Larson T.R. Graham I.A. Plant J. 2001; 25: 115-125Crossref PubMed Google Scholar) at 85 °C for 20 min. HPLC analysis was made using a Thermoquest HPLC system (Thermoquest, Egels-bach, Germany) equipped with a LUNA 150 × 2.0 mm column with phenylhexyl-coated 5 μm silica particles (Phenomenex, Torrance, CA) under the same conditions as described by Larson et al. (28Larson T.R. Edgell T. Byrne J. Dehesh K. Graham I.A. Plant J. 2002; 32: 519-527Crossref PubMed Scopus (65) Google Scholar). Acyl-CoA were identified using saturated and mono-unsaturated acyl-CoAs from Sigma or enzymatically synthesized polyunsaturated acyl-CoAs (18:2Δ9,12-, 18:3Δ6,9,12-, 20:2Δ11,14-, 20:3Δ8,11,14-, and 20:4Δ5,8,11,14-CoA) as standards. Synthesis was achieved in 200 μl reaction mixture containing 100 mm Tris-HCl, pH 8.1, 10 mm MgCl2, 5 mm CoASH, 2 mm dithiothreitol, 25 μm free fatty acid, and 2.5 units of acyl-coenzyme A synthetase from Pseudomanas sp. (Sigma). After2hof incubation at 37 °C, the reaction was stopped with 50 μl of glacial acetic acid/ethanol 1:1 (v/v), and the samples were washed with petroleum ether to extract unreacted fatty acids. Acyl-CoAs were purified on a short reverse phase column (Strata C18-E, Phenomenex, Torrance, CA) using acetonitrile as eluent, dried under argon, and dissolved in 50 mm MES, pH 5.0. Exogenously Supplied Fatty Acids Enter the Yeast Lipid Metabolism via the Acyl-CoA Pool—A large body of evidence suggests that exogenously supplied fatty acids enter the yeast cell by concomitant conversion to acyl-CoAs before their use for various metabolic purposes. Nevertheless, to our knowledge the time course of this conversion has not yet been studied experimentally. For a first approximate evaluation of this time scale, a yeast growing culture was pulsed for 1 min with LA before evaluating the fatty acid profile in three different fractions: the total fatty acids, the esterified fatty acids and the acyl-CoA pool (Fig. 1). These patterns were compared with control samples prepared from the same culture just before the addition of LA. Before adding LA (Fig. 1A), the fatty acid composition of total and esterified fatty acids were practically identical, indicating the absence of a significant pool of free fatty acids. The monounsaturated fatty acids (16:1 and 18:1) represented about 76% of the total fatty acids, whereas the saturated 16:0 and 18:0 accounted for about 18 and 6%, respectively. These major fatty acids were also found in the acyl-CoA pool, but in significantly different proportions. Palmitoleoyl-CoA was the major acyl-CoA species (about 36%), 16:0-CoA, 18:0-CoA, and 18:1-CoA accounted each for about 20%, and myristoyl-CoA represented less than 1%. One minute after adding LA (Fig. 1B), the profiles of the total and esterified fatty acids clearly differed, since LA represented about 50% of the total, but only 4% of the esterified acyl groups. This difference indicates that after 1 min in the presence of LA, only a very small proportion of the exogenously supplied fatty acid has already been channeled into lipids, and that most of the LA, which has been bound to and/or incorporated by the yeast cells, remains in the form of the free fatty acid. In marked contrast, the fatty acid profile of the acyl-CoA pool is dominated by LA, which represents about 60% of the acyl groups, supporting the assumption that exogenously supplied fatty acids are converted to acyl-CoAs when entering the yeast cells. These data also indicated that the acyl-CoA pool could be extensively and specifically flooded with a particular fatty acid within a very short time, enabling the possibility to detect acyl-group modifications taking place in that pool. It should be added that after 24 h, LA was still dominating the acyl-CoA pool, but that the profile of the esterified and total fatty acids were similar (data not shown), indicating that all the LA exogenously supplied had been incorporated and acylated into the yeast lipids. Elongation of GLA Takes Place in the Acyl-CoA Pool—We then used the same approach with a yeast expressing an ELO-type elongase in order to see whether the elongation of GLA takes place within the acyl-CoA pool. For this experiment, a yeast culture transformed with a construct carrying the gene of the Δ6-elongase from P. patens was grown in the presence of galactose for 24 h before adding GLA so that the elongase was present within the cells before supplying its substrate. The fatty acid composition of the three different fractions defined above were then determined before and 1 min after the addition of GLA. Before the pulse, the fatty acid profiles were similar to those reported in Fig. 1A (data not shown). After 1 min in the presence of GLA (Fig. 1C), GLA represented about 40% of the total, but only 2.5% of the esterified fatty acids, confirming the data obtained with LA. In these two fractions, the elongation product of GLA, i.e. 20:3Δ8,11,14, could not be detected. On the other hand, both GLA and 20:3Δ8,11,14 were present in high proportions in the acyl-CoA pool. Besides the predominant 16:1, each represented more than 25% of the acyl-CoA species only 1 min after the addition of GLA, reflecting an elongation of about 50% in that pool. These data strongly suggest that upon entrance into yeast cells exogenously supplied GLA is converted into GLA-CoA and thus becomes immediately available for Δ6-elongation. Next we analyzed the distribution of GLA and 20:3Δ8,11,14 in the different lipids of a yeast culture that had expressed the Δ6-elongase in the presence of GLA for 24 h. After extraction and separation of the major
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