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Arteriosclerosis, Thrombosis, and Vascular Biology. 1999;19:1291-1298

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(Arteriosclerosis, Thrombosis, and Vascular Biology. 1999;19:1291-1298.)
© 1999 American Heart Association, Inc.

Atherosclerosis and Lipoproteins

Role of Group II Secretory Phospholipase A2 in Atherosclerosis

2. Potential Involvement of Biologically Active Oxidized Phospholipids

Norbert Leitinger; Andrew D. Watson; Susan Y. Hama; Boris Ivandic; Jian-Hua Qiao; Joakim Huber; Kym F. Faull; David S. Grass; Mohamad Navab; Alan M. Fogelman; Frederick C. de Beer; Aldons J. Lusis; Judith A. Berliner

From the Departments of Medicine (N.L., A.D.W., S.Y.H., B.I., J.-H.Q., J.H., M.N., A.M.F., A.J.L., J.A.B.), Pathology (N.L., J.A.B.), Psychiatry and Biobehavioral Sciences and the Neuropsychiatric Institute (K.M.F.), University of California, Los Angeles; Chrysalis DNX Transgenic Sciences (D.S.G.), Princeton, NJ; and the Department of Medicine (F.C.d.B.), University of Kentucky, Lexington. Current address for Norbert Leitinger, Department of Vascular Biology and Thrombosis Research, Schwarzspanierstrasse 17, A1090 Vienna, University of Vienna, Austria.

Correspondence to Judith A Berliner, 13-239 CHS, Departments of Pathology and Medicine, Los Angeles, CA 90095-1732. E-mail jberline{at}pathology.medsch.ucla.edu


*    Abstract
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Abstract—Secretory nonpancreatic phospholipase A2 (group II sPLA2) is induced in inflammation and present in atherosclerotic lesions. In an accompanying publication we demonstrate that transgenic mice expressing group II sPLA2 developed severe atherosclerosis. The current study was undertaken to determine whether 1 mechanism by which group II sPLA2 might contribute to the progression of inflammation and atherosclerosis is by increasing the formation of biologically active oxidized phospholipids. In vivo measurements of bioactive lipids were performed, and in vitro studies tested the hypothesis that sPLA2 can increase the accumulation of bioactive phospholipids. We have shown previously that 3 oxidized phospholipids derived from the oxidation of 1-palmitoyl-2-arachidonoyl-sn-glycero-3-phosphorylcholine (PAPC) stimulated endothelial cells to bind monocytes, a process that is known to be an important step in atherogenesis. We now show that these 3 biologically active phospholipids are significantly increased in livers of sPLA2 transgenic mice fed a high-fat diet as compared with nontransgenic littermates. We present in vitro evidence for several mechanisms by which these phospholipids may be increased in sPLA2 transgenics. These studies demonstrated that polyunsaturated free fatty acids, which are liberated by sPLA2, increased the formation of bioactive phospholipids in LDL, resulting in increased ability to stimulate monocyte-endothelial interactions. Moreover, sPLA2-treated LDL was oxidized by cocultures of human aortic endothelial cells and smooth muscle cells more efficiently than untreated LDL. Analysis by electrospray ionization–mass spectrometry revealed that the bioactive phospholipids, compared with unoxidized PAPC, were less susceptible to hydrolysis by human recombinant group II sPLA2. In addition, HDL from the transgenic mice and human HDL treated with recombinant sPLA2 in vitro failed, in the coculture system, to protect against the formation of biologically active phospholipids in LDL. This lack of protection may in part relate to the decreased levels of paraoxonase seen in the HDL isolated from the transgenic animals. Taken together, these studies show that levels of biologically active oxidized phospholipids are increased in sPLA2 transgenic mice; they also suggest that this increase may be mediated by effects of sPLA2 on both LDL and HDL.


Key Words: mice, transgenic • lipid peroxidation • LDL oxidation • inflammation • spectrum analysis, mass • phospholipids


*    Introduction
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 *Introduction
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It is shown in an accompanying paper that fatty streak formation is increased in sPLA2 transgenic mice on a high-fat diet.1 There is considerable evidence that oxidative modification of LDL is directly related to the development of the atherosclerotic lesion.2 3 4 5 6 7 Treatment of endothelial cells with minimally oxidized LDL (LDL that has undergone antioxidant depletion and oxidation of arachidonic acid-containing phospholipids, with little or no protein modification) increased monocyte binding and transmigration in cocultures of endothelial cells and smooth muscle cells.4 8 The major lipids responsible for the biological activity of minimally oxidized LDL (MM-LDL) are derived from the oxidation of arachidonate-containing phospholipids9 such as 1-palmitoyl-2-arachidonoyl-sn-glycero-3-phosphorylcholine (PAPC). Three active lipids accounted for the major bioactivity in MM-LDL and oxidized PAPC (OxPAPC): 1-palmitoyl-2-(5)oxovaleroyl-sn-glycero-3-phosphorylcholine (POVPC, m/z 594.3 (M+H+)), 1-palmitoyl-2-glutaroyl-sn-glycero-3-phosphorylcholine (PGPC, m/z 610.2 (M+H+)), and a molecule with m/z 828.6 (M+H+).10 We demonstrated that these bioactive phospholipids were increased in fatty streak lesions from cholesterol-fed rabbits and they were significantly less abundant when the atherosclerotic diet was supplemented with antioxidants such as vitamin E and probucol.10 Natural antibodies against these phospholipids were found in the serum of apoE null mice, which develop advanced atherosclerotic lesions owing to massive hypercholesterolemia.10 11

Both the liver and the arterial wall are sites of lipid accumulation when mice are fed an atherogenic diet. Moreover, in recombinant inbred strains derived from C57BL/6J and C3H/HeJ mice, hepatic inflammatory gene induction and aortic fatty streak development cosegregated, indicating similar pathophysiology in these tissues.12 In these same recombinant inbred strains, levels of POVPC, PGPC, and the molecule with m/z 828.6 in livers also cosegregated with lesion development.13 In the present study we determined whether levels of biologically active oxidized phospholipids were increased in sPLA2 transgenics and performed in vitro tests of the hypothesis that sPLA2 accelerates their accumulation.


*    Methods
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Materials
L-{alpha}-1-Palmitoyl-2-arachidonoyl-sn-glycero-3-phosphorylcholine was obtained from Avanti Polar Lipids Inc. or Sigma Chemical Co. Soybean lipoxygenases (SLO), sPLA2 (Naja naja), butylated hydroxytoluene (BHT), dimyristoylphosphatidylcholine (DMPC), palmitic acid, stearic acid, oleic acid, linoleic acid, arachidonic acid, and endotoxin-free gelatin, tissue-culture grade, were obtained from Sigma. Aminopropyl solid phase extraction columns were obtained from J.T. Baker. Chloroform, methanol, isopropanol, and ethyl ether (all HPLC grade or better) were obtained from Fisher Scientific. Tissue culture media, serum, and supplements were obtained from sources previously reported.8 Transwells and chamber slides were obtained from Costar. The fluorescent probe DiI (1,1'-dioctadecyl-3,3,3'-tetramethyl-indo-carbocyanine perchlorate) was purchased from Molecular Probes. Human recombinant sPLA2 was a generous gifts from Dr J. Browning, Biogen. The method of preparation of sPLA2 has been previously described.14 Briefly, recombinant sPLA2 was purified from medium of Chinese hamster ovary (CHO) cells transfected with an expression vector encoding human sPLA2.

Animals
sPLA2 transgenic mice were obtained from Chrysalis DNX Transgenic Sciences, Princeton, NJ, and had been produced as described previously.15 The atherogenic diet contained 1.25% cholesterol, 15.75% fat, and 0.5% sodium cholate (TD 90221, Harlan-Teklad). After 12 weeks on the high-fat diet, the mice were sacrificed by cervical dislocation after the administration of isoflurane. All animal procedures were conducted according to the regulations of the University of California Animal Research Committee.

Lipoprotein Isolation and Modification
Human LDL (d=1.019 to 1.069 g/mL) and HDL (d=1.069 to 1.210 g/mL) were isolated from the sera of healthy blood donors by density gradient ultracentrifugation as described16 and used within 1 to 2 weeks of isolation. Concentration of LDL is expressed in terms of protein content as determined according to the method of Lowry et al.17 The concentration of endotoxin in solutions tested in biological assays was less than 50 pg/mL (determined by a chromogenic assay), which is 40-fold less than that required to induce monocyte-endothelial interactions. LDL was modified using purified SLO and PLA2, according to Sparrow et al.18 To remove enzymes from the reaction by centrifugation, SLO and PLA2 were separately bound to CNBr-activated Sepharose beads as described.18 LDL (1 mg/mL) was modified by incubation with SLO (5000 U/mL) alone, SLO+PLA2 (20 U/mL), or SLO+fatty acids (187 mmol/L) at 37°C for 24 hours or at 4°C for 48 hours.19 The enzymes were removed by centrifugation, BHT (100 µmol/L) and EDTA (0.3 mmol/L) were added, and the modified LDL was stored at 4°C and used within 2 weeks.

Mouse HDL were isolated from pooled plasma by fast protein liquid chromatography,20 in the absence of EDTA to avoid inactivation of paraoxonase. HDL were incubated at concentrations of 2 mg protein/mL with recombinant human sPLA2 (0.5 µg/mL) for 16 hours, subsequently isolated again, and then added to the cocultures at a final concentration of 250 to 350 µg/mL.

Cell Culture
Human aortic endothelial cells (HAEC) and rabbit aortic endothelial cells (RAEC) were cultured as described.8 Human aortic smooth muscle cells (HASMC) were isolated as previously described.21 Cocultures of HAEC and HASMC were grown, and monocyte binding and transmigration experiments were performed in transwells or chamber slides as previously described.21 In all experiments, HAEC and HASMC from the same donor were used at passages 4 to 6. Blood monocytes were obtained from a large pool of healthy donors by modification of the Recalde procedure as described previously.22

Monocyte Adhesion Assay
Binding of human monocytes to endothelial cells was performed essentially as described previously.8 HAEC or RAEC were treated with MM-LDL (125 to 200 µg/mL) for 4 hours at 37°C. Treatment media were removed, the cells were rinsed twice with medium, and a suspension of human monocytes (2 to 3x105/well) was added for 12 minutes. Unbound monocytes were removed, and the number of bound monocytes was determined microscopically.

Monocyte Transmigration Assay
These assays were performed essentially as described previously.21 Cocultures were treated with native LDL (250 to 350 mg/mL) in the absence or presence of various test compounds for 24 hours. The culture supernatants were subsequently transferred to untreated cocultures and were incubated for an additional 24 hours. Monocytes were labeled with the fluorescent probe DiI at 4°C for 10 minutes, washed, and resuspended in medium-199 at the desired cell density. Labeled monocytes were added to the treated cocultures at 2x105 cells/cm2 and were incubated for 60 minutes at 37°C. The medium containing nonadherent cells was removed, and the cell layers were washed to remove loosely adherent monocytes. The cocultures were fixed on polycarbonate membranes with 10% neutral buffered formalin at room temperature for 24 hours and were mounted on glass slides. The number of monocytes in the subendothelial space (beneath the endothelial cells) in a minimum of 9 fields was determined under 500x magnification and fluorescence microscopy.

Measurement of Lipid Hydroperoxides
Lipid hydroperoxide formation was measured using the method of Auerbach et al.23

Lipid Extraction
Lipids were extracted from modified LDL and from mouse livers using a modification of the method of Bligh and Dyer.24 Liver tissue samples were homogenized in chloroform/methanol (2:1) containing 0.01% BHT for 5 minutes at 0°C. After addition of water, liver suspensions were vortexed for 2 minutes and centrifuged at 1200g for 15 minutes at 4°C (Beckman J-6B; rotor, Beckman JS-4.2). In separate studies chloroform/methanol/BHT was added to modified LDL in PBS, and these suspensions were vortexed for 2 minutes. The chloroform phase was carefully removed and 5 vol of chloroform was added to the residual aqueous phase. The mixture was vortexed and centrifuged as described above, and the chloroform phase was pooled with the previous extract. Phospholipid recovery typically ranged between 96% and 98% as determined by spiking samples with known quantities of 1-palmitoyl-2-[14C]arachidonoyl-sn-glycero-3-phosphorylcholine.

Solid Phase Extraction
Phospholipids, free fatty acids, and neutral lipids were separated by the method of Kaluzny et al.25 using aminopropyl solid phase extraction chromatography. Total lipid extracts were dried under nitrogen, resuspended in 200 µL of chloroform, and applied to the aminopropyl columns, which had been washed with 3 mL of methanol and preconditioned with 3 mL of hexane. Neutral lipids were eluted with 3 mL of chloroform/2-propanol (2:1), free fatty acids with 3 mL of 3% acetic acid in ethyl ether, and phospholipids with 3 mL of methanol. When phospholipids were used in tissue culture experiments, acetic acid retained in the columns was removed with pure ethyl ether before phospholipid elution. Lipid fractions were dried under nitrogen and either used immediately or resuspended in chloroform containing 0.01% BHT, covered with argon, and stored at -44°C.

Substrate Specificity Assay
OxPAPC was produced by exposing dry PAPC to air for 3 days. Oxidation products, as well as residual unoxidized PAPC, were present in these preparations. Liposomes were produced by sonicating OxPAPC for 10 minutes in Tris buffer, pH 7.4. Fifty microliters containing 500 µg of OxPAPC liposomes was incubated with either human recombinant sPLA2 (1 µg), snake venom sPLA2 (Naja naja; 5 U), or buffer alone for 90 minutes at room temperature in Tris buffer containing 2 mmol/L Ca2+. The reaction was stopped and lipids were extracted by adding 200 µL chloroform/methanol (2:1)+0.01% BHT. After vortexing and centrifugation for 10 minutes, 100 µL of the chloroform phase was transferred into a glass tube, and 500 ng of internal standard (DMPC) was added to each tube.26 Samples were dried under a stream of nitrogen and redissolved in acetonitrile/water/formic acid (50:50:0.1; vol/vol/vol) and analyzed by electrospray ionization–mass spectrometry (ESI-MS).

Electrospray Ionization–Mass Spectrometry
An API III triple-quadrupole biomolecular mass analyzer (Perkin-Elmer Sciex Instruments) was used for mass analysis of phospholipids. Flow-injection analysis was performed as described previously.10 For quantitative measurements, DMPC was used as an internal standard.26 Mass spectrometry of individual phospholipid molecular species represents the most sensitive, discriminating, and direct method to assess alterations in phospholipid molecular species in biological tissues.26 In contrast to previously used techniques like fast atom bombardment mass spectrometry, which results in extensive fragmentation of individual phospholipid ions, electrospray ionization results in the efficient production of molecular ions of phospholipids with negligible fragmentation.27 Therefore, ESI-MS offers a sensitive and reliable tool for quantitative phospholipid analysis in biological tissues. For quantitative measurements we were assuming that possible differential effects of phospholipid oxidation on extraction recovery, ionization efficiency, and molecular ion yield were negligibly small in comparison with the quantitative differences reported. In an extensive examination of phospholipid quantitation by ESI-MS, Han et al26 reported that the effects of unsaturation and differential surface characteristics were minimal. Because any 13C effects on quantitation were constant within the comparisons made, no corrections of the molecular ion signal intensities have been performed. Within the range of concentrations measured, trial experiments indicated that the response was linear, such that increases in phospholipid content were reflected in proportional increases in phospholipid to internal standard ratio.

Statistical Analysis
Statistical analysis was performed using 1-way ANOVA; probability values <0.05 were considered statistically significant.


*    Results
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Evaluation of Phospholipids by ESI-MS
Previous studies have shown that group II sPLA2 was expressed to a significantly greater extent in livers15 and aortas1 of sPLA2 transgenic mice when compared with nontransgenic littermates. Liver was used as a surrogate for aorta as validated in previous studies in Bl-6 mice.12 Phospholipids were separated from a total lipid extract obtained from livers of female sPLA2 transgenic mice and nontransgenic littermates maintained for 12 weeks on an atherogenic diet. Analysis of phospholipids was performed by ESI-MS, and quantification of the different phospholipid species was carried out by addition of DMPC as an internal standard, as described in the Methods. The levels of native phospholipids, 1-palmitoyl-2-arachidonoyl-sn-glycero-3-phosphorylcholine (PLPC, m/z 758) and PAPC, m/z 782, in livers of transgenic female mice were not significantly different from the levels in livers of nontransgenic (Figure 1ADown). In contrast, the levels of POVPC and PGPC were significantly increased in transgenic animals versus nontransgenics (Figure 1BDown; P<0.003 for POVPC, P<0.014 for PGPC). The bioactive molecule with m/z 828.6 was not increased in the transgenic compared with nontransgenic animals (Figure 1BDown). Interestingly, the levels of lysophosphatidylcholine (LPC) (sn-1=palmitic acid (16:0), m/z 496, and sn-1=stearic acid (18:0), m/z 524 were not significantly different between transgenic and nontransgenic animals (Figure 1ADown).



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  		Figure 1. Levels of unoxidized phospholipids and LPC (A) and bioactive derivatives of oxidized PAPC (B) in livers of sPLA2 transgenic (TG) and nontransgenic (nonTG) mice. Levels of phosphatidylcholines in livers were determined using positive-mode ESI-MS. Lipids were extracted with chloroform/methanol and phospholipids separated using solid phase extraction columns after addition of DMPC as an internal standard. The abundance of ions with m/z 782 (PAPC), m/z 758 (PLPC), m/z 496 (LPC 16:0), m/z 524 (LPC 18:0), m/z 594.3 (POVPC), m/z 610.2 (PGPC), and m/z 828.6 was determined using software supplied by PE Sciex and plotted in terms of ng DMPC equivalents/mg liver tissue. Data are presented as mean+SD (n=8 animals in each group). Levels of POVPC and PGPC were significantly increased in the livers of TG compared with nonTG animals (B) (P<0.003 for POVPC and P<0.014 for PGPC). However, there was no significant increase in native phospholipids PAPC or PLPC or of LPC in the livers of the transgenics (A).

Effect of sPLA2 Products on LDL Phospholipid Oxidation
Our group and others have shown that treatment of LDL with sPLA2 from Naja naja combined with SLO leads to the formation of conjugated dienes and bioactive phospholipids.18 19 On the basis of these findings, we hypothesized that treatment of LDL with sPLA2 led to a limited liberation of fatty acids that, when oxidized, could cause oxidation of arachidonate-containing phospholipids, leading to the formation of bioactive phospholipids. To test this hypothesis, LDL was incubated without additions, or in the presence of SLO and linoleic acid. The formation of conjugated dienes was increased 2- to 3-fold in the phospholipid fraction of the treated LDL compared with the untreated LDL (Figure 2ADown). An approximately 10-fold increase in the level of absorbance at 270 nm, an indication for the levels of bioactive phospholipids,9 was also observed (Figure 2BDown). The treated LDL preparation was active in stimulating endothelial cells to bind monocytes, further confirming the presence of biologically active phospholipids (Figure 2CDown). Arachidonic acid had similar effects (data not shown). To further test the hypothesis that oxidized fatty acids play a role in the formation of bioactive phospholipids, LDL was treated with oxidized linoleic acid in the absence of other additives. This treatment also stimulated the formation of bioactive phospholipids (monocyte binding increased by 2.4±0.3-fold). Oxidized fatty acids alone in the absence of LDL had no effect on monocyte binding (data not shown). Several lines of evidence suggest that lipoxygenase may play a significant role in the oxidation of LDL by cells.28 Free polyunsaturated fatty acids (PUFAs) are better substrates than esterified PUFAs for some mammalian lipoxygenases. We tested the hypothesis that treatment of LDL with sPLA2 would increase the ability of cells to oxidize LDL and thereby form bioactive phospholipids. Pretreatment of LDL with different concentrations of human recombinant sPLA2 and subsequent incubation for 24 hours with cocultures of HAEC and HASMC resulted in increased formation of lipid hydroperoxides (Figure 3ADown) and increased bioactivity (Figure 3BDown).



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  		Figure 2. Biological activity, phospholipid-associated conjugated diene formation, and absorbance at 270 nm in untreated and SLO+linoleic acid-treated LDL. LDL were incubated with SLO and linoleic acid (18:2). Lipids were extracted from LDL and phospholipids were isolated by solid phase extraction chromatography and analyzed for conjugated dienes (A) and for presence of bioactive phospholipids (B) by measuring absorbance at 234 and 270 nm, respectively. C, RAEC were incubated with LDL oxidized with SLO and linoleic acid (18:2) or untreated LDL, and then tested for induction of human monocyte adhesion. These data are representative of 4 separate experiments in which each condition was tested in quadruplicate. Values are mean+SD (n=4). *Indicates P<0.001.



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  		Figure 3. Effect of sPLA2 treatment of LDL on oxidation by coculture cells and induction of monocyte transmigration. Human LDL was treated with different concentrations of human recombinant sPLA2 for 4 hours, then incubated with cocultures of endothelial cells and smooth muscle cells for 24 hours. The supernatant was then removed and tested for levels of lipid hydroperoxides (A) and tested on a second set of cocultures for ability to induce monocyte transmigration (B). Lipid hydroperoxide levels were significantly elevated with as little as 0.1 µg/mL of sPLA2. Lower levels of sPLA2-treated LDL versus untreated LDL were more effective in stimulating monocyte transmigration (P<0.05). These data are representative of 4 separate experiments in which each condition was tested in quadruplicate. Mean+SD (n=4).

Substrate Specificity of sPLA2
It was clear from the above studies that sPLA2 could both hydrolyze phospholipids and lead to the formation of bioactive phospholipids. We hypothesized that these 2 observations were compatible with the idea that other phospholipids in OxPAPC were better substrates for sPLA2 than the bioactive oxidized phospholipids. To assess the substrate specificity among the various species of oxidized phospholipids present in OxPAPC, a human recombinant sPLA2 and snake venom sPLA2 were used. Liposomes consisting of OxPAPC (which contained small amounts of native PAPC) were incubated with the enzymes and analyzed by ESI-MS. The bioactive molecules POVPC (m/z 594.3), PGPC (m/z 610.2), and m/z 828.6 were poorly hydrolyzed by the treatment with human recombinant sPLA2, whereas unoxidized PAPC (m/z 782.4) was effectively hydrolyzed (Figure 4Down and the TableDown). After treatment with recombinant sPLA2, the level of LPC was increased to a greater extent than could be accounted for by hydrolysis of native PAPC. This suggested that the excess LPC was because of hydrolysis of other oxidized phospholipids present in OxPAPC. Although the reduction of the >100 peaks (seen by MS) is not necessarily discernible in Figure 4Down, the collective production of LPC from these lipids is highly significant. The snake venom sPLA2 showed a preference for POVPC (m/z 594.3) and unoxidized PAPC (m/z 782.4) among the phospholipids present in OxPAPC. Representative mass spectra are shown in Figure 4Down and the results of these experiments are summarized in the TableDown.



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  		Figure 4. Substrate specificity of human recombinant sPLA2 on liposomes of oxidized PAPC. Representative mass spectra of OxPAPC liposomes under the following conditions: untreated (A), after treatment with human recombinant sPLA2 (B), and after treatment with snake venom sPLA2 (C). Inset B, Difference between human recombinant sPLA2-treated and untreated (A minus B). Inset C, Difference between snake venom sPLA2-treated and untreated (A minus C). For these studies 50 µL containing 500 µg of OxPAPC liposomes was incubated with human recombinant sPLA2 (1 µg), snake venom sPLA2 (Naja naja; 5 U), or buffer alone for 90 minutes at room temperature in Tris buffer containing 2 mmol/L Ca2+. Lipids were extracted and analyzed by ESI-MS as described in the Methods. Relative changes of different phospholipid species were evaluated by comparing relative ion abundance to the internal standard (DMPC, m/z 678.6). (In C, the level of LPC, m/z 496, was approximately 300 as measured on an expanded scale.) These experiments were performed 3 times, and a representative experiment is shown.


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  		Table 1. Substrate Specificity of Human Recombinant sPLA2 Among Oxidized Phosphatidylcholines

HDL From sPLA2 Transgenic Mice Fail to Protect Against LDL Oxidation
HDL has been shown in vitro to prevent LDL oxidation and accumulation of bioactive lipids.4 This protective HDL function may also play an important role in fatty streak formation. We examined the protective effect of HDL from sPLA2 transgenic mice and their nontransgenic littermates using a coculture system in which cell-induced LDL oxidation stimulates the secretion of monocyte chemotactic protein-1 (MCP-1): levels of MCP-1 are assessed as monocyte chemotaxis and transmigration in the coculture.29 As previously reported, the addition of HDL along with LDL to such cocultures inhibited the formation of biologically active oxidized LDL.29 The HDL isolated from the plasma of sPLA2 transgenic mice did not protect against LDL-induced monocyte transmigration, whereas HDL from nontransgenic littermates was protective (Figure 5ADown). In fact, the HDL from the transgenic mice significantly promoted LDL-oxidation and monocyte transmigration in the coculture. The HDL from the sPLA2 transgenics contained one-third the paraoxonase activity of HDL from nontransgenics. To further study the sPLA2-mediated effects on HDL, we performed additional in vitro experiments (Figure 5BDown). First, we preincubated human HDL with sPLA2 for 3 hours with gentle mixing; the HDL was then isolated again and added to the coculture together with human LDL. Monocyte transmigration in the coculture containing sPLA2-treated HDL was increased 2-fold as compared with the addition of LDL alone (no HDL) and {approx}3-fold compared with the addition of LDL and sham-treated HDL. Therefore, sPLA2-treated HDL not only failed to protect against LDL oxidation but also exhibited proinflammatory capacity.



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  		Figure 5. HDL isolated from sPLA2 transgenic mice (A) or HDL treated in vitro with sPLA2 (B) fail to protect against LDL oxidation in a coculture model of the artery wall. A, Human aortic smooth muscle and endothelial cell cocultures were incubated for 18 hours with medium (no additions), 350 µg/mL LDL (no HDL), or LDL plus HDL (nontransgenic HDL, transgenic HDL). B, sPLA2-treated HDL denotes HDL isolated again that had been first incubated with sPLA2 (0.5 µg/mL) for 3 hours; sham-treated HDL, HDL pretreated in a similar fashion omitting sPLA2. Supernatants were then transferred to new cocultures. Labeled human blood monocytes were added and allowed to adhere and migrate into the endothelial cell layer for 1 hour. After washing off nonadherent cells, cocultures were fixed and the recruited monocytes were counted using 625x magnification. The number of monocytes per high-power field are given as mean+SD.


*    Discussion
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Previous studies reported in the accompanying manuscript1 showed that atherosclerosis was increased in sPLA2 transgenic mice. In these studies and those of others, group II sPLA2 was shown to be present in atherosclerotic lesions.1 30 31 A major goal of these studies was to test the hypothesis that sPLA2 could increase the levels of biologically active oxidized phospholipids, previously shown to have atherogenic effects.10 The importance of bioactive lipids in the development of atherosclerosis is suggested by the findings that animals injected with MDA-LDL32 or oxidized LDL33 before cholesterol feeding developed antibodies to the lipids and were partially protected against lesion development during cholesterol feeding. A role for these bioactive phospholipids in other inflammatory processes is suggested by the presence of antibodies to the bioactive phospholipids in a number of diseases of chronic inflammation.34

Because of the limited amount of fatty streak lesion material available for the study we compared the levels of bioactive phospholipids in livers of transgenic and nontransgenic female mice. The results of previous studies, showing that pathological changes in livers of Bl-6 mice fed an atherogenic diet parallel those in the aorta,12 make the liver an appropriate target for study in these animals. We found that the levels of the native phospholipids PLPC and PAPC were not increased in the livers of the transgenic mice compared with the nontransgenics (Figure 1AUp). Somewhat surprisingly, levels of lysophosphatidylcholine were also not increased (Figure 1AUp), perhaps because of high levels of lysophospholipase in the liver.35 However, levels of the biologically active oxidized phospholipids POVPC and PGPC were highly significantly elevated (Figure 1BUp) in the transgenics compared with the nontransgenics (Figure 1BUp).

Our in vitro studies suggest that 1 mechanism by which group II sPLA2 accelerates atherosclerosis is by increasing the levels of biologically active oxidized phospholipids. We present studies suggesting that sPLA2 may lead to more generation and less destruction of active lipids. The hypothetical scheme for the increased formation of bioactive phospholipids in sPLA2 transgenic animals is shown in Figure 6Down.



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  		Figure 6. Scheme demonstrating the hypothetical involvement of sPLA2 in the development of biologically active oxidized phospholipids. The enzymatic reaction of sPLA2 liberates PUFAs from the sn-2 position of phospholipids, mainly phosphatidylethanolamines, because of the substrate specificity of the enzyme. Formation of reactive oxygen species (ROS) in liver cells and arterial wall cells may be increased because of acute-phase conditions caused by the high-fat diet. Mechanisms by which ROS are released may involve cyclooxygenase, lipoxygenase, inducible nitric oxide synthase, cytochrome P450, mitochondrial respiration, or free iron. These oxygen radicals may attack free PUFAs and produce lipid hydroperoxides, which contribute to the propagation of oxidation of esterified PUFAs to form biologically active phospholipids. Biologically active oxidized phospholipids were suggested to be substrates for paraoxonase (PON), an enzyme associated with HDL. The enzymatic reaction of PON would destroy these bioactive phospholipids into molecules that do not evoke the characteristic inflammatory responses in endothelial cells induced by MM-LDL. Decreased serum PON levels, as measured in the sPLA2 transgenic animals, however, may prolong the half-life and cause accumulation of biologically active phospholipids in various tissues and plasma. Induction of monocyte adhesion to endothelial cells by biologically active oxidized phospholipids would then lead to the development of the fatty streak lesion.

We and others have previously shown that treatment of LDL with SLO and PLA2 increases oxidation, resulting in the formation of bioactive LDL.18 19 Treatment of LDL with sPLA2 led to a limited release of PUFAs.36 We now show that treatment of LDL with free PUFAs (which are products of PLA2 activity) in the presence of SLO or with oxidized PUFA causes oxidation of PUFA-containing phospholipids and thereby formation of bioactive phospholipids (Figure 2Up). Others have shown that free PUFAs are much better substrates for mammalian lipoxygenases than are phospholipid-bound PUFAs.37 Furthermore, PUFA radicals can act essentially as catalysts to propagate lipid oxidation. Evidence that sPLA2 enhances the formation of biologically active phospholipids in cellular systems is provided by our studies in cell culture. Treatment of LDL with sPLA2 led to formation of bioactive phospholipids and hydroperoxides (Figure 3Up). Free fatty acids have been shown to diffuse spontaneously across phospholipid bilayers, described as a flip-flop mechanism.38 Recently, active transport in addition to passive diffusion of free fatty acids across cell membranes has been postulated. It has been shown that carrier-mediated uptake of fatty acids in hepatocytes follows an inwardly directed transmembrane proton gradient and is stimulated by the presence of H+ at the outer surface of the plasma membrane.39 Lower pH levels, which have been measured at sites of inflammation, would therefore increase the uptake of free fatty acids into cells. Because the uptake of free fatty acids into mitochondria for subsequent ß-oxidation is highly regulated, accumulating free fatty acids would be a likely substrate for oxidizing enzymes, resulting in increased levels of lipid hydroperoxides. These hydroperoxides could then leave the cells seeding LDL, resulting in the formation of biologically active oxidized phospholipids.

LDL may not be the only source of bioactive oxidized phospholipids in sPLA2 transgenic animals. In vivo, sPLA2 may also liberate PUFAs from other lipoproteins, dying erythrocytes, or membranes of other cells leading to formation of bioactive lipids. Others have shown that whereas normal cell membranes are not good substrates for sPLA2, membranes from damaged cells may become good substrates.40 Although our studies demonstrated that the bioactive phospholipids were not good substrates for human recombinant sPLA2, they also demonstrated that hydrolysis of other oxidized phospholipids in OxPAPC resulted in a significant increase in LPC after treatment (Figure 4Up). Others have demonstrated that, in oxidized phospholipid vesicles that contain phosphatidyl ethanolamine as well as phosphatidyl choline, there is significant hydrolysis of oxidized phospholipids by snake venom sPLA2, caused by increased Ca2+ binding affinity and activation of the enzyme.41 42 The present studies have focused on the role of sPLA2 in the formation of 3 important bioactive phospholipids. However, increased levels of sPLA2 may also increase the levels of other bioactive phospholipids such as lysophosphatidic acid. Fourcade et al43 showed that sPLA2 plays an important role in producing lysophosphatidic acid and in the release from the cell membrane in the form of microvesicles.

Our results suggest that sPLA2 may also increase the levels of biologically active oxidized phospholipids by altering HDL. We show that HDL isolated from the transgenic animals had lost its protective activity. (Figure 5AUp). In addition, we show that in vitro sPLA2-treated HDL also had lost its protective activity and even enhanced monocyte transmigration induced by cell-modified LDL (Figure 5BUp). HDL from sPLA2 transgenic mice had lower levels of paraoxonase,1 an enzyme that can degrade bioactive phospholipids.9 Therefore loss of paraoxonase may be 1 mechanism by which the HDL protective mechanism is compromised, resulting in increased levels of bioactive phospholipids. However, other changes in sPLA2-treated HDL, such as increased cholesterol delivery to cells, shown to occur with other lipases,44 may represent another important mechanism.

In summary, we have presented evidence that the increase in atherosclerosis in sPLA2 transgenic mice may relate to the ability of sPLA2 to release PUFAs, which catalyze the formation of bioactive phospholipids. These bioactive phospholipids are poorly degraded by sPLA2, and the loss of protective activity of HDL in transgenic animals, perhaps in part caused by decreased paraoxonase activity, may further enhance their accumulation.


*    Acknowledgments
 
This work was supported by National Institute of Health grants HL30568 (A.J.L., A.M.F., J.A.B.), AG10886 (F.C.d.B.), the Council for Tobacco Research (F.C.d.B.), and Veterans Administration Research Funds (F.C.d.B.).

Received January 12, 1998; accepted October 5, 1998.


*    References
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