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• W2012194536 abstract "To create the unique properties of a certain cellular membrane, both the composition and the metabolism of membrane phospholipids are key factors. Phospholipase A2(PLA2), with hydrolytic enzyme activities at thesn-2 position in glycerophospholipids, plays critical roles in maintaining the phospholipid composition as well as producing bioactive lipid mediators. In this study we examined the contribution of a Ca2+-independent group IVC PLA2 isozyme (cPLA2γ), a paralogue of cytosolic PLA2α (cPLA2α), to phospholipid remodeling. The enzyme was localized in the endoplasmic reticulum and Golgi apparatus, as seen using green fluorescence fusion proteins. Electrospray ionization mass spectrometric analysis of membrane extracts revealed that overexpression of cPLA2γ increased the proportion of polyunsaturated fatty acids in phosphatidylethanolamine, suggesting that the enzyme modulates the phospholipid composition. We also found that H2O2 and other hydroperoxides induced arachidonic acid release in cPLA2γ-transfected human embryonic kidney 293 cells, possibly through the tyrosine phosphorylation pathway. Thus, we propose that cPLA2γ is constitutively expressed in the endoplasmic reticulum and plays important roles in remodeling and maintaining membrane phospholipids under various conditions, including oxidative stress. To create the unique properties of a certain cellular membrane, both the composition and the metabolism of membrane phospholipids are key factors. Phospholipase A2(PLA2), with hydrolytic enzyme activities at thesn-2 position in glycerophospholipids, plays critical roles in maintaining the phospholipid composition as well as producing bioactive lipid mediators. In this study we examined the contribution of a Ca2+-independent group IVC PLA2 isozyme (cPLA2γ), a paralogue of cytosolic PLA2α (cPLA2α), to phospholipid remodeling. The enzyme was localized in the endoplasmic reticulum and Golgi apparatus, as seen using green fluorescence fusion proteins. Electrospray ionization mass spectrometric analysis of membrane extracts revealed that overexpression of cPLA2γ increased the proportion of polyunsaturated fatty acids in phosphatidylethanolamine, suggesting that the enzyme modulates the phospholipid composition. We also found that H2O2 and other hydroperoxides induced arachidonic acid release in cPLA2γ-transfected human embryonic kidney 293 cells, possibly through the tyrosine phosphorylation pathway. Thus, we propose that cPLA2γ is constitutively expressed in the endoplasmic reticulum and plays important roles in remodeling and maintaining membrane phospholipids under various conditions, including oxidative stress. phospholipase A2 arachidonic acid bovine serum albumin Chinese hamster ovary cytosolic PLA2α (group IVA phospholipase A2) group IVC phospholipase A2 Dulbecco's modified Eagle's medium endoplasmic reticulum electrospray ionization mass spectrometry green fluorescence protein human embryonic kidney 293 Ca2+-independent phospholipase A2 methyl arachidonyl fluorophosphonate mitogen-activated protein phosphatidylcholine phosphatidylethanolamine protein kinase C protein tyrosine kinase polyunsaturated fatty acid reactive oxygen species Biological membranes contain a complicated mixture of phospholipids differing from each other with respect to their head-group structure, hydrocarbon chain length, and degree of unsaturation of the acyl chains. The complexity of these phospholipid structures results in their diverse roles in membrane dynamics, protein regulation, signal transduction, and vesicular secretion. The assembly of membranes requires the coordinate synthesis, catabolism, and transport of phospholipids to create the unique properties of a certain cellular membrane. Considerable numbers of studies have identified many enzymes involved in these multiple pathways (1Dircks L. Sul H.S. Prog. Lipid Res. 1999; 38: 461-479Google Scholar, 2Farooqui A.A. Horrocks L.A. Farooqui T. Chem. Phys. Lipids. 2000; 106: 1-29Google Scholar, 3Lands W.E. Biochim. Biophys. Acta. 2000; 1483: 1-14Google Scholar, 4MacDonald J.I. Sprecher H. Biochim. Biophys. Acta. 1991; 1084: 105-121Google Scholar, 5Ohvo-Rekila H. Ramstedt B. Leppimaki P. Slotte J.P. Prog. Lipid Res. 2002; 41: 66-97Google Scholar, 6Tabas I. Biochim. Biophys. Acta. 2000; 1529: 164-174Google Scholar, 7Yamashita A. Sugiura T. Waku K. J. Biochem. (Tokyo). 1997; 122: 1-16Google Scholar, 8Chilton F.H. Fonteh A.N. Surette M.E. Triggiani M. Winkler J.D. Biochim. Biophys. Acta. 1996; 1299: 1-15Google Scholar). Phospholipase A2(PLA2)1 is a superfamily of enzymes that hydrolyze the sn-2 ester bond in glycerophospholipids, releasing free fatty acids and lysophospholipids (9Six D.A. Dennis E.A. Biochim. Biophys. Acta. 2000; 1488: 1-19Google Scholar, 10Murakami M. Kudo I. J. Biochem. (Tokyo). 2002; 131: 285-292Google Scholar, 11Cho W. Biochim. Biophys. Acta. 2000; 1488: 48-58Google Scholar). To date, at least 19 enzymes have been identified in mammals. Among them, cPLA2α is the only PLA2enzyme that shows significant selectivity toward phospholipids containing arachidonic acid (AA) at the sn-2 position. A number of studies have clarified its structure (12Dessen A. Tang J. Schmidt H. Stahl M. Clark J.D. Seehra J. Somers W.S. Cell. 1999; 97: 349-360Google Scholar), localization (13Glover S. de Carvalho M.S. Bayburt T. Jonas M. Chi E. Leslie C.C. Gelb M.H. J. Biol. Chem. 1995; 270: 15359-15367Google Scholar, 14Schievella A.R. Regier M.K. Smith W.L. Lin L.L. J. Biol. Chem. 1995; 270: 30749-30754Google Scholar, 15Hirabayashi T. Kume K. Hirose K. Yokomizo T. Iino M. Itoh H. Shimizu T. J. Biol. Chem. 1999; 274: 5163-5169Google Scholar), and pathophysiological roles in vivo (16Uozumi N. Kume K. Nagase T. Nakatani N. Ishii S. Tashiro F. Komagata Y. Maki K. Ikuta K. Ouchi Y. Miyazaki J. Shimizu T. Nature. 1997; 390: 618-622Google Scholar, 17Sapirstein A. Bonventre J.V. Biochim. Biophys. Acta. 2000; 1488: 139-148Google Scholar, 18Bonventre J.V. Huang Z. Taheri M.R. O'Leary E. Li E. Moskowitz M.A. Sapirstein A. Nature. 1997; 390: 622-625Google Scholar). In contrast, the information on cPLA2γ, a Ca2+-independent paralogue of cPLA2α, is limited. It has been reported that cPLA2γ is predominantly expressed in the brain, heart, and skeletal muscle in humans (19Pickard R.T. Strifler B.A. Kramer R.M. Sharp J.D. J. Biol. Chem. 1999; 274: 8823-8831Google Scholar, 20Underwood K.W. Song C. Kriz R.W. Chang X.J. Knopf J.L. Lin L.L. J. Biol. Chem. 1998; 273: 21926-21932Google Scholar) and that the enzyme is activated in vivo by serum (21Stewart A. Ghosh M. Spencer D.M. Leslie C.C. J. Biol. Chem. 2002; 277: 29526-29536Google Scholar). However, the fatty acid selectivity is controversial, and its subcellular localization, cellular roles, and regulation of activity have not been documented. In this study we demonstrated that cPLA2γ is localized in the endoplasmic reticulum (ER) and Golgi and is involved in the mobilization of fatty acids in phosphatidylethanolamine (PE). During the search for regulators of the enzymatic activity at the cellular level, we found that reactive oxygen species (ROS) activate the activity of cPLA2γ, possibly through the tyrosine phosphorylation pathway. [1-14C]Arachidonic acid (2.1 GBq/mmol) was from Amersham Biosciences. Bovine serum albumin (BSA, fatty acid-free) and catalase (Aspergillus niger) were from Sigma. Herbimycin A, calphostin C, and sodium orthovanadate were obtained from Calbiochem, and methyl arachidonyl fluorophosphonate (MAFP) was from Cayman Chemical Co. (Ann Arbor, MI). Hydrogen peroxide (H2O2) was from Wako (Osaka, Japan). BODIPY Brefeldin A and MitoTracker Red CMXRos were from Molecular Probes (Eugene, OR). Mammalian expression vector pcDNA3.1/His was obtained from Invitrogen, and pEGFP-C1 was from Clontech. The mammalian expression vector pcDNA3.1/His was used to express proteins fused with the N-terminal Xpress epitope. The cDNA fragments encoding full-length human cPLA2γ were inserted into pcDNA3.1/His to obtain pcDNA3.1/His-cPLA2γ. For confocal microscopy studies, cPLA2γ was subcloned into pEGFP-C1, encoding a green fluorescence protein (GFP) at the N terminus between theBglII and SmaI sites to obtain pEGFP-cPLA2γ. HEK293 cells were cultured in Dulbecco's modified Eagle's medium (DMEM; Sigma) supplemented with 10% fetal calf serum (Sigma), 100 IU/ml penicillin, and 100 μg/ml streptomycin. CHO cells were cultured in Nutrient Mixture F-12 HAM (Sigma) supplemented with 10% fetal calf serum, 100 IU/ml penicillin, and 100 μg/ml streptomycin. LipofectAMINE PLUS (Invitrogen) was used for the transfection of HEK293 and CHO cells according to the manufacturer's protocols. To obtain HEK293 cells stably expressing cPLA2γ, the cells were transfected with pcDNA3.1/His-cPLA2γ, selected with 600 μg/ml G418 (Invitrogen), and maintained in the presence of 300 μg/ml G418. The expression of cPLA2γ was confirmed by immunoblotting. HEK293 cells transfected with the vector alone were also kept in medium with G418 and used as a vector control. CHO cells were seeded on coverslips (10-mm diameter) on glass-bottomed culture dishes (Matsunami, Osaka, Japan) at a density of 1.5 × 104 cells/coverslip, transiently transfected with 1 μg of the expression vector encoding a GFP-cPLA2γ fusion protein (pEGFP-cPLA2γ) or a control vector, pEGFP-C1, with LipofectAMINE PLUS according to the manufacturer's protocol, and used for experiments 48 h after transfection. For double staining, cells transiently expressing GFP-cPLA2γ were incubated with 1 μm BODIPY Brefeldin A (λex = 558 nm, λem = 568 nm) or 100 nm MitoTracker Red CMXRos (λex = 579 nm, λem = 599 nm) in Hanks' balanced salt solution containing 10 mm HEPES and 0.1% BSA for 20 min at 37 °C. The fluorescent signal was observed with an AX-80 analytical microscope system (Olympus, Tokyo, Japan) or a Zeiss LSM510 Laser Scanning Microscope using a Zeiss 100 × 1.3 NA oil immersion lens. About 1 × 107 cells were collected, washed three times with phosphate-buffered saline (5 mm sodium phosphate buffer, pH 7.4, containing 150 mm NaCl), and then suspended in 1 ml of phosphate-buffered saline. The total phospholipids were extracted by the method of Bligh and Dyer (22Bligh E.G. Dyer W.J. Can. J. Biochem. Physiol. 1959; 37: 911-918Google Scholar). The total lipid extract was dried under a gentle stream of nitrogen, dissolved in 400 μl of chloroform/methanol (2:1, v/v), and used for mass spectrometry analysis. Mass spectrometry analysis was performed essentially as described previously (23Taguchi R. Hayakawa J. Takeuchi Y. Ishida M. J. Mass Spectrom. 2000; 35: 953-966Google Scholar). Lipid extracts from the cells were analyzed using a Quattro II tandem quadrupole mass spectrometer (Micromass, Manchester, UK) equipped with an electrospray ion source. 5 μl of samples (500 pmol/μl) dissolved in chloroform/methanol (2:1, v/v) were introduced by means of a flow injector into the ESI chamber at a flow rate of 2 μl/min. The elution solvent was acetonitrile/methanol/water (2:3:1, v/v/v) containing 0.1% ammonium formate (pH 6.4). The mass spectrometer was operated in the positive and negative ion scan modes. The nitrogen drying gas flow rate was 10 liters/min, and its temperature was 80 °C. Essentially, the capillary voltage was set at 3.7 kV, and the cone voltage was set at 30 V in both the positive and negative ion scan modes. Cells were seeded onto 12-well culture plates at a density of 8 × 104 cells/well in DMEM. After 24 h of incubation the medium was removed, and the cells were labeled by incubation for 24 h in 1 ml of DMEM containing 0.05 μCi of [14C]AA and 0.2% (w/v) fatty acid-free BSA. The cells were then washed three times with DMEM containing 0.2% (w/v) fatty acid-free BSA (the control medium) and stimulated with ligands in the same buffer at 37 °C. The radioactivity of the supernatants and cell lysates (in 1% Triton X-100) was measured using a liquid scintillation counter. The amount of radioactivity released into the supernatant was expressed as a percentage of the total incorporated radioactivity. To observe the subcellular localization of cPLA2γ, CHO cells were transiently transfected with a vector coding for cPLA2γ fused to GFP (GFP-cPLA2γ), and the distribution of the chimeric proteins was examined. GFP-cPLA2γ was located in the nuclear envelope and in an extensive reticular pattern that was typical for ER localization (Fig. 1 A). These structures were identified to be ER and Golgi by loading the cells with BODIPY Brefeldin A, which recognizes both ER and Golgi (24Deng Y. Bennink J.R. Kang H.C. Haugland R.P. Yewdell J.W. J. Histochem. Cytochem. 1995; 43: 907-915Google Scholar) (Fig. 1 B). GFP-cPLA2γ did not colocalize with a mitochondrial marker, MitoTracker Red (Fig. 1 B). We also confirmed the absence of cPLA2γ in the lysosome using a lysosomal marker (data not shown). Although we have previously shown different subcellular localization of GFP-cPLA2γ in a preliminary experiment (56Hirabayashi T. Shimizu T. Biochim. Biophys. Acta. 2000; 1488: 124-138Google Scholar), the localization turned out to be caused by a peroxisomal targeting signal generated incidentally in the linker region of the chimeric proteins. Western blotting analysis of protein extracts from cPLA2γ-overexpressing HEK293 cells confirmed the assignment of the localization of cPLA2γ to the microsomal fraction (data not shown), in accordance with the results reported previously for cPLA2γ expressed in CHO cells (19Pickard R.T. Strifler B.A. Kramer R.M. Sharp J.D. J. Biol. Chem. 1999; 274: 8823-8831Google Scholar) and Sf9 (20Underwood K.W. Song C. Kriz R.W. Chang X.J. Knopf J.L. Lin L.L. J. Biol. Chem. 1998; 273: 21926-21932Google Scholar). These results suggest that cPLA2γ is localized predominantly in the ER and Golgi membranes. Although the enzymatic properties of cPLA2γ in vitro have previously been characterized (19Pickard R.T. Strifler B.A. Kramer R.M. Sharp J.D. J. Biol. Chem. 1999; 274: 8823-8831Google Scholar, 20Underwood K.W. Song C. Kriz R.W. Chang X.J. Knopf J.L. Lin L.L. J. Biol. Chem. 1998; 273: 21926-21932Google Scholar, 21Stewart A. Ghosh M. Spencer D.M. Leslie C.C. J. Biol. Chem. 2002; 277: 29526-29536Google Scholar), its roles at the cellular level have not been clarified. First, we hypothesized that cPLA2γ might change the phospholipid composition in the membrane, because the enzyme is membrane-bound, possesses Ca2+-independent PLA2 activity (19Pickard R.T. Strifler B.A. Kramer R.M. Sharp J.D. J. Biol. Chem. 1999; 274: 8823-8831Google Scholar, 20Underwood K.W. Song C. Kriz R.W. Chang X.J. Knopf J.L. Lin L.L. J. Biol. Chem. 1998; 273: 21926-21932Google Scholar, 21Stewart A. Ghosh M. Spencer D.M. Leslie C.C. J. Biol. Chem. 2002; 277: 29526-29536Google Scholar), and is localized in the ER (Fig. 1) where phospholipid biosynthesis occurs. To test this possibility and also to determine intracellular substrates for cPLA2γ, we analyzed the molecular species of phospholipids in HEK293 cells stably expressing cPLA2γ or control cells by ESI-MS. Examination of the PE molecular species in the negative-ion mode demonstrated a relative abundance of polyunsaturated fatty acid (PUFA) species such as 1-alkyl or alkenyl-2-acyl 16:1–20:4 (m/z 722.6) and diacyl 18:0–20:4 (m/z 766.7) in cPLA2γ-overexpressing HEK293 cells. In contrast, saturated or monounsaturated fatty acid species such as diacyl 16:0–16:1 (m/z 688.6), diacyl 16:0–18:1 (m/z 716.6) and diacyl 18:0–18:0 (m/z 746.7) were unchanged or rather decreased in cPLA2γ-expressing HEK293 cells (Fig.2 A). On the other hand, analysis of the phosphatidylcholine (PC) molecular species in the positive ion mode demonstrated no major difference between the two cell lines (Fig. 2 B). To search for stimuli which regulate cPLA2γ activity at the cellular level, we examined the effect of various compounds on AA release and found that H2O2 enhanced AA release. As shown in Fig.3 A, 1 mmH2O2 enhanced AA release from cells in a cPLA2γ-dependent manner. Although the effect was also detected in control cells, much higher release was observed in cPLA2γ-overexpressing cells. Addition of glucose and glucose oxidase to the medium also enhanced AA release from cells expressing cPLA2γ (data not shown). These effects were completely inhibited by adding catalase to the reaction medium. Other ROS, such as cumene hydroperoxide, enhanced the H2O2-induced AA release in a cPLA2γ-dependent manner (data not shown). 4-Hydroxy 2-nonenal, a major oxidized product of fatty acids or hydroxy radicals produced by the combination of Fe2+ and H2O2, did not induce AA release (data not shown). The effect of H2O2 on AA release was evident at 5 min and lasted for at least 30 min (data not shown). Under the assay conditions, no significant loss or floating of cells from the plates were seen (data not shown). Similar results were obtained in three independent stable clones. To further address the involvement of cPLA2γ in the H2O2-induced AA release, we tested the effect of MAFP, which inhibits both cPLA2α and Ca2+-independent PLA2 (iPLA2) and has also been reported to inhibit cPLA2γ (21Stewart A. Ghosh M. Spencer D.M. Leslie C.C. J. Biol. Chem. 2002; 277: 29526-29536Google Scholar). Pretreatment with 100 μm MAFP exerted a significant inhibitory effect on the H2O2-induced AA release from cPLA2γ-overexpressing cells. MAFP also inhibited H2O2-induced AA release from control cells, probably through inhibition of intrinsic cPLA2α or iPLA2. We investigated H2O2-initiated signaling pathways, because we could not detect the activation of cPLA2γ by H2O2 in vitro (data not shown). ROS cause protein phosphorylation through the activation of protein tyrosine kinase (PTK) and protein kinase C (PKC) in different cell types (25Jin N. Hatton N.D. Harrington M.A. Xia X. Larsen S.H. Rhoades R.A. Free Radic. Biol. Med. 2000; 29: 736-746Google Scholar, 26Konishi H. Tanaka M. Takemura Y. Matsuzaki H. Ono Y. Kikkawa U. Nishizuka Y. Proc. Natl. Acad. Sci. U. S. A. 1997; 94: 11233-11237Google Scholar, 27Guyton K.Z. Liu Y. Gorospe M. Xu Q. Holbrook N.J. J. Biol. Chem. 1996; 271: 4138-4142Google Scholar), including vascular smooth muscle and cultured endothelial cells. We then tested various inhibitors that regulate the tyrosine or serine/threonine phosphorylation. As shown in Fig.4, calphostin C, an inhibitor of PKC, did not have any inhibitory effects on H2O2-induced AA release but rather a slightly up-regulatory effect. On the other hand, herbimycin A, a PTK inhibitor, inhibited H2O2-induced AA release, whereas orthovanadate (Na3VO4), a protein tyrosine phosphatase inhibitor, enhanced H2O2-induced AA release. We could not detect any effect induced by addition of Na3VO4 by itself (data not shown). Thus, cPLA2γ enhanced H2O2-induced AA release through a tyrosine phosphorylation pathway. We could not detect the direct tyrosine phosphorylation of cPLA2γ by immunoprecipitation and Western blotting (data not shown). The mechanisms involved in this activation process remain to be elucidated. The major findings of the present study are as follows. 1) cPLA2γ is localized in the ER and Golgi, where membrane phospholipids are abundantly produced. 2) expression of cPLA2γ changes the fatty acid composition in PE but not in PC, as determined by ESI-MS. 3) cPLA2γ is involved in H2O2-induced AA release. ESI-MS analysis showing that the expression of cPLA2γ changed the fatty acid composition in PE (Fig. 2) suggests that cPLA2γ has a substrate preference for PE in intact cells. In an in vitro assay using cell lysates derived from HEK293 cells overexpressing cPLA2γ, the enzyme preferred PE over PC or phosphatidylinositol. 2K. Asai, T. Hirabayashi, N. Uozumi, and T. Shimizu, unpublished data. The precise mechanism by which a relative abundance of PUFA at thesn-2 position of PE was produced in cPLA2γ-overexpressing cells (Fig. 2 A) remains uncertain. Saturated or monounsaturated fatty acids at thesn-2 position of PE may be preferentially hydrolyzed by the PLA2 activity of cPLA2γ, and the derived lysoPE may be reacylated with PUFA by other enzymes with acyltransferase activities. Alternatively, cPLA2γ may have transacylase or acyltransferase activity in itself, as in the case of other PLA2s (28Reynolds L.J. Hughes L.L. Louis A.I. Kramer R.M. Dennis E.A. Biochim. Biophys. Acta. 1993; 1167: 272-280Google Scholar, 29Lio Y.C. Dennis E.A. Biochim. Biophys. Acta. 1998; 1392: 320-332Google Scholar). Lysophospholipase activity of cPLA2γ (21Stewart A. Ghosh M. Spencer D.M. Leslie C.C. J. Biol. Chem. 2002; 277: 29526-29536Google Scholar) may also contribute to the remodeling of the phospholipids. ER is a major organelle wherein the biosynthesis of various kinds of phospholipids, including PE, occurs (30Lykidis A. Baburina I. Jackowski S. J. Biol. Chem. 1999; 274: 26992-27001Google Scholar, 31Spencer A.G. Woods J.W. Arakawa T. Singer I.I Smith W.L. J. Biol. Chem. 1998; 273: 9886-9893Google Scholar, 32Rusinol A.E. Cui Z. Chen M.H. Vance J.E. J. Biol. Chem. 1994; 269: 27494-27502Google Scholar, 33Henneberry A.L. Wright M.M. McMaster C.R. Mol. Biol. Cell. 2002; 13: 3148-3161Google Scholar). Due to the fact that PE has a unique property in that it possesses high proportion of AA or docosahexaenoic acid (22:6) at the sn-2 position, the fatty acid composition in PE may be rearranged to the proper proportion in the ER by cPLA2γ. The membranes of the ER and Golgi, especially those of the smooth ER, constantly participate in protein and lipid mass transport by local vesicle formation and fusion. cPLA2γ may be involved in the local vesicle formation and fusion as suggested for other PLA2s (34de Figueiredo P. Drecktrah D. Katzenellenbogen J.A. Strang M. Brown W.J. Proc. Natl. Acad. Sci. U. S. A. 1998; 95: 8642-8647Google Scholar, 35Choukroun G.J. Marshansky V. Gustafson C.E. McKee M. Hajjar R.J. Rosenzweig A. Brown D. Bonventre J.V. J. Clin. Invest. 2000; 106: 983-993Google Scholar). As we only determined the subcellular localization of cPLA2γ by the overexpression system, we have to await definite localization in native cells until a good quality antibody is available. H2O2-induced AA release by cPLA2γ suggests that ROS regulate the activity of cPLA2γ at the cellular level. To determine whether the activity of cPLA2γ can be regulated at the cellular level, we searched for stimuli that enhanced the AA release and found H2O2 to be a candidate. H2O2 is one of the highly important ROS because of its ability to penetrate cellular membranes; as a precursor of the hydroxy radical, a powerful free radical, it can cause severe oxidative damage to membrane phospholipids, especially by oxidizing polyunsaturated fatty acids at the sn-2 position. Any oxidative modification of membrane phospholipids is a deleterious process, altering membrane fluidity, protein structure and cell signaling (36Imai K. Aimoto T. Shima T. Nakashima T. Sato M. Kimura R. Biol. Pharm. Bull. 2000; 23: 415-419Google Scholar, 37Rapoport S.M. Schewe T. Biochim. Biophys. Acta. 1986; 864: 471-495Google Scholar, 38Uchida K. Shiraishi M. Naito Y. Torii Y. Nakamura Y. Osawa T. J. Biol. Chem. 1999; 274: 2234-2242Google Scholar). The best way for repairing the phospholipids is the selective cleavage of the peroxidized fatty acid residues and their subsequent replacement by native fatty acids. Although several studies have shown that ROS cause AA release from cells through PLA2, such as iPLA2 or cPLA2α (39Birbes H. Gothie E. Pageaux J.F. Lagarde M. Laugier C. Biochem. Biophys. Res. Commun. 2000; 276: 613-618Google Scholar, 40Martinez J. Moreno J.J. Arch Biochem. Biophys. 2001; 392: 257-262Google Scholar, 41Buschbeck M. Ghomashchi F. Gelb M.H. Watson S.P. Borsch-Haubold A.G. Biochem. J. 1999; 344: 359-366Google Scholar, 42Mori A. Yasuda Y. Murayama T. Nomura Y. Eur. J. Pharmacol. 2001; 417: 19-25Google Scholar, 43McHowat J. Swift L.M. Arutunyan A. Sarvazyan N. Cancer Res. 2001; 61: 4024-4029Google Scholar, 44Cummings B.S. McHowat J. Schnellmann R.G. Am. J. Physiol. Renal Physiol. 2002; 283: F492-F498Google Scholar, 45Rao G.N. Runge M.S. Alexander R.W. Biochim Biophys Acta. 1995; 1265: 67-72Google Scholar, 46Hayama M. Inoue R. Akiba S. Sato T. Am. J. Physiol. Renal Physiol. 2002; 282: F485-F491Google Scholar, 47Nigam S. Schewe T. Biochim. Biophys. Acta. 2000; 1488: 167-181Google Scholar, 48Chaitidis P. Schewe T. Sutherland M. Kuhn H. Nigam S. FEBS Lett. 1998; 434: 437-441Google Scholar, 49Rashba-Step J. Tatoyan A. Duncan R. Ann D. Pushpa-Rehka T.R. Sevanian A. Arch. Biochem. Biophys. 1997; 343: 44-54Google Scholar), capabilities in these enzymes for selective cleavage of the peroxidized fatty acid residues have not been addressed to date. So far, the platelet-activating factor acetylhydrolases of type II, which cleave preferentially peroxidized or lipoxygenated phospholipids, are proposed to be competent for the repair (47Nigam S. Schewe T. Biochim. Biophys. Acta. 2000; 1488: 167-181Google Scholar, 50Stafforini D.M. Rollins E.N. Prescott S.M. McIntyre T.M. J. Biol. Chem. 1993; 268: 3857-3865Google Scholar). Taken together, cPLA2γ may release AA in response to oxidative stress and increase the AA level to repair the oxidized fatty acids in phospholipids. Furthermore, considering the major tissue distribution of cPLA2γ in skeletal muscle and heart, it may contribute to the repair of the oxidized phospholipids in these distinct tissues in which oxidative stress on the unique phospholipid compositions were noted (51Waku K. Uda Y. Nakazawa Y. J. Biochem. (Tokyo). 1971; 69: 483-491Google Scholar, 52Okano G. Matsuzaka H. Shimojo T. Biochim. Biophys. Acta. 1980; 619: 167-175Google Scholar). Regarding the mechanism by which H2O2 activates cPLA2γ, at least three possibilities exist: 1) modification of the enzyme itself or associated molecules; 2) changes in the phospholipids environment; and 3) existence of a recognition mechanism for the oxidized product or oxidative state. Several signal transduction pathways in cultured mammalian cells have been activated by the application of H2O2, including PTKs, mitogen-activated protein (MAP) kinases, PKC isoforms, and the epidermal growth factor receptor (25Jin N. Hatton N.D. Harrington M.A. Xia X. Larsen S.H. Rhoades R.A. Free Radic. Biol. Med. 2000; 29: 736-746Google Scholar, 26Konishi H. Tanaka M. Takemura Y. Matsuzaki H. Ono Y. Kikkawa U. Nishizuka Y. Proc. Natl. Acad. Sci. U. S. A. 1997; 94: 11233-11237Google Scholar, 27Guyton K.Z. Liu Y. Gorospe M. Xu Q. Holbrook N.J. J. Biol. Chem. 1996; 271: 4138-4142Google Scholar). For example, cPLA2α is reported to be activated by H2O2 through the activation of extracellular signal-regulated kinase and p38 MAP kinase in mesangial cells (46Hayama M. Inoue R. Akiba S. Sato T. Am. J. Physiol. Renal Physiol. 2002; 282: F485-F491Google Scholar). Unlike cPLA2α, where MAP kinase phosphorylation sites exist (53Borsch-Haubold A.G. Bartoli F. Asselin J. Dudler T. Kramer R.M. Apitz-Castro R. Watson S.P. Gelb M.H. J. Biol. Chem. 1998; 273: 4449-4458Google Scholar, 54Lin L.L. Wartmann M. Lin A.Y. Knopf J.L. Seth A. Davis R.J. Cell. 1993; 72: 269-278Google Scholar, 55Nemenoff R.A. Winitz S. Qian N.X. Van Putten V. Johnson G.L. Heasley L.E. J. Biol. Chem. 1993; 268: 1960-1964Google Scholar), cPLA2γ is postulated to possess consensus sequence for PKC and PTK phosphorylation. In this work we showed that cPLA2γ is activated by H2O2through the tyrosine phosphorylation pathway, as demonstrated by the inhibitor assays. In our preliminary experiments we could not detect direct tyrosine phosphorylation of cPLA2γ by immunoprecipitation and Western blotting (data not shown). These results suggest that there may be associated molecules that regulate the cPLA2γ activity in response to the oxidative stress. Other possibilities remain to be explored. In conclusion, we have provided several new insights into the properties of human cPLA2γ, including PE preferencein vivo, localization in the ER and Golgi, and involvement of H2O2-induced AA release. Herein we have elucidated the possible roles of cPLA2γ in mammalian cells, i.e. membrane remodeling and oxidative stress-induced AA release. The physiological and pathological functions of the enzyme would be clarified by analyzing cPLA2γ knockout mice. We thank Drs. M. Murakami and I. Kudo at Showa University for invaluable suggestions and Drs. S. Hoshiko, F. Osawa, and Y. Akamatsu at the Pharmaceutical Research Center, Meiji Seika Kaisha, Ltd. for the encouragement they gave us." @default.
• W2012194536 created "2016-06-24" @default.
• W2012194536 creator A5000716531 @default.
• W2012194536 creator A5008934626 @default.
• W2012194536 creator A5029011785 @default.
• W2012194536 creator A5029449740 @default.
• W2012194536 creator A5048776946 @default.
• W2012194536 creator A5068102473 @default.
• W2012194536 date "2003-03-01" @default.
• W2012194536 modified "2023-10-14" @default.
• W2012194536 title "Human Group IVC Phospholipase A2(cPLA2γ)" @default.
• W2012194536 cites W1487384455 @default.
• W2012194536 cites W1510592624 @default.
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