Paraquat-induced phosphatidylserine oxidation and apoptosis are independent of activation of PLA2

James P. Fabisiak1, Valerian E. Kagan2, Yulia Y. Tyurina2, Vladimir A. Tyurin2, and John S. Lazo1

Departments of 1 Pharmacology and 2 Environmental and Occupational Health, Schools of Medicine and Public Health, University of Pittsburgh, Pittsburgh, Pennsylvania 15261

    ABSTRACT
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Abstract
Introduction
Methods
Results
Discussion
References

Paraquat is a pneumotoxin that causes lung injury by enhancing oxidative stress; however, the cellular responses to these redox events are undefined. We previously showed that paraquat produced selective peroxidation of phosphatidylserine that preceded apoptosis in 32D cells. We now report that the phospholipase A2 (PLA2) inhibitor quinacrine can attenuate phosphatidylserine oxidation and also block paraquat-induced apoptosis. Therefore, we investigated the potential for PLA2 to mediate apoptosis after paraquat. We found that, in contrast to quinacrine, the PLA2 inhibitors manoalide, aristolochic acid, and arachidonyl trifluoromethylketone failed to prevent paraquat-induced apoptosis. Moreover, no evidence of PLA2 activation was observed within 7 h after paraquat exposure. Finally, quinacrine failed to inhibit basal and 4-bromo-A-23187-induced release of [3H]arachidonic acid at concentrations that protected paraquat-induced apoptosis. We conclude that paraquat-induced phosphatidylserine oxidation and apoptosis occurred in the absence of PLA2 activation and that quinacrine protected phosphatidylserine and cell viability after paraquat in a PLA2-independent manner.

phospholipase A2; oxidative stress; lipid peroxidation; quinacrine; arachidonic acid

    INTRODUCTION
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Abstract
Introduction
Methods
Results
Discussion
References

PARAQUAT (PQ), a bipyridyl herbicide, serves as a prototypical environmental pneumotoxin, the toxic effects of which have been well described in both humans and animals (38). The mechanisms of cellular damage after PQ involve the P-450 reductase-dependent formation of reactive oxygen species and subsequent interactions with lipids, proteins, and nucleic acids (10). Because reactive oxygen species have been implicated in the final common pathway that triggers apoptosis (4), peroxidation of membrane lipids during oxidative stress represents one potential target for signaling apoptosis (14). It has previously been demonstrated that PQ induces apoptosis in 32D cells (8). In addition, it was found that the selective oxidation of phosphatidylserine (PS) precedes PQ-induced apoptosis. The externalization of PS to the outer leaflet of the plasma membrane, as measured by annexin V binding (9), now appears to be one hallmark of apoptosis, although the mechanism responsible for its translocation is not known. Inhibition of PS oxidation by overexpression of the antiapoptotic gene bcl-2 (8) suggests that Bcl-2 can protect cells from one of the consequences of oxidative stress, namely, lipid peroxidation, and further supports the hypothesis that PS oxidation is an important signal during apoptosis.

Membrane lipid peroxidation can affect membrane fluidity, alter the function of membrane-bound proteins, perturb ion fluxes, and generate toxic oxidized lipid products (18). Of particular interest is the potential for oxidative stress to activate phospholipase A2 (PLA2). Lipid peroxidation accelerates phospholipid hydrolysis in vascular endothelial cells (30), and oxidized phospholipids provide better substrates for PLA2 than native phospholipids (35). In addition, H2O2 appears to directly activate a signal-responsive PLA2 independent of lipid peroxidation and changes in intracellular Ca2+, perhaps through stimulation of enzyme phosphorylation (3). Activation of phospholipid hydrolysis can further amplify membrane damage (25) and generate potent eicosanoid and other signaling molecules (7).

Tumor necrosis factor (TNF) is a cytokine with potent cytolytic properties capable of inducing apoptosis in sensitive cells (21, 22). The ability of TNF to kill cells has been linked to its ability to activate intracellular oxidative stress (37) and PLA2 (13, 40, 42). The role of PLA2 in apoptosis after oxidants and other stimuli, however, remains unexplored. It is thus possible that PLA2 activation coincident with and resulting from lipid peroxidation can participate in the early signaling of apoptosis after PQ-induced oxidative stress.

We used our previously characterized model of PQ-induced apoptosis in 32D cells to examine the potential role of PLA2 (8). We tested the ability of multiple pharmacological PLA2 inhibitors to protect cells from PQ-induced apoptosis. We also assessed PLA2 activation after PQ exposure to determine any association between phospholipid hydrolysis and PS oxidation. Our results support the hypothesis that PLA2 is not required to mediate specific oxidation of PS and subsequent apoptosis after exposure to PQ.

    METHODS
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Abstract
Introduction
Methods
Results
Discussion
References

Materials. PQ dichloride, quinacrine (QN) hydrochloride, and manoalide were obtained from Sigma (St. Louis, MO). Arachidonyl trifluoromethylketone (AACOCF3) was from ICN (Costa Mesa, CA), and aristolochic acid was from Biomol (Plymouth Meeting, PA). Proteinase K, RNase A, and RNase T1 were from Boehringer Mannheim (Indianapolis, IN). [3H]arachidonic acid (AA) ([5,6,8,9,11,12,14,15-3H]AA; 180-240 Ci/mmol) and BetaMax scintillation fluid were obtained from DuPont NEN (Boston, MA). cis-Parinaric acid (cis-PNA), 4-bromo-A-23187 (4-Br-A-23187), and Hoechst 33342 were obtained from Molecular Probes (Eugene, OR). All tissue culture media and reagents were from GIBCO BRL (Gaithersburg, MD) except for fetal bovine serum (FBS), which was purchased from Hyclone (Logan, UT). HPLC-grade solvents and ScintiVerse scintillation cocktail were from Fisher Scientific (Pittsburgh, PA). All other chemicals and reagents were of molecular biology or ultrapure grade.

Cell culture. The murine myeloid cell lines 32D (clone 23) and WEHI-3B were kindly provided by Dr. Robert Redner (Univ. of Pittsburgh) and were cultured essentially as previously described (8). We used 32D cells transfected with an empty mammalian expression vector containing the neomycin resistance gene (32D/neo cells) that had served as negative control cells in previous studies examining Bcl-2 overexpression (8). These cells are phenotypically identical to wild-type 32D cells with the exception that they are capable of growth in G-418-containing media. Before drug exposure, 32D cells were seeded at 1.5 × 105 cells/ml in complete media (RPMI with 10% FBS) and cultured for 48 h. The complete RPMI growth media contained 10% media conditioned by WEHI-3B as a source of interleukin (IL)-3, a required survival factor for 32D cells. PQ and QN were freshly prepared as concentrated stocks in serum-free media and were then added to cells at indicated concentrations. PLA2 inhibitors AACOCF3, manoalide, and aristolochic acid were administered from stock solutions prepared in DMSO and stored at -70°C until use. DMSO concentrations never exceeded 0.4% (vol/vol), and control treatments contained DMSO vehicle alone. For IL-3 withdrawal experiments, 32D cells were cultured in complete growth media for 48 h. Cells were then collected, washed three times with serum-free media in the absence of WEHI-conditioned medium, resuspended, and cultured in the original volume of RPMI, with 10% FBS but lacking WEHI-conditioned medium.

Nuclear morphology. Samples of 32D cells (0.5-1 ml) were obtained at the indicated times after treatment and were collected by centrifugation (1,000 g, 2 min). Cells were then washed in PBS, fixed with 2% paraformaldehyde in PBS, and stained with Hoechst 33342 (1 µg/ml) as previously described (8). At least 300 cells were viewed under fluorescent microscopy, with excitation at 340-380 nm, and nuclear morphology was scored as either normal or apoptotic. Apoptotic nuclei were characterized by small, bright-staining nuclei, often very rounded and sometimes fragmented into distinct sections.

DNA fragmentation. Internucleosomal DNA fragmentation was determined by conventional gel electrophoresis in 2% agarose as previously described (8). Briefly, DNA was extracted from 1 × 106 cells by lysis in 10 mM EDTA, 0.5% sarkosyl, and 50 mM Tris (pH 8.0) and sequential digestion in proteinase K (1 mg/ml, 55°C) followed by RNase A (0.5 mg/ml) and RNase T1 (1,000 units/ml) at 37°C. Electrophoresis was performed at 60 V for ~4 h. Gels were then stained with ethidium bromide (1 µg/ml) and then photographed under ultraviolet illumination.

PLA2 activation and AA release. Labeling of cellular phospholipids with [3H]AA and release of radioactivity into tissue culture media were performed essentially as described by Jäättellä (15). 32D/neo cells were obtained from exponential growth-phase cultures, centrifuged (400 g, 10 min), and resuspended (0.8 × 106/ml) in fresh RPMI containing 10% FBS and 10% WEHI-conditioned medium. [3H]AA was added (0.2 µCi/ml), and cells were incubated overnight at 37°C in a 5% CO2 incubator. The following morning, the radiolabeled cells were harvested and washed three times with 10 ml of serum-free medium. Radioactivity in aliquots of the total cell suspension obtained before and after the first centrifugation indicated that ~70% of the radiolabeled AA was incorporated into 32D/neo cells. Cell number and viability (routinely >95%) were determined in the last wash suspension by trypan blue exclusion and cell counting, and the final cell pellet was resuspended in complete medium at 4 × 105 cells/ml. Cells were then seeded into 24-well plates (0.5 ml/well) that contained 100 µl of medium containing test substances or vehicle controls. Individual plates were removed from the incubator at the indicated times and centrifuged (400 g, 10 min), and aliquots (300 µl) of the cell-free medium were placed in scintillation vials. Radioactivity was then determined by scintillation counting after addition of 3 ml of ScintiVerse counting fluid.

The above-mentioned technique was modified to include an analysis of [3H]AA hydrolysis from individual phospholipids after PQ and Ca2+-ionophore stimulation. For these studies, cellular phospholipids were labeled as described above. After overnight incubation, cells were washed as described above with the exception that the first wash in serum-free media contained 0.5 mg/ml of fatty acid-free human serum albumin to assist in the removal of free AA. Washed cells were resuspended to 4 × 105 cells/ml, and 15 ml were seeded into a T-25 flask for each condition. Test substances were then applied, and cells were incubated at 37°C for 3 h, at which time two 7-ml aliquots were removed from each flask and centrifuged (400 g, 10 min). Supernatants were decanted, and cell pellets were resuspended in 1.5 ml of PBS and quick- frozen in a methanol-dry ice bath followed by storage at -70°C until assay. Lipids were extracted from thawed samples by addition of 10 ml of chloroform-methanol followed by vigorous mixing (4 min), centrifugation, and collection of the lower organic phase. The remaining aqueous phase was reextracted with 4 ml of chloroform and combined with the first organic phase. Organic phase was then evaporated to dryness under nitrogen, and the residue was reconstituted in 250 µl of chloroform-methanol (2:1). Pi was determined on 30 µl of this sample as described (1), and the remainder was applied to a 5 × 5-cm silica G TLC plate (Whatman, Clifton, NJ). Lipid classes were separated by two-dimensional high-performance TLC using a solvent system of chloroform-methanol-28% ammonium hydroxide (65:35:5) in the first direction and chloroform-acetone-methanol-glacial acetic acid-water (50:20:10:10:5) in the second. Lipids were visualized by exposure to iodine vapor and identified by comigration relative to purified standards. Identified spots were then scraped from the plate into scintillation vials, lipids were extracted from gel by addition of 250 µl of chloroform-methanol (1:1), and radioactivity was determined after the addition of 3 ml of BetaMax scintillation fluid.

Lipid peroxidation. The use of cis-PNA to measure oxidation of specific phospholipid classes has previously been described (8). Briefly, 32D/neo cells in log-phase growth were allowed to incorporate cis-PNA (4 µg/ml, final concentration) in serum-free RPMI containing 5% WEHI-conditioned medium for 2 h (1 × 106 cells/ml). cis-PNA was given in complex with human serum albumin previously prepared by addition of 500 µg of cis-PNA to 50 mg of human serum albumin in 1 ml of PBS. After repeated washing of cells to remove unincorporated cis-PNA, cells were placed in complete media and incubated with the various test substances for two additional hours. Aliquots of cells were then removed, centrifuged at 1,000 g for 2 min, and washed once with PBS. Cell suspensions were immediately transferred to cold methanol containing butylated hydroxytoluene (0.1 mg/ml). Total lipids were extracted with a modified Folch procedure and subjected to HPLC for separation of phospholipids as previously described (8). The amount of cis-PNA fluorescence in individual phospholipid classes was normalized to the amount of Pi contained within the total lipid extract. Pi was determined spectrophotometrically using the method of Chalvardjian and Rubnicki (5).

Statistical analyses. The effects of various PLA2 inhibitors on cell viability, apoptotic morphology, and lipid peroxidation treatments were first assessed by a one-way ANOVA followed by Dunnett's multiple comparisons with control group, which in most cases represented the PQ alone-treated group. Levels for significant difference were set at P < 0.05.

    RESULTS
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Abstract
Introduction
Methods
Results
Discussion
References

Effect of PLA2 inhibitors on PQ-induced apoptosis and PS oxidation. Apoptosis in response to the prototypical oxidant PQ, which is characterized by chromatin condensation and fragmentation, internucleosomal DNA cleavage, and loss of cell viability within 24 h of toxin exposure, was previously described (8). We first determined whether PQ-induced apoptosis was affected by PLA2 inhibitors.

Figure 1 shows the concentration-dependent ability of the PLA2 inhibitor QN to inhibit the changes in nuclear morphology and cell viability after exposure to PQ. Figure 1A shows that 400 µM PQ alone killed ~30% of the 32D/neo cells within 24 h. In contrast, when QN was simultaneously included during the PQ exposure, there was a concentration-dependent preservation of cell viability that achieved >50% protection at 5 µM QN. Similarly, Fig. 1B demonstrates that QN also inhibited the PQ-induced nuclear changes. Viability of cells receiving 1 and 5 µM QN alone were unchanged compared with untreated control cells (89.3% for 5 µM QN, 89.6% for 1 µM), and changes in nuclear morphology were not observed. Preliminary studies revealed that higher concentrations of QN (10-50 µM) were toxic by themselves (data not shown) and not included for further analyses. Inhibition of PQ-induced apoptosis by QN was confirmed by analysis of internucleosomal DNA fragmentation. Figure 2 shows that 5 µM QN reduced the formation of low-molecular-weight DNA fragments after PQ exposure.


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Fig. 1.   Quinacrine (QN) protected 32D cells transfected with an empty mammalian expression vector containing the neomycin resistance gene (32D/neo cells) from paraquat (PQ)-induced cytotoxicity and apoptosis. 32D/neo cells were seeded at 1.5 × 105/ml in T-25 flasks and cultured for 48 h. Cells were then treated with or without PQ (400 µM) in presence or absence of QN (1 and 5 µM). Twenty-four hours later, cells were harvested, viability was determined by trypan blue exclusion (A), and percentage of apoptotic nuclei was determined using Hoechst 33342 fluorescence (B). Data are means ± SE of 4 individual observations. Each group was compared relative to PQ treatment alone using 1-way ANOVA and Dunnett's multiple comparison with control cells (* P < 0.05, ** P < 0.01).


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Fig. 2.   QN reduced internucleosomal DNA fragmentation in 32D/neo cells after PQ exposure. 32D/neo cells were cultured and treated as described in Fig. 1. After 24 h of drug treatment, 1 × 106 cells were obtained, and DNA was prepared by sequential proteinase K and RNase digestion. DNA was subjected to electrophoresis in a 2% agarose gel. Gel was stained with ethidium bromide and photographed under ultraviolet illumination. Note formation of extensive internucleosomal DNA fragmentation, corresponding to 180- to 200-bp ladders, that occurs with PQ (400 µM) and was substantially reduced with simultaneous inclusion of 5 µM QN. +, Presence; -, absence.

To mechanistically attribute the protective actions of QN to PLA2 inhibition, it was important to assess the protective potential of multiple and structurally diverse PLA2 inhibitors. Because the specificity of QN for various PLA2 types [secretory and cytosolic PLA2 (sPLA2 and cPLA2, respectively)] is not firmly established, we chose pharmacological agents with varying specificity for sPLA2 and cPLA2. Manoalide produces selective inhibition of sPLA2, whereas AACOCF3 specifically antagonizes cPLA2. Aristolochic acid possesses mixed activity toward both enzyme types. Figure 3 shows the effects of these three specific PLA2 inhibitors on PQ-induced apoptosis. 32D/neo cells were treated with 400 µM PQ, which produced ~40% apoptotic cells within 24 h. None of these PLA2 inhibitors attenuated PQ-induced apoptosis. Surprisingly, two agents, AACOCF3 (50 µM) and aristolochic acid (50 and 250 µM), enhanced apoptosis to over 80% when combined with PQ treatment, which is similar to the apoptosis seen with AACOCF3 and aristolochic acid alone. None of the compounds reduced DNA fragmentation compared with PQ alone. Thus, among inhibitors of PLA2, QN was unique in its ability to ameliorate the apoptotic process after exposure to the prototypical oxidant PQ.


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Fig. 3.   Specific inhibitors of phospholipase A2 (PLA2) failed to protect 32D cells from PQ-induced apoptosis. 32D/neo cells were seeded as described in METHODS and cultured for 48 h. Cells were treated with PQ (400 µM) in absence or presence of indicated concentrations of PLA2 inhibitors (brackets): arachidonyl trifluoromethyl ketone (AACOCF3), aristolochic acid, and manoalide. Twenty-four hours later, nuclear morphology was evaluated using Hoechst 33342 fluorescence (similar to Fig. 1B), and genomic DNA was analyzed by gel electrophoresis (similar to Fig. 2). Note that none of these PLA2 inhibitors could inhibit apoptosis similar to QN and, in several instances, enhanced PQ-induced apoptosis. Nos. on top, concentrations of respective inhibitors in µM.

PLA2 activity is not modulated by PQ or QN. Because oxidative stress can potentially regulate PLA2 activity, we next determined the activity of this important regulatory enzyme after PQ exposure. We also examined whether the concentration of QN used to protect 32D cells from PQ apoptosis was sufficient to inhibit PLA2 activity in these cells. For these experiments, PLA2 activity was assessed by the release of metabolically incorporated [3H]AA in the presence and absence of PQ-induced oxidative stress. The Ca2+ ionophore 4-Br-A-23187 was used as a positive control. Figure 4 shows the time-dependent release of [3H]AA into the medium over 7 h of treatment in the absence (Fig. 4A) and presence (Fig. 4B) of 400 µM PQ. In addition, Fig. 4C depicts the activation of PLA2 after 1 µM 4-Br-A-23187. Note the rapid eightfold increase in AA release within 1 h of 4-Br-A-23187 treatment compared with control treatment, which indicated the expression of functional Ca2+-dependent PLA2 within these cells. In contrast, oxidative stress induced by 400 µM PQ failed to lead to significant enzyme activation. The inclusion of 5 µM QN in all cases failed to attenuate the basal or Ca2+-activated release of [3H]AA and indicated that antiapoptotic concentrations of QN were below the level required to inhibit PLA2 activity.


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Fig. 4.   PQ and QN did not affect PLA2 activation. 32D/neo cells were labeled overnight with [3H]arachidonic acid (AA) as described in METHODS. Radiolabeled cells were then harvested, washed, and seeded into 24-well plates (4 × 105 cells/0.5 ml) containing 100 µl of vehicle control (A), PQ (final concentration, 1 mM; B), or 4-bromo-A-23187 (4-Br-A-23187; 10 µM; C) with or without QN (5 µM). At indicated times after drug treatment, plates were removed and centrifuged. Aliquots of cell-free media were then removed for quantification of radioactivity. Data are means ± SE (SE within size of symbol) of triplicate observations from 1 experiment that was replicated a second time. Note 8-fold increase in release of [3H]AA from cells into medium after 4-Br-A-23187 stimulation. PQ produced no release of [3H]AA above that seen in untreated cells. QN had no effect on basal or Ca2+ ionophore-induced PLA2 activity.

We also directly evaluated phospholipid hydrolysis after PQ and QN treatment of 32D cells by preparing total lipid extracts from [3H]AA metabolically labeled cells. The amount of [3H]AA-derived radioactivity was then determined in individual phospholipid and free fatty acid pools after separation by TLC to assess PLA2 activation. A typical chromatogram of lipids extracted from 32D/neo cells is illustrated in Fig. 5. Phosphatidylcholine (PC) represented about one-half (51.1 ± 2.1%) of the total phospholipid, with phosphatidylethanolamine (PEA) being the next most prominent phospholipid (20.1 ± 1.6%). Additional phospholipids detectable on the plates of abundance were sphingomyelin (8.9 ± 0.8%), PS (8.0 ± 1.0%), phosphatidylinositol (PI; 8.0 ± 0.9%), diphosphatidylglycerol (2.8% ± 0.4%), lysophosphatidylcholine (0.6% ± 0.2%), and phosphatidic acid (0.3 ± 0.1%). No significant differences in the pattern of distribution of phospholipid classes were apparent between control, PQ-treated, and 4-Br-A-23187-treated cells. As shown in Table 1, essentially no release of [3H]AA was observed from any of the phospholipid classes studied after a 3-h PQ exposure. These data substantiate the conclusion that PQ failed to activate PLA2 before apoptosis. In contrast, the Ca2+ ionophore 4-Br-A-23187 induced release from several major phospholipid classes. After a 3-h treatment with 1 µM 4-Br-A-23187, ~50% of the [3H]AA was lost from PEA and 40% was lost from PC and PI. Thus the Ca2+-regulated PLA2 activity in 32D/neo cells appeared to recognize multiple phospholipid substrates. Notably, PS was resistant to hydrolysis by PLA2 despite preferential oxidation of this phospholipid during PQ-induced oxidative stress. Essentially identical results were observed when QN was included during the 4-Br-A-23187 treatment, with 43, 47, and 33% of the [3H]AA released from PEA, PI, and PC, respectively. In addition, the neutral lipid fraction containing triglycerides lost ~30% of its content of [3H]AA after 4-Br-A-23187 treatment. Despite the significant release of [3H]AA from multiple lipid sources within 32D/neo cells after 4-Br-A-23187 treatment, no significant increase in radioactivity was observed within the free fatty acid pool within these cells (data not shown), indicating that hydrolyzed AA was rapidly transported from the cell into the extracellular medium.


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Fig. 5.   A typical high-performance TLC 2-dimensional chromatogram of a total lipid extract from 32D cells. Lipids were isolated and subjected to 2-dimensional TLC as described in METHODS. FFA, free fatty acids; NL, neutral lipids; DPG, diphosphatidylglycerol; PI, phosphatidylinositol; PEA, phosphatidylethanolamine; PS, phosphatidylserine; PC, phosphatidylcholine; SPH, sphingomyelin; PA, phosphatidic acid; LPC, lysophosphatidylcholine.

                              
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Table 1.   Release of [3H]AA from individual phospholipid classes after paraquat and Ca2+-ionophore treatment of 32D/neo cells

Table 2 shows the specific activity of [3H]AA incorporation normalized to Pi content of each lipid for four major classes of phospholipid and demonstrates that differences in specific incorporation of AA did not determine substrate specificity. Note that PEA is approximately eightfold enriched in AA compared with PC, and each appears to lose a similar percentage of [3H]AA after 4-Br-A-23187 stimulation. PS was found to be relatively resistant to hydrolysis; however, PS and PC showed similar specific incorporation of AA. Thus, although PS and PC are similarly enriched in AA, PS does not represent as efficient a substrate for PLA2 as does PC in 32D cells.

                              
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Table 2.   Specific incorporation activities of AA in various phospholipid classes in 32D cells

QN inhibits early PS oxidation after PQ. We next examined whether QN could act during the initiation of apoptosis after PQ. On the basis of the previous observation of selective early oxidation of PS early during PQ-induced apoptosis (8), we measured PQ-dependent oxidation of cis-PNA incorporated into individual phospholipid classes with or without QN. Figure 6 shows that QN blocked the selective oxidation of PS after PQ. Within 2 h, PQ oxidized ~25% of the cis-PNA covalently incorporated into PS, whereas no significant loss was observed in other phospholipid classes, including PC, PEA, PI, and sphingomyelin. The simultaneous inclusion of 5 µM QN, however, completely prevented the PQ-dependent oxidation of PS. Exposure of 32D/neo cells to QN alone had no effect on the peroxidation of any of the phospholipid classes studied here.


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Fig. 6.   QN blocked selective peroxidation of PS after PQ exposure. 32D/neo cells with incorporated cis-parinaric acid (cis-PNA) were treated with or without PQ (1 mM for 2 h) in presence or absence of QN (5 µM), followed by extraction of phospholipids and resolution by HPLC. Unoxidized cis-PNA content of each phospholipid class was quantified using arbitrary fluorescence units in each peak and then normalized to Pi content of each peak. Data are means ± SE of cis-PNA content of each phospholipid class [PC (A), PEA (B), PS (C), PI (D), and SPH (E)] based on 7 observations obtained from 2 separate experiments. Statistical analysis was performed by 1-way ANOVA and Dunnett's multiple comparisons with control group. * Significant difference from PQ treatment alone (P < 0.01).

QN does not inhibit apoptosis after IL-3 withdrawal. We next studied whether the effects of QN were specific for PQ-induced apoptosis or whether the cytoprotective effects of this compound extended to other apoptotic stimuli. 32D cells depend on the cytokine IL-3 for continued survival in culture and undergo extensive apoptosis after the removal of this growth factor. Figure 7 shows the time course of apoptosis (Fig. 7A) and the formation of internucleosomal DNA cleavage (Fig. 7B) after IL-3 withdrawal in the presence and absence of 5 µM QN. Note that the time-dependent increase in cells that exhibited nuclear condensation and fragmentation 48 h after IL-3 withdrawal was similar in the absence and presence of QN. Thus, in contrast to its ability to inhibit PQ-induced apoptosis, QN failed to protect 32D/neo cells from apoptosis after IL-3 withdrawal.


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Fig. 7.   QN failed to inhibit apoptosis after interleukin (IL)-3 withdrawal. 32D/neo cells were seeded at 1.5 × 105 cells/ml for 48 h. Cells were washed and resuspended in complete culture medium with or without IL-3-containing WEHI-conditioned medium. QN (5 µM) was also supplied to indicated cells at this time. After 24 or 48 h, aliquots of cells were processed to assess nuclear morphology (A) and internucleosomal DNA fragmentation (B). Data are means ± SE of 3-4 individual observations. Note that QN had no effect on induction of apoptotic nuclear morphology or internucleosomal fragmentation after IL-3 withdrawal. Stds, standards.

    DISCUSSION
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Abstract
Introduction
Methods
Results
Discussion
References

PQ-induced PS oxidation and apoptosis are independent of early PLA2 activation. Considerable evidence has emerged supporting a role for PLA2 in the potentiation of cell injury and death. TNF is a cytokine capable of mediating apoptotic cell death on sensitive cell types and serves as an example of PLA2 involvement with cell death. First, Voelkel-Johnson et al. (42) reported a positive correlation between cPLA2 activity in various cell lines and their ability to be killed by TNF. Second, inhibition of cPLA2 activity with AACOCF3 and anti-sense constructs ameliorates the cytotoxic effect of TNF on adenovirus-infected cells (40). Third, TNF-resistant L929 cells isolated by Hayakawa et al. (13) lack cPLA2, and sensitivity to TNF can be partially restored by transfection of cPLA2 cDNA into these cells. PLA2 has also been implicated in other examples of cell death, including glutamate neuronal toxicity (12) and ischemic injury (2, 32).

Therefore, we initially hypothesized that early PLA2 activation could play a role in mediating apoptosis after PQ exposure. Several observations, however, refute this notion. First, several specific inhibitors of PLA2, such as AACOCF3, manoalide, and aristolochic acid, could not inhibit PQ-induced apoptosis. Second, using two distinct assays, we observed absolutely no evidence for activation of PLA2 after PQ exposure. Third, the concentrations of QN required to inhibit apoptosis in our studies were significantly less than those frequently used by others to inhibit PLA2 in vitro (3, 23) and failed to exert an inhibitory effect on PLA2 measured in these studies. Thus PLA2 activation does not appear to immediately follow PS oxidation and participate in apoptosis after PQ. Therefore, PQ-induced apoptosis does not resemble TNF-mediated cell death in terms of its dependence on PLA2 activation.

It is important to note, however, that PLA2 activation may play a role in other forms of cell death, such as necrosis, after oxidant exposure. Our goal was to specifically study apoptosis; therefore, we chose a concentration of PQ that killed cells primarily by apoptosis (8). In this previous study, ~50% of 32D/neo cells were nonviable after 24-h exposure to 400 µM PQ. On analyzing apoptotic nuclei, we extrapolated that over 80% of those dead cells had undergone apoptosis. Thus nonapoptotic cell death contributes little to the overall cytotoxicity of PQ under these conditions. It is possible that PLA2 blockers may have protective effects in blocking necrotic cell death; however, experimental testing of this hypotheses requires a model of cell death different from that applied here.

It could be argued that the failure of PLA2 inhibitors to attenuate apoptosis resulted from using concentrations below those required for enzyme inhibition. We therefore assessed the ability of these PLA2 inhibitors to attenuate AA release in our experimental system. Neither 2 h of basal or Ca2+-induced AA release was affected by 1 or 5 µM manoalide. This inhibitor, however, is relatively specific for the 14-kDa sPLA2 and has a reported EC50 of 0.2-0.3 µM for inhibiting Ca2+-induced eicosanoid production in peritoneal macrophages (26). Thus the failure of manoalide to inhibit PLA2 in our studies suggests the predominance of an 85-kDa cPLA2 in controlling Ca2+-dependent AA release in 32D cells. In contrast, 250 µM aristolochic acid reduced 4-Br-A-23187-induced AA release by ~40% but failed to protect against PQ apoptosis. Although still somewhat selective for sPLA2, aristolochic acid probably possesses efficacy against cPLA2 at this concentration (43). The cPLA2-specific drug AACOCF3 paradoxically increased basal AA release (2-fold at 10 µM and 9-fold at 50 µM). This release of AA, however, could not be temporally separated from a dramatic and rapid incidence in apoptosis in these cells (Fabisiak, unpublished observations). The nature of this AA release is currently under investigation but probably does not represent cPLA2 activation, since the AACOCF3 concentrations used here were above those required to inhibit cPLA2 in other cell types (33). Instead, it may represent the shedding of membrane-bound apoptotic bodies or sPLA2 activation that could occur as a sequela to apoptosis. Regardless of how these data are interpreted, they still support that notion that PLA2 activation is not responsible for the initiation of PQ-induced apoptosis.

Surprisingly, we found no evidence for the activation of PLA2 during PQ-induced oxidative stress and lipid peroxidation in these cells. These data contrast those of Salgo et al. (35), who found oxidized phospholipids to be preferred substrates for PLA2. Thus PS oxidation observed after PQ may not be a sufficient signal to enhance PLA2 activity. It is possible, however, that because PS incorporated relatively little AA, PS oxidation occurs primarily within other polyenoic fatty acids. Thus hydrolysis and repair of oxidized PS could have gone undetected using AA release in these studies. It should be stressed, however, that even small amounts of oxidized PS could participate in modulation of various aspects of apoptosis. For example, PS oxidation could lead to PS externalization by specific interaction and inhibition with lipid-transporting activities such as aminophospholipid translocase. It is also possible that PLA2 activities can be differentially regulated under various oxidative stresses. Although cPLA2 appears activated after H2O2 treatment (3), porcine pancreatic sPLA2 appears to be directly inhibited by PQ and other superoxide anion-generating systems (11). Thus different lipid-based signaling pathways may be operative for the induction of apoptosis: those that are PLA2 dependent, such as after TNF exposure, and those dependent on PS oxidation, such as after PQ, that are independent on PLA2 activation.

To our knowledge, this is the first report of PLA2 substrate specificity in these nontransformed 32D murine myeloid progenitor cells. Multiple PLA2 types have been described that differ in structure, substrate specificity, and Ca2+ sensitivity (7). It is possible that activation of a Ca2+-independent PLA2 could be involved in signaling apoptosis, since most of the inhibitors chosen for this study are restricted in their effects to the Ca2+-requiring enzymes. This is unlikely, however, in light of the fact that we detected no AA release or phospholipid hydrolysis after PQ, which would reflect activity of any activated PLA2. Although we have not as yet characterized the specific PLA2 types responsible for AA liberation after 4-Br-A-23187 stimulation of 32D cells, it is possible to state that Ca2+-dependent PLA2 in 32D cells can utilize AA-containing PEA, PI, and PC as substrates. In contrast, PS appears to be a relatively poor substrate for hydrolysis by PLA2 despite its preferential oxidation during PQ-induced apoptosis.

QN has antiapoptotic effects not shared by other PLA2 inhibitors. We have shown that PQ-induced apoptosis can be inhibited by the agent QN. The cytoprotective effects of QN were clearly demonstrated by its ability to maintain cell viability and inhibit several characteristics of apoptosis, including internucleosomal DNA fragmentation and chromatin condensation. Importantly, our data also indicate that QN acted early after PQ exposure to disrupt a potential proapoptotic signal, namely, PS oxidation. Selective oxidation of PS is associated with apoptosis and is inhibited by bcl-2 overexpression (8). Thus QN and Bcl-2 may inhibit signal transduction pathways that utilize oxidation of specific membrane phospholipids as downstream mediators for apoptotic signaling after oxidative stress. Our laboratory has also observed that PS oxidation associated with apoptosis appears uniquely resistant to vitamin E analogs (unpublished observations).

It is important to note, however, that QN was ineffective at inhibiting apoptosis after IL-3 withdrawal, whereas overexpression of bcl-2 greatly attenuates apoptosis in this model (29) (data not shown). Thus significant differences must exist between these two models with respect to the action of QN. Many diverse stimuli, including oxidative stress (4, 14, 8), growth factor withdrawal (29), and TNF exposure (22) among others, presumably converge at some point to initiate a common apoptotic signaling cascade. It is likely that Bcl-2 acts early to disrupt the initiation of this final common pathway of apoptosis that includes release of cytochrome c from mitochondria (20), activation of caspase proteases (31), and possibly PS oxidation. One potential locus for the QN effect observed here is within an oxidant-specific pathway not shared by other stimuli of apoptosis such as growth factor deprivation.

QN inhibits TNF cytotoxicity (21) and neuronal cell death after glutamate exposure (12). In addition, QN protects myocytes and neurons from ischemic cell death (2, 32) and the kidney from cyclosporin-mediated toxicity (19). These studies usually attribute the cytoprotective actions of QN to its ability to act as an inhibitor of PLA2 (6, 24). QN, however, is a prototypical cationic amphiphilic compound similar to local anesthetics and chloroquine, with the potential to interact directly with membrane phospholipids (16) and stabilize a variety of organelle membranes including lysosomes (23). The presence of secondary and tertiary amine groups within the QN molecule makes possible their protonation at physiological pH and physical interaction specifically with acidic phospholipids such as PS. In addition, QN can modulate a variety of other membrane-associated functions such as Ca2+ and K+ channels (28, 34) and acetylcholine receptors (16). QN itself, however, did not appear to modulate Ca2+-dependent cellular processes in these studies, since it failed to alter either basal or Ca2+ ionophore-induced [3H]AA release (Fig. 4). It is also possible that QN cytoprotection may be due to a direct antioxidant action. QN can inhibit the release of oxygen radicals from human alveolar macrophages (39) as well as suppress superoxide production in a cell-free xanthine-xanthine oxidase system (23, 41). The effects of QN on blocking PQ-induced PS oxidation support this hypothesis. Studies are currently underway to further elucidate the molecular mechanisms of QN cytoprotection during oxidative stress.

In summary, we showed that QN inhibited selective oxidation of PS and subsequent apoptosis after PQ exposure. These effects appeared independent of PLA2 inhibition. In addition, no evidence of PLA2 activation was observed after PQ. Therefore, PS oxidation and other components of apoptosis after PQ did not require PLA2 activation and suggest that multiple apoptotic signaling pathways exist. Some, as in the case of TNF, appear to depend on PLA2 activation, whereas others such as PQ utilize selective oxidation of membrane phospholipids and are PLA2 independent. In addition, these data point to a novel mechanism for QN cytoprotection not shared by other PLA2 inhibitors. Clearly, care must be taken in the interpretation of experiments utilizing QN as a putative PLA2 inhibitor. Data suggest the potential usefulness of compounds structurally similar to QN as tissue protective agents during exposure to oxidative agents such as PQ.

    ACKNOWLEDGEMENTS

We thank Dr. Daniel Johnson for the kind gift of the 32D/neo cells and for helpful comments in the preparation of this manuscript.

    FOOTNOTES

This work was funded, in part, by American Lung Association Research Grant (to J. P. Fabisiak), National Cancer Institute Grant CA-61299 (to J. S. Lazo), American Institute for Cancer Research Grants 9A50 (to J. S. Lazo) and 97B128 (to V. E. Kagan), Johns Hopkins Center for Alternatives to Animal Testing Grant 96008 (to V. E. Kagan), National Cancer Institute Oncology Research Faculty Development Program (V. A. Tyurin), and International Neurological Science Fellowship Program F05 NS10669, administered by National Institute of Neurological Disorders and Stroke, National Institutes of Health in collaboration with Unit of Neuroscience, Division of Mental Health and Substance Abuse, World Health Organization (Y. Y. Tyurina).

V. A. Tyurin and Y. A. Tyurina are on leave from the Institute of Evolutionary Physiology and Biochemistry, Russian Academy of Science, St. Petersburg, Russia.

Address for reprint requests: J. P. Fabisiak, Dept. of Pharmacology, E1313 Biomedical Science Tower, Univ. of Pittsburgh, Pittsburgh, PA 15261.

Received 15 September 1997; accepted in final form 6 February 1998.

    REFERENCES
Top
Abstract
Introduction
Methods
Results
Discussion
References

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