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• W2061943793 abstract "A pathogen-inducible oxygenase in tobacco leaves and a homologous enzyme from Arabidopsis were recently characterized (Sanz, A., Moreno, J. I., and Castresana, C. (1998) Plant Cell 10, 1523–1537). Linolenic acid incubated at 23 °C with preparations containing the recombinant enzymes underwent α-oxidation with the formation of a chain-shortened aldehyde, i.e., 8(Z),11(Z),14(Z)-heptadecatrienal (83%), an α-hydroxy acid, 2(R)-hydroxy-9(Z),12(Z),15(Z)-octadecatrienoic acid (15%), and a chain-shortened fatty acid, 8(Z),11(Z),14(Z)-heptadecatrienoic acid (2%). When incubations were performed at 0 °C, 2(R)-hydroperoxy-9(Z),12(Z),15(Z)-octadecatrienoic acid was obtained as the main product. An intermediary role of 2(R)-hydroperoxy-9(Z),12(Z),15(Z)-octadecatrienoic acid in α-oxidation was demonstrated by re-incubation experiments, in which the hydroperoxide was converted into the same α-oxidation products as those formed from linolenic acid. 2(R)-Hydroperoxy-9(Z),12(Z),15(Z)-octadecatrienoic acid was chemically unstable and had a half-life time in buffer of about 30 min at 23 °C. Extracts of cells expressing the recombinant oxygenases accelerated breakdown of the hydroperoxide (half-life time, about 3 min at 23 °C), however, this was not attributable to the recombinant enzymes since the same rate of hydroperoxide degradation was observed in the presence of control cells not expressing the enzymes. No significant discrimination between enantiomers was observed in the degradation of 2(R, S)-hydroperoxy-9(Z)-octadecenoic acid in the presence of recombinant oxygenases. A previously studied system for α-oxidation in cucumber was re-examined using the newly developed techniques and was found to catalyze the same conversions as those observed with the recombinant enzymes, i.e. enzymatic α-dioxygenation of fatty acids into 2(R)-hydroperoxides and a first order, non-stereoselective degradation of hydroperoxides into α-oxidation products. It was concluded that the recombinant enzymes from tobacco and Arabidopsis were both α-dioxygenases, and that members of this new class of enzymes catalyze the first step of α-oxidation in plant tissue. A pathogen-inducible oxygenase in tobacco leaves and a homologous enzyme from Arabidopsis were recently characterized (Sanz, A., Moreno, J. I., and Castresana, C. (1998) Plant Cell 10, 1523–1537). Linolenic acid incubated at 23 °C with preparations containing the recombinant enzymes underwent α-oxidation with the formation of a chain-shortened aldehyde, i.e., 8(Z),11(Z),14(Z)-heptadecatrienal (83%), an α-hydroxy acid, 2(R)-hydroxy-9(Z),12(Z),15(Z)-octadecatrienoic acid (15%), and a chain-shortened fatty acid, 8(Z),11(Z),14(Z)-heptadecatrienoic acid (2%). When incubations were performed at 0 °C, 2(R)-hydroperoxy-9(Z),12(Z),15(Z)-octadecatrienoic acid was obtained as the main product. An intermediary role of 2(R)-hydroperoxy-9(Z),12(Z),15(Z)-octadecatrienoic acid in α-oxidation was demonstrated by re-incubation experiments, in which the hydroperoxide was converted into the same α-oxidation products as those formed from linolenic acid. 2(R)-Hydroperoxy-9(Z),12(Z),15(Z)-octadecatrienoic acid was chemically unstable and had a half-life time in buffer of about 30 min at 23 °C. Extracts of cells expressing the recombinant oxygenases accelerated breakdown of the hydroperoxide (half-life time, about 3 min at 23 °C), however, this was not attributable to the recombinant enzymes since the same rate of hydroperoxide degradation was observed in the presence of control cells not expressing the enzymes. No significant discrimination between enantiomers was observed in the degradation of 2(R, S)-hydroperoxy-9(Z)-octadecenoic acid in the presence of recombinant oxygenases. A previously studied system for α-oxidation in cucumber was re-examined using the newly developed techniques and was found to catalyze the same conversions as those observed with the recombinant enzymes, i.e. enzymatic α-dioxygenation of fatty acids into 2(R)-hydroperoxides and a first order, non-stereoselective degradation of hydroperoxides into α-oxidation products. It was concluded that the recombinant enzymes from tobacco and Arabidopsis were both α-dioxygenases, and that members of this new class of enzymes catalyze the first step of α-oxidation in plant tissue. pathogen-inducible oxygenase 2,6-di-tert-butyl-4-methylphenol 9(S)-hydroperoxy-10(E),12(Z)-octadecadienoic acid bis(trimethylsilyl)trifluoroacetamide gas-liquid chromatography gas-liquid chromatography-mass spectrometry (−)-menthoxycarbonyl O-methyloxime trimethylsilyl reversed phase high performance liquid chromatography Fatty acid hydroperoxides serve as important intermediates in the oxylipin pathway of fatty acid oxygenation in plants and fungi (1Hamberg M. Gardner H.W. Biochim. Biophys. Acta. 1992; 1165: 1-18Crossref PubMed Scopus (224) Google Scholar, 2Mueller M.J. Physiol. Plant. 1997; 100: 653-663Crossref Google Scholar, 3Blée E. Prog. Lipid Res. 1998; 37: 33-72Crossref PubMed Scopus (255) Google Scholar, 4Grechkin A. Prog. Lipid Res. 1998; 37: 317-352Crossref PubMed Scopus (260) Google Scholar). Further metabolism of the hydroperoxide derivatives of linoleic and linolenic acids results in the formation of fatty acid epoxides and epoxy alcohols (5Hamberg M. Herman C.A. Herman R.P. Biochim. Biopys. Acta. 1986; 877: 447-457Crossref PubMed Scopus (57) Google Scholar, 6Hamberg M. Hamberg G. Arch. Biochem. Biophys. 1990; 283: 409-416Crossref PubMed Scopus (72) Google Scholar, 7Blée E. Schuber F. J. Biol. Chem. 1990; 265: 12887-12894Abstract Full Text PDF PubMed Google Scholar), dihydroxy acids (8Hamberg M. Gerwick W.H. Arch. Biochem. Biophys. 1993; 305: 115-122Crossref PubMed Scopus (49) Google Scholar, 9Brodowsky I.D. Hamberg M. Oliw E.H. J. Biol. Chem. 1992; 267: 14738-14745Abstract Full Text PDF PubMed Google Scholar), short-chain aldehydes (10Gardner H.W. Biochim. Biophys. Acta. 1991; 1084: 221-239Crossref PubMed Scopus (508) Google Scholar, 11Hatanaka A. Phytochemistry. 1993; 34: 1201-1218Crossref Scopus (557) Google Scholar), and divinyl ethers (12Galliard T. Phillips D.R. Biochem. J. 1972; 129: 743-753Crossref PubMed Scopus (98) Google Scholar, 13Proteau P.J. Gerwick W.H. Lipids. 1993; 28: 783-787Crossref PubMed Scopus (43) Google Scholar, 14Grechkin A.N. Fazliev F.N. Mukhtarova L.S. FEBS Lett. 1995; 371: 159-162Crossref PubMed Scopus (55) Google Scholar, 15Hamberg M. Lipids. 1998; 33: 1061-1071Crossref PubMed Scopus (44) Google Scholar). One specific hydroperoxide isomer, i.e. the 13(S)-hydroperoxide derivative of linolenic acid, is converted into jasmonic acid by a series of reactions catalyzed by allene oxide synthase, allene oxide cyclase, reductase, and β-oxidation enzymes (2Mueller M.J. Physiol. Plant. 1997; 100: 653-663Crossref Google Scholar). This pathway is of biological importance in plants because it produces compounds which are involved in defense reactions against insects and other phytopathogens (16Farmer E.E. Ryan C.A. Proc. Natl. Acad. Sci. U. S. A. 1990; 87: 7713-7716Crossref PubMed Scopus (1100) Google Scholar), in mechanical responses such as tendril coiling (17Weiler E.W. Albrecht T. Groth B. Xia Z.-Q. Luxem M. Liss H. Andert L. Spengler P. Phytochemistry. 1993; 32: 591-600Crossref Scopus (175) Google Scholar), and pollen development (18McConn M. Browse J. Plant Cell. 1996; 8: 403-416Crossref PubMed Google Scholar). A variety of conditions, such as mechanical perturbation, osmotic stress, attack by plant pathogens and wounding, elicit increased formation of jasmonates and other biologically active oxylipins in plant leaves (19Creelman R.A. Mullet J.E. Annu. Rev. Plant Physiol. Plant Mol. Biol. 1997; 48: 355-381Crossref PubMed Scopus (1482) Google Scholar). This is partly a consequence of liberation of free linolenic acid from its esterified forms (20Conconi A. Miquel M. Browse J.A. Ryan C.A. Plant Physiol. 1996; 111: 797-803Crossref PubMed Scopus (205) Google Scholar, 21Ryu S.B. Wang X. Biochim. Biophys. Acta. 1998; 1393: 193-202Crossref PubMed Scopus (93) Google Scholar) but may also depend on increased levels of enzymes catalyzing hydroperoxide formation and metabolism. In a recent study, tobacco leaves were found to accumulate a 75-kDa protein in response to bacterial infection (22Sanz A. Moreno J.I. Castresana C. Plant Cell. 1998; 10: 1523-1537Crossref PubMed Scopus (122) Google Scholar). This protein, as well as a protein from Arabidopsis showing a 75% homology in amino acid sequence, were expressed in insect cells and found to cause uptake of molecular oxygen in the presence of polyunsaturated fatty acids such as linolenic acid, linoleic acid, and arachidonic acid. Interestingly, the tobacco enzyme, called “pathogen-inducible oxygenase” (PIOX),1 showed significant homology to prostaglandin-endoperoxide H synthases-1 and -2 present in animal tissue (22Sanz A. Moreno J.I. Castresana C. Plant Cell. 1998; 10: 1523-1537Crossref PubMed Scopus (122) Google Scholar). The present study was carried out with the aim of identifying the catalytic function of the pathogen-induced oxygenase from tobacco leaves and its homologous enzyme from Arabidopsis. Evidence will be presented that both enzymes are fatty acid α-dioxygenases which catalyze conversion of linolenic acid and other fatty acids into their 2(R)-hydroperoxy derivatives. The mode of degradation of these unstable hydroperoxides into chain-shortened aldehydes and other α-oxidation products has also been studied. [1-14C]Linolenic, [1-14C]linoleic, and [9,10-3H2]oleic acids were purchased from NEN Life Science Products Inc. (Boston, MA). Dilution with unlabeled materials (Nu-Chek-Prep, Elysian, MN) followed by purification by SiO2 chromatography afforded specimens having specific radioactivities of 8.9, 3.8, and 184 kBq/μmol, respectively. In the same way, [9,10,12,13,15,16-3H6]linolenic acid (American Radiolabeled Chemicals, St. Louis, MO) was diluted with unlabeled linolenic acid and purified to make a specimen having a specific radioactivity of 22.2 kBq/μmol. Seeds of Thymus vulgaris (23Smith C.R. Wolff I.A. Lipids. 1969; 4: 9-14Crossref PubMed Scopus (31) Google Scholar) were ground in an electric coffee mill and the powder (25 g) was extracted under an argon atmosphere for 2 h with hexane (250 ml) containing BHT (12 ppm) in a Soxhlet apparatus. The oil (8.5 g) was subjected to methanolysis and fractionated by SiO2 open column chromatography. Elution with diethyl ether/hexane (7:93, v/v) afforded methyl 2-hydroxylinolenate (1.0 g) having a purity of 98% according to GLC analysis. Treatment with 1 m NaOH in 50 ml of 50% methanol containing BHT afforded the free acid. The identity of the material obtained with 2-hydroxylinolenic acid was confirmed by analysis of the methyl-esterified material by GC-MS. Prominent ions were observed at m/z 308 (M+; 4% relative intensity), 279 (M+ − 29; loss of ⋅CH2CH3; 1), 252 (2Mueller M.J. Physiol. Plant. 1997; 100: 653-663Crossref Google Scholar), 161 (6Hamberg M. Hamberg G. Arch. Biochem. Biophys. 1990; 283: 409-416Crossref PubMed Scopus (72) Google Scholar), 135 (12Galliard T. Phillips D.R. Biochem. J. 1972; 129: 743-753Crossref PubMed Scopus (98) Google Scholar), 108 (32Hamberg M. Fahlstadius P. Plant Physiol. 1992; 99: 987-995Crossref PubMed Scopus (40) Google Scholar), 79 (100), 67 (74), and 55 (48Namai T. Kato T. Yamaguchi Y. Hirukawa T. Biosci. Biotech. Biochem. 1993; 57: 611-613Crossref Scopus (42) Google Scholar). The mass spectrum of the Me3Si derivative of the methyl ester showed ions at m/z 380 (M+; 3%), 365 (M+ − 15; loss of ·CH3; 15), 321 (M+ − 59; loss of ⋅COOCH3; 13), 161 (Me3SiO+ = CH-COOCH3; 10), 159 (14Grechkin A.N. Fazliev F.N. Mukhtarova L.S. FEBS Lett. 1995; 371: 159-162Crossref PubMed Scopus (55) Google Scholar), 89 (Me3SiO+; 40), 79 (52), and 73 (Me3Si+; 100). The stereochemistry of the hydroxy acid, which was not unequivocally determined in the previous work (23Smith C.R. Wolff I.A. Lipids. 1969; 4: 9-14Crossref PubMed Scopus (31) Google Scholar), was established as “R” by GLC analysis of the MC derivative of the methyl ester. The percentage of the 2(S) isomer was less than 1%. 2(R,S)-Hydroxylinolenic acid was prepared by reduction of methyl 2-ketolinolenate (see below) with NaBH4 followed by saponification. The methyl ester of 2(R)-hydroxylinolenic acid (31 mg) was oxidized with chromium trioxide-pyridine complex in methylene chloride (24Ratcliffe R. Rodehurst R. J. Org. Chem. 1970; 35: 4000-4002Crossref Scopus (821) Google Scholar). Purification by open column SiO2 chromatography afforded methyl 2-ketolinolenate (12 mg) having a purity in excess of 98%. The mass spectrum showed prominent ions at m/z 306 (M+; 3%), 250 (2Mueller M.J. Physiol. Plant. 1997; 100: 653-663Crossref Google Scholar), 247 (M+ − 59; loss of⋅COOCH3; 2), 187 (3Blée E. Prog. Lipid Res. 1998; 37: 33-72Crossref PubMed Scopus (255) Google Scholar), 159 (7Blée E. Schuber F. J. Biol. Chem. 1990; 265: 12887-12894Abstract Full Text PDF PubMed Google Scholar), 93 (50), 79 (100), 67 (81), and 55 (58). The Fourier transform-infrared spectrum (film) showed absorption bands at 1754 and 1732 cm−1 due to the ester and keto carbonyls, respectively. 2(R)-Hydroxylinolenic acid (400 mg) was treated under an argon atmosphere with sodium periodate (1.5 g) in acetone (40 ml) containing glacial acetic acid (20 ml) and water (10 ml) at 50 °C for 21 h (cf. Ref.25Yanuka Y. Katz R. Sarel S. Tetrahedron Lett. 1968; 14: 1725-1728Crossref PubMed Scopus (25) Google Scholar). The product obtained by extraction with light petroleum (about 80% of aldehyde and 15% of unoxidized hydroxy acid) was subjected to SiO2 open column chromatography. Elution with diethyl ether/hexane (5:95, v/v) afforded 8(Z),11(Z),14(Z)-heptadecatrienal (200 mg; purity, 98%). A faint odor of fresh seaweed was noted for the sample. The Fourier transform-infrared spectrum (film) showed absorption bands at inter alia 1727 cm−1(aldehyde carbonyl) and 2715 cm−1 (C-H stretching in aldehyde group). The mass spectrum showed prominent ions at m/z 248 (M+; 2%), 219 (1Hamberg M. Gardner H.W. Biochim. Biophys. Acta. 1992; 1165: 1-18Crossref PubMed Scopus (224) Google Scholar), 192 (3Blée E. Prog. Lipid Res. 1998; 37: 33-72Crossref PubMed Scopus (255) Google Scholar), 135 (6Hamberg M. Hamberg G. Arch. Biochem. Biophys. 1990; 283: 409-416Crossref PubMed Scopus (72) Google Scholar), 121 (8Hamberg M. Gerwick W.H. Arch. Biochem. Biophys. 1993; 305: 115-122Crossref PubMed Scopus (49) Google Scholar), 108 (23Smith C.R. Wolff I.A. Lipids. 1969; 4: 9-14Crossref PubMed Scopus (31) Google Scholar), 93 (45Landino L.M. Crews B.C. Gierse J.K. Hauser S.D. Marnett L.J. J. Biol. Chem. 1997; 272: 21565-21574Abstract Full Text Full Text PDF PubMed Scopus (60) Google Scholar), 79 (100), 67 (79), and 55 (48Namai T. Kato T. Yamaguchi Y. Hirukawa T. Biosci. Biotech. Biochem. 1993; 57: 611-613Crossref Scopus (42) Google Scholar). The O-methyloxime (MO) derivative showed two peaks on GC-MS analysis due to syn/anti isomerism. The mass spectra recorded on these peaks were virtually identical and showed prominent ions at 277 (M+, 0.5%), 262 (M+ − 15; loss of⋅CH3; 1), 246 (M+ − 31; loss of⋅OCH3; 21), 190 (7Blée E. Schuber F. J. Biol. Chem. 1990; 265: 12887-12894Abstract Full Text PDF PubMed Google Scholar), 166 (7Blée E. Schuber F. J. Biol. Chem. 1990; 265: 12887-12894Abstract Full Text PDF PubMed Google Scholar), 108 (23Smith C.R. Wolff I.A. Lipids. 1969; 4: 9-14Crossref PubMed Scopus (31) Google Scholar), 93 (42Shine W.E. Stumpf P.K. Arch. Biochem. Biophys. 1974; 162: 147-157Crossref PubMed Scopus (73) Google Scholar), 79 (100), 67 (75), and 55 (52). Mixed fatty acid methyl esters prepared from the seed oil of T. vulgaris (see above) were saponified and subjected to RP-HPLC using solvent system II. 8(Z),11(Z),14(Z)-Heptadecatrienoic acid (nor-linolenic acid, Ref. 23Smith C.R. Wolff I.A. Lipids. 1969; 4: 9-14Crossref PubMed Scopus (31) Google Scholar) appeared with an elution volume of 37.9 ml (linolenic acid, 54.7 ml). The mass spectrum of the methyl ester showed prominent ions at m/z 278 (M+; 7%), 247 (M+ − 31; loss of ⋅OCH3; 2), 222 (3Blée E. Prog. Lipid Res. 1998; 37: 33-72Crossref PubMed Scopus (255) Google Scholar), 177 (3Blée E. Prog. Lipid Res. 1998; 37: 33-72Crossref PubMed Scopus (255) Google Scholar), 149 (9Brodowsky I.D. Hamberg M. Oliw E.H. J. Biol. Chem. 1992; 267: 14738-14745Abstract Full Text PDF PubMed Google Scholar), 135 (10Gardner H.W. Biochim. Biophys. Acta. 1991; 1084: 221-239Crossref PubMed Scopus (508) Google Scholar), 121 (17Weiler E.W. Albrecht T. Groth B. Xia Z.-Q. Luxem M. Liss H. Andert L. Spengler P. Phytochemistry. 1993; 32: 591-600Crossref Scopus (175) Google Scholar), 108 (27Galliard T. Matthew J.A. Biochim. Biophys. Acta. 1976; 424: 26-35Crossref PubMed Scopus (55) Google Scholar), 93 (48Namai T. Kato T. Yamaguchi Y. Hirukawa T. Biosci. Biotech. Biochem. 1993; 57: 611-613Crossref Scopus (42) Google Scholar), 79 (100), 67 (72), and 55 (49). 2-Hydroperoxyoleic acid and 2-hydroperoxypalmitic acid were prepared by modification of a method previously described (26Konen D.A. Silbert L.S. Pfeffer P.E. J. Org. Chem. 1975; 40: 3253-3258Crossref Scopus (76) Google Scholar). For preparation of the first mentioned compound, butyllithium (0.8 mmol) was added at −78 °C to an argon-purged solution of diisopropylamine (0.1 ml) in 1.5 ml of dry tetrahydrofuran. A solution of [9,10-3H2]oleic acid (111 mg; 25 MBq) and hexamethylphosphoric triamide (63 μl) in 0.25 ml of tetrahydrofuran was added and the mixture was kept under argon at 50 °C for 30 min. The material was added under a period of 30 min to 25 ml of oxygen-saturated diethyl ether at −78 °C. Oxygen gas was continuously bubbled into the reaction mixture in order to maintain oxygen saturation and to effect rapid dispersion of the introduced oleate dianion. Material obtained by extraction with diethyl ether was analyzed by RP-HPLC using solvent system IV. Three main peaks were observed, i.e. 2-hydroperoxyoleic acid (13.0 ml effluent), 2-hydroxyoleic acid (13.8 ml), and oleic acid (27.1 ml). Effluent containing 2-hydroperoxyoleic acid was collected and rapidly extracted with diethyl ether. The ether phase was dried over MgSO4and taken to dryness in vacuo. The residual 2-hydroperoxyoleic acid was dissolved in dry acetone and stored at −25 °C. Analysis of the specimen by radio-HPLC showed a radiochemical purity in excess of 95%. The specific radioactivity was 32 kBq/μmol (the drop in specific activity compared with that of the starting material was due to a slight chromatographic separation between ditritio- and unlabeled molecules in the RP-HPLC purification step resulting in partial loss of the 3H2compound). As expected, the 2-hydroperoxyoleic acid was reducible into 2-hydroxyoleic acid by treatment with SnCl2 or triphenylphosphine. Furthermore, treatment of the hydroperoxide with sodium borodeuteride afforded 2-hydroxyoleic acid with no detectable incorporation of deuterium. An aliquot of the hydroperoxide was treated with BSTFA and analyzed by GC-MS. Considerable degradation with formation of 8(Z)-heptadecenal (30%, 6.3 min retention time), the Me3Si ester of 2-ketooleic acid (9%, 10.2 min), the Me3Si ether/ester of 2-hydroxyoleic acid (18%, 10.7 min), the Me3Si ether/ester of the enol form of 2-ketooleic acid (16%, 10.9 min) took place, however, a portion of the hydroperoxide derivative (27%) chromatographed as the intact Me3Si peroxide and appeared as a peak at 11.3 min. The mass spectrum recorded on this material showed prominent ions at m/z 443 (M+ − 15; loss of⋅CH3; 1%), 426 (M+ − 32; rearrangement with loss of CH3OH; 9), 341 (M+ − 117; elimination of ⋅COOSiMe3; 6), 251 (341-90; 3), 163 (Me3Si-O-O+ = SiMe2; 13), 147 (Me3Si-O+ = SiMe2; 27), 89 (Me3SiO+; 53), and 73 (Me3Si+; 100). Recombinant oxygenases were obtained from insect cells infected with baculovirus carrying pFASTBAC (vector), pFASTBAC-tob.A5.2, or pFASTBAC-ara.N38086 (22Sanz A. Moreno J.I. Castresana C. Plant Cell. 1998; 10: 1523-1537Crossref PubMed Scopus (122) Google Scholar). Cell suspensions in 1–2 ml of sonication buffer (22Sanz A. Moreno J.I. Castresana C. Plant Cell. 1998; 10: 1523-1537Crossref PubMed Scopus (122) Google Scholar) were sonicated at 0 °C by four bursts of 10 s and subsequently centrifuged at 12,000 ×g for 6 min. The supernatants (protein, 1–2 mg/ml) were diluted with 0.1 m potassium phosphate buffer, pH 6.7 or 7.4, and directly used for the incubations. Two crude preparations of the α-oxidation system of cucumber (27Galliard T. Matthew J.A. Biochim. Biophys. Acta. 1976; 424: 26-35Crossref PubMed Scopus (55) Google Scholar, 28Baardseth P. Slinde E. Thomassen M.S. Biochim. Biophys. Acta. 1987; 922: 170-176Crossref Scopus (13) Google Scholar, 29Andersen Borge G.I. Slinde E. Nilsson A. Biochim. Biophys. Acta. 1997; 1344: 47-58Crossref PubMed Scopus (11) Google Scholar) were obtained in the following way. Cucumber fruits were peeled and the flesh and seed parts (175 g) were diced and added to 0.1 m potassium phosphate buffer, pH 6.7 or 7.4 (175 ml). The tissue was homogenized at 0 °C for two periods of 60 s using an Ultra-Turrax and subsequently filtered through gauze. The filtrate (protein, 0.6–0.7 mg/ml) was directly used for small scale incubations carried with various fatty acid substrates. For larger scale incubations carried out in order to prepare 2-hydroperoxy acids, the filtrate was centrifuged for 15 min at 1,100 × g. The supernatant was decanted and further centrifuged at 48,000 × g for 30 min. The sediment fraction was either suspended in potassium phosphate buffer (140 ml; protein, 0.2–0.3 mg/ml) and used for the incubations or frozen and stored at −25 °C. The 105,000 × g particle fraction of homogenate of seeds of Vicia faba L. was prepared as described previously (6Hamberg M. Hamberg G. Arch. Biochem. Biophys. 1990; 283: 409-416Crossref PubMed Scopus (72) Google Scholar). Suspensions of this material in 0.1 m potassium phosphate buffer, pH 6.7 (protein, 0.6 mg/ml), were used as a source of peroxygenase (6Hamberg M. Hamberg G. Arch. Biochem. Biophys. 1990; 283: 409-416Crossref PubMed Scopus (72) Google Scholar). Incubations of the tobacco and Arabidopsis oxygenases, and of whole homogenate of cucumber, were carried out with 50–250 μm fatty acid at 23 or 0 °C for the times indicated. The mixtures were diluted with 1 volume of distilled water, acidified to pH 4, and extracted twice with diethyl ether. The combined ether phases were washed with water and taken to dryness in vacuo. In most incubations, the material obtained was immediately dissolved in HPLC mobile phase, centrifuged, and analyzed by RP-radio-HPLC. For incubations with peroxygenase, suspensions (0.5 ml) of the membrane fraction from V. faba seeds were preincubated at 23 °C for 5 min with the lipoxygenase inhibitor 5,8,11,14-eicosatetraynoic acid (50 μm). Subsequently, [9,10-3H2]oleic acid (100 μm) and hydroperoxide (30–264 μm) were added and stirring continued for 15 min. The reaction products were extracted with diethyl ether, and the material obtained was analyzed by RP-radio-HPLC. Linolenic acid (5.9 mg; concentration, 152 μm) was stirred at 0 °C for 30 min with a suspension (140 ml) of the 48,000 × g particle fraction of homogenate of cucumber. The mixture was acidified to pH 4 and rapidly extracted with 2 volumes of diethyl ether. The material obtained following evaporation of the solvent was suspended in HPLC mobile phase (0.4 ml). After centrifugation, aliquots of 0.1 ml were subjected to RP-HPLC using solvent system I at a flow rate of 2 ml/min. Effluent containing the hydroperoxide (37.0–39.4 ml) was immediately extracted with diethyl ether and the solution dried over MgSO4. Hydroperoxide obtained from several such incubations was dissolved in 0.5 ml of dry acetone (concentration, 10 mm) and stored at −25 °C. The yield of hydroperoxide from the incubated linolenic acid was 5–10% and the radiochemical purity was in excess of 95%. The identity of the hydroperoxide with 2(R)-hydroperoxy-9(Z),12(Z),15(Z)-octadecatrienoic acid was based on chemical and spectral analyses as described under “Results.” The rate of breakdown of 2-hydroperoxides in enzyme preparations or in buffer was determined by using one of two methods. In method A, tritium-labeled 2(R)-hydroperoxylinolenic acid (35 μm) was stirred at 23 °C with enzyme preparation or buffer (0.8 or 1 ml). At different times of stirring, the sample was directly subjected to RP-radio-HPLC using a column protected with a pre-column and a solvent system consisting of acetonitrile/water (80:20, v/v) at a flow rate of 2 ml/min. Remaining hydroperoxide was eluted as its salt (3.6–6.0 ml effluent), well separated from the main product of hydroperoxide breakdown, i.e. 8(Z),11(Z),14(Z)-heptadecatrienal (16.8–19.2 ml effluent). The rate of hydroperoxide breakdown was estimated by plotting the integrated radioactivity associated with the peak of unconverted hydroperoxide versus time. When 2-hydroperoxylinolenic acid was allowed to degrade in the presence of enzyme preparations, a portion of the product consisted of 2-hydroxylinolenic acid and nor-linolenic acid. These compounds are expected to elute as their salts together with the hydroperoxide, however, because of the small and variable amounts of these decomposition products (together 10% or less), no attempt was made to correct for their presence. In method B, 2(R,S)-hydroperoxyoleic acid (30 μm) was stirred with the test preparation (6 ml) at 23 °C. Aliquots of 1 ml were removed at different times and added to 5 ml of ethanol containing 25 mg of stannous chloride. After 20 min at 23 °C, an internal standard of tetracosanoic acid (70 nmol) was added and the mixtures were extracted with diethyl ether. Aliquots of the methyl-esterified product was subjected to GLC (column temperature, 270 °C) and the peak areas of methyl 2-hydroxyoleate (retention time, 3.6 min) formed by reduction of 2-hydroperoxyoleic acid remaining in the incubation mixture, and of methyl tetracosanoate (retention time, 7.8 min) due to the added internal standard were determined. The rate of hydroperoxide breakdown was calculated from plots of the ratio between the peak areas of methyl 2-hydroxyoleate and methyl tetracosanoate versus time. As with method A, no attempt was made to correct for the small amount of 2-hydroxy acid produced from the hydroperoxide during the incubation period. In some experiments, the reduced samples containing methyl 2-hydroxyoleate were derivatized with (−)-menthoxycarbonyl chloride, purified by TLC, and subjected to GC-MS operated in the selected ion monitoring mode using the ions m/z 294 and 262. By combining the peak areas of the MC derivatives of methyl 2(S)-hydroxyoleate (retention time, 13.15 min) and methyl 2(R)-hydroxyoleate (13.37 min) with the half-life data, it was possible to separately monitor breakdown of the “R” and “S” enantiomers of 2-hydroperoxyoleate. Configurational determination of 2-hydroxy acids were performed by analysis of MC derivatives by GLC or GC-MS (30Hamberg M. Anal. Biochem. 1971; 43: 515-526Crossref PubMed Scopus (219) Google Scholar). MO derivatives of carbonyl compounds and Me3Si ethers of hydroxy compounds were prepared as described previously (31Hamberg M. Eur. J. Biochem. 1968; 6: 136-146Google Scholar). [2H9]Me3Si derivatives, occasionally needed to verify correct interpretation of mass spectra, were prepared by derivatization with [2H18]N,O-bis(trimethylsilyl)acetamide (98%, Cambridge Isotope Laboratories, Andover, MA) at 23 °C for 30 min. For analysis of 2-hydroperoxy acids by GC-MS, the hydroperoxides (10–50 μg) were derivatized with BSTFA (0.1 ml) and an aliquot of 1–2 μl containing the Me3Si peroxide/Me3Si ester was directly injected onto the column. Hydroperoxides were reduced into alcohols by treatment with SnCl2 in ethanol (5 mg/ml) at room temperature for 10 min, or with triphenyl phosphine in diethyl ether (10 mg/ml) at room temperature for 1 h. Catalytic hydrogenation was performed with platinum catalyst (3 mg) and methanol (1 ml) as the solvent. Oxidative ozonolysis was carried out as described (30Hamberg M. Anal. Biochem. 1971; 43: 515-526Crossref PubMed Scopus (219) Google Scholar) using an ozone generator model T-12 purchased from TriO3 Industries, Fort Pierce, FL. Incubations under 18O gas were conducted in an all-glass apparatus attached to a high vacuum line.18O2 (isotopic purity, 96%) was obtained from Larodan AB, Malmö, Sweden. RP-radio-HPLC was performed with columns of Nucleosil 100-5 C18 (250 × 4.6 mm) purchased from Macherey-Nagel (Düren, Germany). The solvent systems consisted of mixtures of acetonitrile, water, 2m hydrochloric acid in volume proportions 55:45:0.013 (system I), 60:40:0.013 (system II), 65:35:0.013 (system III), or 80:20:0.013 (system IV). The absorbance (210 nm) and radioactivity of HPLC effluents were determined on-line using a Spectromonitor III ultraviolet detector (Laboratory Data Control, Riviera Beach, FL) and a liquid scintillation counter (IN/US Systems, Tampa, FL), respectively. GLC was performed with a Hewlett-Packard (Avondale, PA) model 5890 gas chromatograph equipped with a methylsilicone capillary column (length, 25 m; film thickness, 0.33 μm) and a flame ionization detector. Helium at a flow rate of 25 cm/s was used as the carrier gas. Retention times found on GLC were converted into C-values as described (31Hamberg M. Eur. J. Biochem. 1968; 6: 136-146Google Scholar). GC-MS was carried out with a Hewlett-Packard model 5970B mass selective detector connected to a Hewlett-Packard model 5890 gas chromatograph fitted with a 5% phenylmethylsilicone capillary column (length, 12 m; film thickness, 0.33 μm). In most runs the initial column temperature was 120 °C and raised at 10 °C/min until 240 °C. Ultraviolet spectra were recorded with a Hitachi (Tokyo, Japan) model" @default.
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• W2061943793 title "α-Oxidation of Fatty Acids in Higher Plants" @default.
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