Involvement of Cytosolic Phospholipase A₂ and Secretory Phospholipase A₂ in Arachidonic Acid Release from Human Neutrophils

Materials

Butylated hydroxytoluene, leupeptin, pepstatin, PMSF, fMLP, and [²D₈]AA were from Sigma (St. Louis, MO). Reagents for Krebs-Ringer-phosphate dextrose buffer (KRPD), including NaCl, KCl, Na₂HPO₄, K₂HPO₄, MgCl₂, and CaCl₂, were obtained from Mallinckrodt (Paris, KY). [³H₈]AA (sp. act., 180 Ci/mmol) was from New England Nuclear (Mississauga, ON, Canada). Organic solvents were from Fisher (Toronto, ON, Canada), and were HPLC grade or better. N-Methyl-N-(tert-butyldimethylsilyl) trifluoroacetamide was from Pierce (Rockford, IL). Aminopropyl columns were obtained from Burdick and Jackson (Muskegon, MI), and Sep-Pak silica columns were obtained from Waters (Toronto, ON, Canada). The DB-23 cyanopropyl gas chromatography column was purchased from J&W Scientific (Folsom, CA). The PLA₂ inhibitors HELSS and MAFP were obtained from Calbiochem (San Diego, CA), and the anti-cPLA₂ Ab was from Santa Cruz Biotechnology (Santa Cruz, CA). SLO toxin was obtained from Scantibodies (South San Francisco, CA). The sPLA₂ inhibitor 3-(3-acetamide-1-benzyl-2-ethylindolyl-5-oxy)propanephosphonic acid (LY311-727) was the kind gift of Dr. E. Mihelich (Lilly Research Laboratories, Indianapolis, IN), and the lysosomal-type Ca²⁺-independent PLA₂ inhibitor 1-hexadecyl-3-trifluoroethylglycero-sn-2-phosphomethanol (MJ33) (27) was kindly provided by Dr. M. K. Jain (Department of Chemistry and Biochemistry, University of Delaware, Newark, DE). The neutralizing anti-sPLA₂ Ab 3F10 (28) and sPLA₂ inhibitor SB203347 (29) were kindly provided by Dr. Lisa Marshall, Smith Kline Beecham Pharmaceuticals (King of Prussia, PA). pldA⁻ Escherichia coli was the kind gift of Dr. Peter Elsbach, New York University (New York, NY).

Isolation of neutrophils

Neutrophils from whole blood were isolated by dextran sedimentation and discontinuous plasma Percoll gradient centrifugation and resuspended in KRPD buffer without Ca²⁺ at 2 × 10⁷ cells/ml (30, 31). This procedure yielded >95% PMN with >98% viability, as judged by trypan blue exclusion. PMN were treated with 1 mM DFP for 10 min on ice before experimental treatments, radiolabeling, cell lysis, subcellular fractionation, or other experimental procedures. We found that DFP, which prevents degradation of PMN proteins (32, 33), decreased nonspecific background activation and prevented cell clumping, and markedly enhanced cell viability over extended treatment and experimental regimes.

Experimental protocol

PMN were treated with 0.1% DMSO, 10 μM MAFP, 16 μM HELSS, 10 μM SB203347, 10 μM Scalaradial, 10 μM LY311-727, or 1 mM DTT for 0.5 h at 37°C, as indicated in the figure legends. PMN were then pelleted and resuspended at 2 × 10⁷ cells/ml in KRPD with 0.25% fatty acid-free human albumin and 1 mM CaCl₂. Following readdition of 10 μM SB203347, 10 μM Scalaradial, 10 μM LY311-727, or 1 mM DTT, which do not bind covalently and could therefore be washed out when cells were resuspended in KRPD with CaCl₂ and albumin, neutrophils were equilibrated for 5 min at 37°C. Cells were subsequently treated with either 0.1% DMSO or 5 μM cytochalasin B for 2 min and 100 nM fMLP for 3 min at 37°C. Experiments were terminated by centrifugation for 10 s at 14,000 × g.

Acid extraction of sPLA₂ from neutrophils

PMN (2 × 10⁷/ml in KRPD) were mixed with an equal volume of water, brought to pH 1.6 with concentrated H₂SO₄, and stirred overnight at 4°C. Following centrifugation at 14,000 × g for 15 min, the supernatant was dialyzed against 500 vol of 10 mM sodium acetate buffer, pH 4.5, for 24 h. The dialysate was collected and centrifuged at 14,000 × g for 15 min, and the supernatant was used as a source of sPLA₂.

Determination of extracellular and cell-associated sPLA₂ activity: collection of extracellular and cell-associated sPLA₂

PMN (2 × 10⁷) were stimulated with fMLP, and the extracellular sPLA₂ was collected in the supernatant by brief centrifugation at 14,000 × g. The supernatant was concentrated over a Centricon 3000 NMWL ultrafilter and diluted to a total of 270 μl in assay buffer A (150 mM NaCl, 10 mM CaCl₂, 25 mM HEPES, pH 7.4, and 0.25% fatty acid-free human albumin). The cell pellet was resuspended in 270 μl sPLA₂ assay buffer and disrupted on ice water by three 15-s bursts of a 50-W probe sonicator set to 20% amplitude (Sonics and Materials, Danbury, CT).

Radiolabeling of E. coli membranes

The PLA₂-deficient strain of E. coli (pldA⁻) was radiolabeled during the log growth phase with [³H₈]AA, washed three times in KRPD with 0.25% fatty acid-free BSA, autoclaved and rewashed exactly as described (34), and then used as a substrate in the sPLA₂ assay and cPLA₂ assays.

sPLA₂ assay

A total of 60 μl of assay buffer A was combined with 10 μl of [³H₈]AA-labeled E. coli membranes (∼50,000 dpm/assay) and 5 μl of DMSO or MAFP (10 μM), HELSS (16 μM), LY311-727 (10 μM), or 3F10 (1 μg/ml) on ice. sPLA₂ reactions were initiated by the addition of 25 μl of cell lysates, cell supernatants, or sPLA₂ from acid-extracted neutrophils. After 30 min at 37°C, reactions were terminated by the addition of 900 μl of tetrahydrofuran, followed by centrifugation at 14,000 × g for 15 min at 4°C. The reaction was then applied to an aminopropyl column, and the fatty acid fraction was selectively eluted with tetrahydrofuran:acetic acid (49:1), followed by liquid scintillation counting (28). Results are expressed as the percentage of free fatty acid hydrolyzed (dpm generated − nonspecific hydrolysis)/total dpm added.

[³H₈]AA release from [³H₈]AA-labeled, SLO-permeabilized neutrophils

Neutrophils were labeled with [³H₈]AA for 2 h at 37°C, washed three times in KRPD with 0.25% BSA, resuspended in KRPD with 1 mM CaCl₂ and 0.25% BSA, and incubated with 0.1% DMSO, 10 μM LY311-727, or 1 μg 3F10/ml for 5 min at 37°C. Cells were then treated with either SLO toxin (approximately 0.1 μg per ml) or vehicle (KRPD) for 2.5 min in the presence of 5 μM cytochalasin B before stimulation with fMLP. The quantity of SLO added was optimized for each experiment. Incubations were stopped after 10 min by centrifugation, and the supernatant was collected and counted for [³H₈]AA release using a liquid scintillation counter (Beckmann 6500) after the method of Mira et al. (35).

AA mass release

Following cellular stimulation and centrifugation for 15 s at 14,000 × g, 1.5 ml of supernatant from 3 × 10⁷ PMN was collected and immediately transferred to siliconized borosilicate test tubes with 6 ml of chloroform-methanol (2:1, v/v) and 0.01% butylated hydroxytoluene. A total of 8 pmol of the internal standard deuterated AA ([²D₈]AA) was then added to each sample. After vortexing and addition of 1.5 ml NaCl, samples were centrifuged at 1000 × g for 5 min, and the lower phase was collected. The upper phase was then reextracted twice with chloroform-methanol-0.58% NaCl (86:14:1) (36). Lipids from the pooled lower phase were then dried under a stream of N₂, reconstituted in hexane-methyl tert-butyl ether (200:3), and applied to a silicic acid column. Fatty acids were selectively eluted from the column with hexane-methyl tert-butyl ether-acetic acid (100:2:0.2) (37), dried under N₂, and derivatized with N-methyl-N-(tert-butyldimethylsilyl)trifluoroacetamide (38, 39). The tert-butyldimethylsilyl ether of AA was then separated from other fatty acids by gas chromatography (model 5890; Hewlett Packard, Palo Alto, CA) on a DB-23 0.2 mm × 25 m cyanopropyl-methyl column. Following electrospray ionization, AA mass was quantified by selectively monitoring the intensities of AA (m/z 361) and the internal standard [²D₈]AA (m/z 369) with a quadrupole mass spectrometer (model 5971; Hewlett Packard).

Cell fractionation

A total of 6 × 10⁷ cells were lysed in 600 μl 20 mM HEPES, pH 7.4, 150 mM KCl, 5% glycerol, 1 mM EDTA, 1 mM EGTA, 1 mM NaF, 1 mM sodium orthovanadate, 2 mM DFP, 1 mM PMSF, 1 mM benzamidine hydrochloride, 50 μM leupeptin, 50 μM pepstatin, and 50 μM chymostatin by brief pulses of probe sonication on ice water. The insoluble cellular components, including nuclei, were removed by centrifugation at 14,000 × g for 5 min at 4°C. The supernatant was then separated into microsomal and cytosolic fractions by centrifugation at 150,000 × g for 20 min at 4°C.

Determination of PLA₂ activity in PMN cytosol

SDS-PAGE

Cytosolic, microsomal, or nuclear fractions (200 μl) were combined with 50 μl of 5× sample buffer (0.312 M Tris, pH 6.8, 50% sucrose (w/v), 25 mM DTT, 10% SDS, and 0.5% bromophenol blue) to a total volume of 250 μl. The samples were boiled for 5 min, and 25-μl aliquots were resolved on 12% 0.75-mm gels in a Bio-Rad (Richmond, CA) mini protean II vertical electrophoresis apparatus at 100 V.

Western blots

Gels were placed against nitrocellulose membranes in 25 mM Tris and 192 mM glycine with 20% (v/v) methanol and transferred at 100 V for 1 h. The blots were stained with 0.1% Ponceau Red to ensure equal protein loading before destaining in water. Blots were blocked for 2 h in TBST (20 mM Tris, pH 7.3, 0.24 M NaCl, 2.6 mM KCl, and 0.05% Tween 20) containing 5% (w/v) skim milk powder and 1% (v/v) goat normal serum. The blot was then washed 2 × 5 min in 10 ml TBST and incubated with 1/1000 anti-cPLA₂ overnight at 4°C before washing 1 × 5 min in 10 ml TBST and incubation with a 1/20,000 dilution of goat anti-mouse secondary Ab conjugated to HRP for 1 h. Blots were then washed 3 × 5 min in TBST before detection with 1.24 μM 5-amino-2,3-dihydro-1,4-phthalazinedione (luminol), 0.65 μM 4-hydroxycinnamic acid (p-coumaric acid), and 0.0001% hydrogen peroxide against KODAK ECL blue film.

Statistical analysis

Data were analyzed by ANOVA, followed by comparison of all means with the Tukey-Kramer honestly significant difference (HSD) test using SAS software (SAS Institute, Carey, NC). Results were considered to be significantly different when p < 0.05 was observed.

A minor role for extracellular sPLA₂ in PMN AA release

We examined the kinetics of sPLA₂ release by PMN in response to fMLP and found that approximately 30% of the apparent cell-associated sPLA₂ activity was lost within the first 30 s after stimulation (Fig. 1⇓A). Concurrently, the sPLA₂ activity observed in the supernatant rapidly increased in the first 30 s after treatment with fMLP and then remained constant (Fig. 1⇓B). Under these assay conditions, cell-associated and extracellular PLA₂ activity were each inhibited more than 98% by coincubation with 10 μM LY311-727 (Fig. 2⇓A), which selectively inhibits group IIa (16) and group V sPLA₂ activity (17), but has no effect on cPLA₂ or iPLA₂ activity (17). In a similar pattern to sPLA₂ activity release, the greatest change in [³H₈]AA release from neutrophils occurred within 30 s after fMLP stimulation, with a decline in the rate of release over the next several minutes (Fig. 1⇓C). Hence, there was a strong temporal correlation between the secretion of sPLA₂ and the release of [³H₈]AA from fMLP-stimulated PMN.

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FIGURE 1.

Effect of fMLP on cellular sPLA₂ activity, release of sPLA₂ into the extracellular fluid, and [³H₈]AA release. Neutrophils were incubated for 5 min at 37°C, stimulated with fMLP (time = 0 min), and centrifuged. The sPLA₂ activity in cell lysates (A) or extracellular fluid (B) was then measured, as described in Materials and Methods. C, Cells were metabolically labeled with [³H₈]AA, and the time course of [³H₈]AA release following stimulation with fMLP was assessed. Incubation with 0.1% DMSO did not result in any changes in cell-associated sPLA₂ activity, extracellular sPLA₂ activity, or [³H₈]AA release. Results shown are the mean ± SE of three determinations. Treatment differences by ANOVA were significant (p < 0.01).

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FIGURE 2.

Effect of LY311-727 or DTT on extracellular sPLA₂ activity and fMLP-stimulated neutrophil AA mass release. PMN were treated with vehicle (DMSO) or fMLP in the presence of DMSO, LY311-727, or DTT. The release of sPLA₂ activity (A) or AA mass (B) into the supernatant was then measured, as described in Materials and Methods. Results shown are the mean ± SE of four determinations. Different lower case letters indicate differences by the Tukey-Kramer HSD test, p < 0.05.

As the release of sPLA₂ activity correlated with extracellular [³H₈]AA release, we sought to determine whether extracellular sPLA₂ mediated AA mass release. Stimulating PMN with fMLP in the presence of either DTT or the cell-impermeable sPLA₂ inhibitor LY311-727 (16) significantly inhibited extracellular sPLA₂ activity (Fig. 2⇑A). In contrast, incubation with either DTT or LY311-727 only decreased fMLP-stimulated AA mass release by approximately 15%, a difference that failed to reach statistical significance (Fig. 2⇑B). Therefore, sPLA₂-mediated hydrolysis of AA from glycerophospholipids on the extracellular face of the plasma membrane of PMN does not appear to be a major source of released AA under these conditions.

A central role for cPLA₂ in PMN AA release

cPLA₂ has been implicated in AA release from human neutrophils (40). Using immunoblot analysis, we found that most of the cPLA₂ in unstimulated neutrophils is present in the cytosolic fraction (Fig. 3⇓). The cPLA₂ in the cytosolic fraction was the more rapidly migrating form, indicating that most of the cPLA₂ in this fraction was not phosphorylated (41). Comparatively little cPLA₂ was detected in the nuclear or microsomal fraction of unstimulated PMN. In fMLP-stimulated cells, cPLA₂ was detected in both nuclear and microsomal fractions, and a concomitant decrease in the amount of cPLA₂ in the cytosolic fraction was observed. In addition, the majority of cPLA₂ detected in fMLP-stimulated cells migrated at a relatively higher m.w., demonstrating that most of the cPLA₂ that translocated to nuclear and microsomal fractions was phosphorylated (41).

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FIGURE 3.

Effect of fMLP on cPLA₂ translocation and electrophoretic mobility. PMN were treated with DMSO or fMLP, disrupted by sonication, and separated into cytosolic, microsomal, and nuclear fractions by centrifugation. After SDS-PAGE, immunoblot analysis was performed with an anti-cPLA₂ Ab. cPLA₂ has a calculated molecular mass of 85 kDa, but migrates at a molecular mass of approximately 110 kDa on SDS-PAGE (41 ). cPLA₂ phosphorylation was detected by a decrease in electrophoretic mobility. Similar results were obtained in three separate experiments.

We then examined the effect of the cPLA₂ inhibitor MAFP on AA mass release from fMLP-stimulated cells. We controlled these pharmacological experiments with the putative sPLA₂ inhibitors SB203347 and Scalaradial, which have previously been shown to inhibit AA release (23, 24), and with the iPLA₂ inhibitor HELSS. We observed that MAFP decreased fMLP-induced AA mass release from intact PMN to levels similar to those observed with the cell-permeable sPLA₂ inhibitor SB203347, while preincubation with Scalaradial resulted in a further decrease in AA mass release (Fig. 4⇓). The iPLA₂ inhibitor HELSS had little effect on AA release at a concentration that has previously been shown to decrease cytosolic PMN iPLA₂ activity by 80% (42). Importantly, MAFP had no effect on the catalytic activity of sPLA₂ that had been acid extracted (43) from neutrophils (Fig. 5⇓). As expected, neutrophil sPLA₂ activity was significantly decreased by LY311-727 and the neutralizing anti-sPLA₂ Ab 3F10, and was not affected by coincubation with HELSS (Fig. 5⇓).

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FIGURE 4.

Effect of cell-permeable PLA₂ inhibitors on neutrophil AA mass release. Cells were pretreated with DMSO, HELSS, MAFP, SB203347, or Scalaradial before treatment with DMSO or fMLP. AA mass release was then assessed by gas chromatography-selective ion monitoring mass spectrometry, as described in Materials and Methods. Results shown are the mean ± SE of three determinations. Different lower case letters indicate differences by the Tukey-Kramer HSD test, p < 0.05.

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FIGURE 5.

Effect of PLA₂ inhibitors on acid-extracted neutrophil sPLA₂ activity. sPLA₂ was partially purified from neutrophils by incubation with H₂SO₄. The effect of LY311-727, 3F10, HELSS, or MAFP on acid-extracted sPLA₂ activity was then assessed, as described in Materials and Methods. Results shown are the mean ± SE of four determinations. Different lower case letters indicate differences by the Tukey-Kramer HSD test, p < 0.05.

We then evaluated the effect of MAFP on cPLA₂ activity in intact cells by incubating cells with MAFP and evaluating PLA₂ activity in the cytosolic fraction of these cells. We found that the PLA₂ activity in PMN cytosol, using [³H₈]AA-labeled bacterial membranes as a substrate, was almost completely inhibited by MAFP (Fig. 6⇓). To determine whether the PLA₂ activity in PMN cytosol was attributable to cPLA₂ activity, we controlled this assay against the presence of iPLA₂ activity using the iPLA₂-specific inhibitor HELSS and against sPLA₂ activity using the sPLA₂ inhibitor LY311-727 (16). We found that neither HELSS nor LY311-727 had any effect on AA hydrolysis from bacterial membranes under these conditions (Fig. 6⇓). Two other iPLA₂ inhibitors (PACOCF₃ and MJ33) also failed to inhibit the PLA₂ activity in PMN cytosol (data not shown). Hence, we observed that MAFP significantly blocked cPLA₂ activity in the cytosol of human PMN in vitro, a finding consistent with the capacity of MAFP to inhibit fMLP-induced AA release from intact cells.

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FIGURE 6.

Effect of PLA₂ inhibitors on neutrophil cPLA₂ activity. Cells were disrupted by sonication, and the cytosolic fraction was isolated by centrifugation. The effect of MAFP, HELSS, or LY311-727 on cPLA₂ activity was then measured, as described in Materials and Methods. Results shown are the mean ± SE of four determinations. Different lower case letters indicate differences by the Tukey-Kramer HSD test, p < 0.05.

No role for iPLA₂ in fMLP-induced PMN AA release

We used the suicide substrate HELSS (44) to study the role of iPLA₂ in PMN AA release. HELSS has previously been shown to selectively constrain the activity of iPLA₂, and to be a poor inhibitor of cPLA₂ activity (26). At a concentration that significantly decreased PMN iPLA₂ activity (42) and obliterated PMN superoxide production (45), HELSS failed to inhibit fMLP-induced AA mass release (Fig. 4⇑) or to have any effect on neutrophil sPLA₂ activity (Fig. 5⇑). These findings do not support a role for iPLA₂ in fMLP-stimulated neutrophil AA release.

A potential role for an intracellular or cell-associated sPLA₂-like activity in PMN AA release

We have provided evidence that extracellular sPLA₂ and iPLA₂ make little or no contribution to AA release (Figs. 1⇑ and 4⇑), and that the function of cPLA₂ appears to be required for fMLP-induced AA release (Figs. 3⇑ and 4⇑). As a control for the AA mass release experiments with the soluble cPLA₂ inhibitor MAFP, we examined the effects of the putative sPLA₂-specific inhibitors SB203347 and Scalaradial (23, 24), which, like MAFP, readily penetrate intact cells. We noted that pretreatment with SB203347 or Scalaradial inhibited AA mass release (Fig. 4⇑), findings consistent with a role for an intracellular or cell-associated sPLA₂ in this process. In this regard, activity measurements showed that approximately 70% of cellular sPLA₂ remained within the cell following stimulation with fMLP (Fig. 1⇑A), and thus was inaccessible to LY311-727 or DTT, sPLA₂ inhibitors that failed to affect fMLP-induced AA mass release (Fig. 2⇑B). Hence, we considered whether an intracellular or cell-associated sPLA₂-like molecule could play a role in PMN AA metabolism. To evaluate this hypothesis, we permeabilized PMN with SLO to deliver LY311-727 or 3F10 directly to the putative intracellular or cell-associated sPLA₂. As noted above, LY311-727 and 3F10 inhibit neutrophil sPLA₂ activity by approximately 98% and 80%, respectively (Fig. 5⇑). [³H₈]AA release from permeabilized cells treated with DMSO or DTT were used as controls. After PMN were permeabilized with SLO, incubation with either LY311-727 or 3F10 resulted in a 50% inhibition of [³H₈]AA release (Fig. 7⇓, +SLO), while DTT decreased [³H₈]AA release from permeabilized cells by approximately 40% (data not shown). When [³H₈]AA-labeled cells were not permeabilized with SLO, the cell-impermeable sPLA₂ inhibitor LY311-727 and the neutralizing anti-sPLA₂ Ab 3F10 failed to prevent [³H₈]AA release (Fig. 7⇓, −SLO). Hence, we were able to provide some evidence that an intracellular or cell-associated sPLA₂-like molecule participates in fMLP-stimulated AA release.

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FIGURE 7.

Effect of LY311-727 and 3F10 on [³H₈]AA release from permeabilized neutrophils. Cells were metabolically labeled with [³H₈]AA, washed, and treated with SLO toxin (+SLO) or vehicle (−SLO). Neutrophils were then treated with vehicle or fMLP in the presence of DMSO, LY311-727, or 3F10. Following centrifugation, the release of [³H₈]AA was measured, as described in Materials and Methods. Exposure of SLO-permeabilized cells (i.e., +SLO) to fMLP induced a mean release of 2598 cpm, which was arbitrarily taken as 100%. Results shown are the mean ± SE of four determinations. Different lower case letters indicate differences by the Tukey-Kramer HSD test, p < 0.05.

No direct role for iPLA₂ in fMLP-stimulated PMN AA release

The role of iPLA₂ in AA metabolism appears to vary in different cell lines. In pancreatic β cells, smooth muscle cells, and cardiomyocytes, treatment with the iPLA₂ inhibitor HELSS decreased agonist-stimulated AA release (46, 47, 48). HELSS has been shown to be approximately 1000-fold more selective for inhibition of iPLA₂ compared with cPLA₂ or sPLA₂ under some conditions, and has no effect on other enzymes involved in AA metabolism, including arachidonyl-CoA synthetase, lysophosphatidylcholine:arachidonyl acyl transferase, or CoA-independent transacylase (26, 47, 49, 50, 51). Therefore, the results obtained with HELSS were interpreted to indicate a role for iPLA₂ in AA release from pancreatic β cells, smooth muscle cells, and cardiomyocytes. In addition, human embryonic kidney 293 cells transfected with iPLA₂ cDNA demonstrated increased basal release of [³H₈]AA and increased [³H₈]AA release in response to serum compared with control cells (52). In contrast, studies with P388D₁ macrophages in which HELSS or antisense iPLA₂ oligonucleotides were used to inhibit or deplete iPLA₂ demonstrated that iPLA₂ participates in phospholipid remodeling, and did not play a role in PAF-stimulated AA release from these cells. In the present study, we found that HELSS, at a concentration that significantly inhibits neutrophil iPLA₂ activity (42), had no significant effect on fMLP-stimulated AA release. We conclude that iPLA₂ did not significantly contribute to fMLP-stimulated PMN AA release under our assay conditions.

Central role for cPLA₂ in PMN AA release

Multiple studies have implicated cPLA₂ activation as a critical step in PMN AA release (18, 53, 54, 55). In agreement with these studies, immunoblot analysis of fMLP-stimulated cells demonstrated translocation of cPLA₂ from cytosolic to microsomal and nuclear fractions. In addition, exposure to fMLP resulted in a decrease in the electrophoretic mobility of cPLA₂, a finding consistent with cPLA₂ phosphorylation (41), which is known to increase the catalytic activity of cPLA₂ in vitro (55). Pretreatment of neutrophils with MAFP, which covalently binds to and inactivates cPLA₂ (56), inhibited fMLP-induced AA mass release. In parallel studies conducted in vitro, MAFP obliterated the PLA₂ activity in neutrophil cytosol. While these results are consistent with a role for cPLA₂ in neutrophil AA release, they do not rule out the possibility that MAFP inhibited AA release through an effect on sPLA₂. To evaluate this possibility, sPLA₂ was partially purified from PMN by acid extraction with H₂SO₄, pH 1.6 (43), and the effect of MAFP on neutrophil sPLA₂ activity was assessed. Coincubating acid-extracted sPLA₂ with MAFP had no effect on sPLA₂ activity (Fig. 5⇑), indicating that MAFP did not decrease neutrophil AA release through an effect on sPLA₂. MAFP also inhibits iPLA₂ in vitro (56), but studies with HELSS indicated that iPLA₂ does not participate in neutrophil AA release. Therefore, the observation that fMLP stimulated cPLA₂ phosphorylation and translocation, and that MAFP inhibited fMLP-stimulated neutrophil AA mass release and cPLA₂ activity supports a central role for cPLA₂ in governing AA release from human PMN.

Role for sPLA₂ in PMN AA release

We showed that stimulation with fMLP for 30 s resulted in a significant release of both sPLA₂ activity and [³H₈]AA from PMN. These findings are consistent with the notion that sPLA₂ released by PMN into the extracellular space can bind to the cell membrane and hydrolyze membrane glycerophospholipids (26). In support of this model, addition of exogenous recombinant synovial PLA₂ to P388D₁ macrophages or group V PLA₂ to unstimulated human PMN both led to a significant release of AA (21, 22). In addition, the agonist-induced translocation of phosphatidylserine and phosphatidylethanolamine from the intracellular to the extracellular face of the phospholipid membrane favors membrane hydrolysis by an extracellular sPLA₂, as sPLA₂ binding to lipid bilayers is promoted by negative charges (57, 58), and group V sPLA₂ preferentially hydrolyzes phosphatidylethanolamine vesicles compared with phosphatidylcholine vesicles (59). Our results did not strongly support a role for an extracellular sPLA₂ in PMN AA release, as inhibition of extracellular sPLA₂ activity with LY311-727 or DTT, which constrains the catalytic activity of group IIa and group V sPLA₂ (16, 17), only had a small inhibitory effect on AA mass release from fMLP-stimulated cells. This discrepancy may be explained by the finding that brief exposure to exogenous group IIa or group V PLA₂ caused an initial release of fatty acids from human white blood cells that was followed by resistance to further hydrolysis of membrane phospholipids by either enzyme, a phenomenon that may be mediated by sPLA₂ binding to a membrane receptor (60). Alternatively, it is possible that the affinity of the surface sPLA₂ receptors rapidly changed in response to agonist stimulation, and that stimulated cells did not effectively bind endogenous extracellular sPLA₂. In support of this, we have previously shown that sPLA₂ expression on the surface of PMN increases 7-fold after 15 s of stimulation with fMLP, but that surface sPLA₂ expression returned to baseline levels within 1 min of stimulation (61). We conclude from our data that the sPLA₂ that is released from human PMN in response to fMLP did not directly mediate a major portion of the AA released from these cells.

Possible role for an intracellular or cell-associated sPLA₂-like enzyme in PMN AA release

Previous studies have indicated that a sPLA₂ might play a role in AA release from within the cell (23, 24). Three independent lines of evidence were identified in this study that support this concept. First, we observed that approximately 70% of cellular sPLA₂ activity remained associated with the cell following stimulation with fMLP. Second, treatment with the cell-permeable sPLA₂ inhibitors SB203347 or Scalaradial both prevented AA mass release to an extent similar to MAFP (Fig. 4⇑). However, studies with SB203347 or Scalaradial must be interpreted cautiously (24), as both of these inhibitors may constrain the activity of cPLA₂, and Scalaradial may affect AA release though inhibition of Ca²⁺ mobilization (25). As the reducing environment of the cytosol would inactivate sPLA₂ activity, sPLA₂ would likely have to be confined to intracellular granules (9) or some other intracellular location to retain catalytic activity. To examine the potential role of an intracellular or cell-associated sPLA₂ in AA release under our conditions, we used the highly specific sPLA₂ inhibitor LY311-727 (16) and the neutralizing anti-sPLA₂ mAb 3F10 (28). As neither LY311-727 nor 3F10 was likely to be able to enter cells effectively during the time frame of these studies (minutes), we permeabilized PMN with SLO so that the sPLA₂ inhibitors could gain direct access to the putative intracellular or cell-associated sPLA₂. The nonspecific reducing agent DTT, which inhibits sPLA₂, but not cPLA₂ or iPLA₂ activity, was used as a control. We found that both LY311-727 and 3F10 inhibited the release of [³H₈]AA from permeabilized PMN (Fig. 7⇑) to an extent that was comparable with that of the nonspecific reducing agent DTT, thereby providing a third line of evidence that an intracellular or cell-associated sPLA₂ participates in PMN AA release. In contrast to our results, Bauldry et al. showed that DTT had little effect on [³H₈]AA release from [³H₈]AA-labeled, fMLP-stimulated, SLO-permeabilized PMN (62). This discrepancy may be explained by the fact that the cytosolic buffer used for the permeabilization studies conducted by Bauldry et al. (62) did not support sPLA₂ catalytic activity, most likely because a Ca²⁺ concentration of 500 nM was used. In the present study, cells were simultaneously exposed to fMLP and 1 mM Ca²⁺, a concentration of Ca²⁺ that supports maximal sPLA₂ activity.

As SLO permitted a free diffusion of small molecules between cytoplasm and medium (35), it was not clear whether the [³H₈]AA release inhibited by LY311-727 or 3F10 in the permeabilized cell was in fact destined for extracellular release, or if the [³H₈]AA leaked out of the cell from intracellular compartments such as granules, through the SLO-induced pores. Since LY311-727 did not inhibit cPLA₂ activity (Fig. 6⇑), a finding consistent with a recent report (17), we conclude that a low m.w. sPLA₂ activity remained within a noncytosolic compartment of the neutrophils, such as the granules (9), and participated in AA metabolism in response to the bacterial peptide fMLP.

In summary, our results demonstrate that cPLA₂ plays a central role in fMLP-stimulated AA release from human PMN. We also present some evidence in support of a role for an intracellular or cell-associated sPLA₂ in PMN AA release. The possibility that sPLA₂ activity is regulated by the prior activation of cPLA₂ (26) in human PMN is currently being evaluated.