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J. Biol. Chem., Vol. 275, Issue 24, 18248-18258, June 16, 2000

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Distinct Roles of Two Intracellular Phospholipase A₂s in Fatty Acid Release in the Cell Death Pathway

PROTEOLYTIC FRAGMENT OF TYPE IVA CYTOSOLIC PHOSPHOLIPASE A₂ INHIBITS STIMULUS-INDUCED ARACHIDONATE RELEASE, WHEREAS THAT OF TYPE VI Ca²⁺-INDEPENDENT PHOSPHOLIPASE A₂ AUGMENTS SPONTANEOUS FATTY ACID RELEASE^*

Gen-ichi Atsumi, Makoto Murakami, Kayoko Kojima, Atsuyoshi Hadano, Masae Tajima, and Ichiro Kudo^§

From the Department of Health Chemistry, School of Pharmaceutical Sciences, Showa University, 1-5-8 Hatanodai, Shinagawa-ku, Tokyo 142, Japan

Received for publication, January 13, 2000, and in revised form, February 24, 2000

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Cytosolic phospholipase A₂ (cPLA₂; type IVA), an essential initiator of stimulus-dependent arachidonic acid (AA) metabolism, underwent caspase-mediated cleavage at Asp⁵²² during apoptosis. Although the resultant catalytically inactive N-terminal fragment, cPLA₂(1-522), was inessential for cell growth and the apoptotic process, it was constitutively associated with cellular membranes and attenuated both the A23187-elicited immediate and the interleukin-1-dependent delayed phases of AA release by several phospholipase A₂s (PLA₂s) involved in eicosanoid generation, without affecting spontaneous AA release by PLA₂s implicated in phospholipid remodeling. Confocal microscopic analysis revealed that cPLA₂(1-522) was distributed in the nucleus. Pharmacological and transfection studies revealed that Ca²⁺-independent PLA₂ (iPLA₂; type VI), a phospholipid remodeling PLA₂, contributes to the cell death-associated increase in fatty acid release. iPLA₂ was cleaved at Asp¹⁸³ by caspase-3 to a truncated enzyme lacking most of the first ankyrin repeat, and this cleavage resulted in increased iPLA₂ functions. iPLA₂ had a significant influence on cell growth or death, according to cell type. Collectively, the caspase-truncated form of cPLA₂ behaves like a naturally occurring dominant-negative molecule for stimulus-induced AA release, rendering apoptotic cells no longer able to produce lipid mediators, whereas the caspase-truncated form of iPLA₂ accelerates phospholipid turnover that may lead to apoptotic membranous changes.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Phospholipase A₂ (PLA₂)¹ comprises a growing family of distinct enzymes that exhibit different substrate specificity, cofactor requirements, subcellular localization, and cellular functions (1-11). The family includes at least nine low molecular weight secretory PLA₂ (sPLA₂) isozymes (IB, IIA, IID, IIC, IIE, IIF, III, V, and X), three cytosolic PLA₂ (cPLA₂) isozymes (IVA (), IVB (), and IVC ()), several splicing variants of Ca²⁺-independent PLA₂ (iPLA₂) (VI), and intracellular (VIIB and VIII (1 and 2)) and secretory (VIIA) platelet-activating factor acetylhydrolases. cPLA₂ and several sPLA₂ isozymes are considered signaling PLA₂s, regulating stimulus-induced arachidonic acid (AA) metabolism that is linked to production of bioactive eicosanoids (12-20). iPLA₂ plays a major role in regulation of phospholipid remodeling (21), and recent evidence suggests that it also takes part in lipid signaling under certain conditions (16, 17, 22).

Apoptosis is a process of regulated cell suicide that is crucial for the development and homeostasis of multicellular organisms and is characterized by chromatin condensation, cell shrinkage, and plasma membrane blebbing (23). Although a number of changes in cytosolic and nuclear proteins and chromosomal DNA occur in apoptotic cells (23), little is known about how glycerophospholipid metabolism is affected. Nevertheless, several lines of evidence suggest that perturbation of membrane lipid turnover affects ongoing apoptotic processes (24-28). Extracellular and intracellular PLA₂ isozymes have been implicated in several types of apoptosis. The plasma membrane (or microvesicle) phospholipids of apoptotic or damaged cells are the preferred substrates for several sPLA₂s (29, 30). Type IVA cPLA₂ has been reported to be involved in AA release during cell death in several cell types (31-33). In these studies, suppression of cPLA₂ activity led to a decrease in cell death, whereas overexpression of cPLA₂ enhanced cell death. On the contrary, subsequent studies showed that cPLA₂ is degraded and catalytically inactivated by caspases (34-36), raising the question as to whether cPLA₂ plays some general role in apoptosis. Rather, our recent study showed that increased fatty acid release during Fas-induced apoptosis was sensitive to inhibitors of iPLA₂ rather than of cPLA₂ (34).

In this study, we provide evidence that iPLA₂ indeed mediates enhanced release of fatty acids from apoptotic cells. Deletion of the N-terminal first ankyrin repeat by the action of caspase-3 renders iPLA₂ more active than the uncleaved form. In contrast, the catalytically inactive N-terminal cPLA₂ fragment produced by caspase-3 shows a higher affinity for membranes than does the intact enzyme and behaves like a dominant-negative inhibitor of stimulus-induced AA release by cPLA₂ as well as by sPLA₂-IIA. The latter observation provides further support for a functional linkage between cPLA₂ and signaling sPLA₂.

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Materials-- A23187 was obtained from Calbiochem. The agonistic anti-Fas antibody CH-11 (37) and caspase-3/CPP32 Colorimetric protease assay kit were purchased from Medical & Biological Laboratories. Human recombinant tumor necrosis factor  (TNF) and human recombinant interleukin-1 (IL-1) were purchased from Genzyme. Methylarachidonyl fluorophosphonate (MAFP), which inhibits both iPLA₂ and cPLA₂ (38), bromoenol lactone (BEL), an iPLA₂ inhibitor (39), and rabbit polyclonal anti-iPLA₂ antibody were obtained from Cayman Chemical. Rabbit polyclonal anti-cPLA₂ antibody was from Santa Cruz Biotechnology. The anti-FLAG antibody M2, cycloheximide (CHX), and etoposide were purchased from Sigma. cDNAs for mouse cPLA₂ and hamster iPLA₂ have been described previously (16). Human caspase-3 cDNA was a generous gift from Dr. K. Takahashi (Showa University, Tokyo, Japan). Phospholipids and neutral lipids used as standards for thin layer chromatography (TLC) were purchased from Avanti. MACS apoptotic cell isolation kit was purchased from Miltenyi Biotec. Protease inhibitors and all other regents, which were of analytical grade, were obtained from Wako. Human monocytic U937 cells, human cervix epithelioid carcinoma HeLa cells, and mouse fibroblastic L929 cells were obtained from RIKEN Cell Bank. Human embryonic kidney (HEK) 293 cells were from Health Science Research Resources Bank.

Induction of Apoptosis in U937 Cells-- U937 cells were maintained in RPMI 1640 medium (Nissui Pharmaceutical) supplemented with 10% (v/v) fetal calf serum (FCS) (Intergen), 2 mM glutamine, 100 units/ml penicillin, and 100 µg/ml streptomycin at 37 °C in humidified air containing 5% CO₂. The cells were preincubated with 0.1 µCi/ml [³H]AA (NEN Life Science Products) for 24 h, washed three times, resuspended at 1 × 10⁷ cells/ml in cultured medium, and then treated with 50 or 100 ng/ml anti-Fas antibody or 100 units/ml TNF in the presence or absence of 10 µg/ml CHX for various periods. In some experiments, MAFP or BEL was added to the cells during incubation. The [³H]AA contents of neutral lipids were assessed by counting the radioactivity of a fraction extracted using the method of Dole and Meinertz (40). Cell viability was assessed by the trypan blue dye exclusion test.

TLC Analysis-- The total lipids in radiolabeled cells and their supernatants were extracted using the method of Bligh and Dyer (41) and developed by two-dimensional TLC on Silica Gel 60 plates (Merck), as described previously (34). The first and second solvent systems consisted of chloroform/methanol/acetic acid/water (65/25/4/2, v/v) and chloroform/methanol/formic acid (65/25/8.8, v/v), respectively. The zones on the silica gel corresponding to neutral lipids (a mixture of free fatty acids and other neutral lipids) and phospholipids were identified by comparing their mobilities with those of authentic standards, which were visualized with iodine vapor. The zones were each scraped into a vial, and the radioactivity was counted using a liquid -scintillation counter (Aloka). In order to separate further the free fatty acids from other neutral lipids, the total lipids were developed on Silica Gel 60 plates using a solvent system of hexane/ether/acetic acid (80/30/1, v/v). The radioactivity associated with each lipid was expressed as a percentage of that associated with the total lipids.

Measurement of iPLA₂ Activity-- The cells were washed once with a buffer comprising 10 mM HEPES (pH 7.5), 1 mM EDTA, and 340 mM sucrose, suspended (3.3 × 10⁷ cells/ml) in the same buffer containing 1 mM dithiothreitol, and lysed by sonication for 1 min with a Branson Sonifier (power 30, 50% pulse cycle). The cell lysates were centrifuged at 100,000 × g for 1 h at 4 °C, and the resulting supernatants were incubated at 40 °C for 30 min in 250 µl of buffer comprising 100 mM HEPES (pH 7.5), 5 mM EDTA, 0.4 mM Triton X-100, 0.1 mM ATP, and 15 µM 1-palmitoyl-2-[¹⁴C]arachidonyl phosphatidylethanolamine as a substrate (42). The [¹⁴C]AA released was extracted using the method of Dole and Meinertz (40), and the associated radioactivity was counted.

Assessment of Phosphatidylserine (PS) Externalization-- Cells were magnetically labeled with annexin V conjugated with MACS colloidal super-paramagnetic MicroBeads, according to the instructions for the MACS apoptotic cell isolation kit, and a population of cells that expressed PS on the outer leaflets of their plasma membranes was collected by passing them through a separation column. Briefly, 5 × 10⁶ cells prelabeled with 0.1 µCi/ml [³H]AA were incubated with annexin V MicroBeads in a binding buffer containing Ca²⁺ for 15 min at 10 °C and washed with the binding buffer. The magnetically labeled apoptotic cells were applied to the MS+ separation column, which was placed in the magnetic field of a MiniMACS magnet separator, and the non-apoptotic cells were passed through the column. The column was rinsed with the binding buffer and removed from the magnet separator. The apoptotic cells, which were magnetically retained on the column, were eluted with the binding buffer, and their radioactivity was counted.

Measurement of Caspase-3 Activity-- Caspase-3 activity was assayed using the CPP32/Caspase-3 colorimetric protease assay kit, which is based on spectrophotometric detection of the chromophore p-nitroanilide that is produced after cleavage of the labeled substrate DEVD-p-nitroanilide. Briefly, cells were lysed with the lysis buffer and incubated on ice for 10 min. The lysates were centrifuged at 10,000 × g for 10 min at 4 °C, and the protein concentrations of the supernatants were determined using the BCA protein assay kit (Pierce). Then, 50 µg of protein equivalents were incubated with DEVD-p-nitroanilide for 2 h at 37 °C, and the absorbance of each solution at 405 nm was measured spectrophotometrically using a microtiter plate reader (Bio-Rad).

Immunoblotting-- Cells were washed with phosphate-buffered saline (PBS) and then lysed in PBS containing 100 µM p-4(2-aminoethyl)-benzenesulfonyl fluoride, 5 µM iodoacetamide, 5 mM EDTA, 1 µM pepstatin, 1 µg/ml soybean trypsin inhibitor, and 100 µM leupeptin by sonication for 1 min with a Branson Sonifier (power 30, 50% pulse cycle). The samples (10 µg of protein equivalents/lane) were subjected to 10% (w/v)SDS-polyacrylamide gel electrophoresis (PAGE) under reducing conditions and electroblotted onto nitrocellulose membranes (Schleicher & Schuell), which were probed with the antibodies and visualized with the ECL Western blot analysis system (Amersham Pharmacia Biotech), as described previously (13).

Plasmids-- A cPLA₂ deletion mutant, cPLA₂(1-522), was constructed by polymerase chain reaction (PCR) of the cPLA₂ coding sequence with ex Taq polymerase (Takara) using the oligonucleotide pair 5'-ATGTCATTTATAGATCCTTAC-3' and 5'-TCAGTCGAGCTCGTCATCGAA-3', as described previously (34). The PCR product was ligated into pCR^TM3.1 (Invitrogen) and was transfected into Top10F' supercompetent cells (Invitrogen). Colonies were picked up, and the plasmids were isolated and sequenced using a Taq cycle sequencing kit (Takara) and an auto-fluorometric DNA sequencer (DSQ-1000L, Shimadzu).

Construction of iPLA₂ mutants was carried out by PCR with KlenTaq polymerase (CLONTECH) using hamster iPLA₂ cDNA (Dr. S. Jones, Genetics Institute) as a template. An iPLA₂ deletion mutant, iPLA₂(184-C), was generated using the oligonucleotide pair 5'-ATG TAT CCG TAT GAT GTT CCT GAT TAT GCT AGC CTC AAC AAA GGA GAG ACG G-3' (HA epitope underlined) and 5'-TCA CTT GTC ATC GTC GTC CTT GTA GTC TGA TGA GGG CGA CAG CAG C-3' (FLAG epitope underlined, which was attached as required for the experiments). HA-tagged iPLA₂ was constructed using the primers 5'-ATG TAT CCG TAT GAT GTT CCT GAT TAT GCT AGC CTC ATG CAG TTC TTC GGA C-3' (HA epitope underlined) and 5'-TCA GGG CGA CAG CAG CAT TTG-3'. FLAG-tagged iPLA₂ was constructed using the primers 5'-ATG CAG TTC TTC GGA CGC C-3' and 5'-TCA CTT GTC ATC GTC GTC CTT GTA GTC TGA TGA GGG CGA CAG CAG C-3' (FLAG epitope underlined). PCR conditions were 94 °C for 30 s, 55 °C for 30 s, and 72 °C for 1 min, for 25 cycles. The products of the expected size were subcloned into the pCR3.1 vector and sequenced as noted above.

Transfection Studies-- HEK293 cells stably expressing each PLA₂ were established as described previously (16, 17). To obtain cells expressing cPLA₂(1-522) and epitope-tagged or truncated iPLA₂, their cDNAs subcloned into pCR3.1 were transfected into 293 cells using LipofectAMINE Plus (Life Technologies, Inc.) according to the manufacturer's instruction. Briefly, 1 µg of plasmid was mixed with 5 µl of LipofectAMINE Plus in 200 µl of Opti-MEM medium, left for 15 min, and then added to cells that had attained 40-60% confluency in 6-well plates (Iwaki) in 1 ml of Opti-MEM. After incubation for 6 h, the medium was replaced with 2 ml of fresh culture medium (RPMI 1640 containing 10% FCS). After overnight culture, the medium was replaced again with 2 ml of fresh medium, and culture was continued at 37 °C in a CO₂ incubator flushed with 5% CO₂ in humidified air. For transient expression analyses, the cells were harvested 3 days after transfection and used immediately. In order to establish stable transfectants, the cells were cloned by limiting dilution in 96-well plates in culture medium supplemented with 1 mg/ml G418 (Life Technologies, Inc.). After culture for 2-4 weeks, wells containing a single colony were chosen, and the expression was assessed by RNA blotting as well as immunoblotting using appropriate probes or antibodies. These established clones were expanded and used in the experiments described below.

To establish double transformants, 293 cells stably expressing cPLA₂(1-522) were subjected to a second transfection with cPLA₂, sPLA₂-IIA, or iPLA₂ cDNA, which had been subcloned into pcDNA3.1/Zeo(+) (Invitrogen). Three days after transfection, the cells were seeded into 96-well plates in the presence of 50 µg/ml zeocin (Invitrogen) in order to establish stable transformants expressing both cPLA₂(1-522) and either PLA₂ isozyme.

RNA Blotting-- Approximately equal amounts (~10 µg) of total RNA obtained from the transfected cells were applied to individual lanes of 1.2% (w/v) formaldehyde-agarose gels, electrophoresed, and transferred to Immobilon-N membranes (Millipore). The resulting blots were then probed with the relevant cDNA probes, which had been labeled with [³²P]dCTP (Amersham Pharmacia Biotech) by random priming (Takara Shuzo). All hybridizations were carried out as described previously (15).

Activation of HEK293 Cells-- HEK293 transfectants (5 × 10⁴ cells in 1 ml of culture medium) were seeded into 24-well plates. In order to assess AA release, 0.1 µCi/ml [³H]AA was added to the cells on day 3, when they had nearly reached confluent, and culture was continued for another day. After two washes with fresh medium, 250 µl of RPMI 1640 with or without 10 µM A23187, 1 ng/ml IL-1, and/or 10% FCS was added to each well, and the amount of free [³H]AA released into the supernatant during culture (30 min with A23187 and up to 8 h with IL-1) was measured. The percentage release of AA was calculated using the formula (S/(S + P)) × 100, where S and P are the radioactivities measured in equal portions of the supernatant and cell pellet, respectively. All of these procedures are described previously in detail (16-19).

Induction of Apoptosis in 293 Cells-- HEK293 transfectants (5 × 10⁴ cells in 1 ml of culture medium) were seeded into 24-well plates. 0.1 µCi/ml [³H]AA was added to the cells on day 2, when they are 50% confluent, and culture was continued for another day. After two washes with fresh medium without FCS, 250 µl of RPMI 1640 containing 1% FCS with or without CHX or etoposide was added to each well, and the amount of radioactivity released into the supernatant during culture was measured. MAFP or BEL was added to cells during treatment with CHX or etoposide as required for the experiments. To assess oleic acid (OA) release, 0.5 µCi/ml [³H]OA (NEN Life Science Products) was added to the cells instead of [³H]AA.

In Vitro Transcription and Translation-- [³⁵S]Methionine-labeled cPLA₂ and its truncated mutant cPLA₂(1-522) were synthesized using a PROTEINscript^TM kit (Ambion). Briefly, plasmids containing mouse cPLA₂ or cPLA₂(1-522) cDNA were transcribed using RNA polymerase and then incubated with [³⁵S]methionine (NEN Life Science Products) and rabbit reticulocyte lysate. The products were subjected to SDS-PAGE and visualized autoradiographically. The procedure was described in our previous report (34).

Confocal Microscopic Analysis-- cDNAs for native and truncated forms of cPLA₂ and those for HA-tagged native and truncated forms of iPLA₂ were subcloned into the pEGFP-C3 and -C1 vectors (CLONTECH), respectively, at the EcoRI site. These plasmids were transfected into 293 cells seeded onto collagen-coated coverglasses (Iwaki Glass) using LipofectAMINE 2000 (Life Technologies, Inc.). After culture for 3 days, the cells were fixed with 2% (w/v) paraformaldehyde in PBS for 30 min at room temperature. The coverslips were mounted on glass slides using Perma Fluor (Japan Tanner) and examined using a FLUOVIEW laser fluorescence microscope (Olympus).

Statistics-- All values shown are means ± S.E. for three to six separate experiments. Differences between means were determined by Student's t test, and those at p < 0.05 were considered significant.

	RESULTS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Caspase-cleaved cPLA₂ Suppresses the Functions of Native cPLA₂-- We have previously reported that cPLA₂ is cleaved at Asp⁵²² by caspase-3 in U937 cells undergoing Fas-mediated apoptosis (34). This proteolytic process destroys the catalytic dyad (Ser²²⁸ and Asp⁵⁴⁹) essential for cPLA₂ activity, thereby leading to its enzymatic inactivation. Similar cPLA₂ degradation was observed when U937 (see below), HeLa (35), or L929 cells (data not shown) were killed with TNF in combination with CHX. Thus, it is likely that the caspase-directed cleavage of cPLA₂ is an event generally occurring in the apoptotic process.

In order to investigate whether the cleaved cPLA₂ plays some roles in cellular AA metabolism, death, survival, or proliferation, we prepared cDNA for the mutant cPLA₂ truncated at Asp⁵²² (cPLA₂(1-522)) and transfected it into HEK293 cells. When a sonicate of HEK293 cells expressing native cPLA₂ was centrifuged at 100,000 × g and then subjected to immunoblotting using anti-cPLA₂ antibody, most of the enzyme, as expected, was found to be recovered mainly in the supernatant cytosolic fraction (Fig. 1A). On the other hand, overexpressed cPLA₂(1-522) was distributed mainly in the membrane fraction (Fig. 1A), even though the lysate was prepared in the presence of EDTA, which chelates Ca²⁺ that is thought to be essential for the translocation of native cPLA₂ to the phospholipid membrane (5, 12). To assess whether cPLA₂(1-522) endogenously generated during apoptosis also has altered subcellular distribution, lysates of U937 cells before and 24 h after treatment with agonistic anti-Fas antibody were separated by centrifugation at 100,000 × g into supernatant and pellet and then subjected to immunoblotting. Under the experimental conditions employed, endogenous native cPLA₂ was distributed evenly in supernatant and membrane fractions, whereas the 78-kDa fragment was exclusively detected in the membrane fraction (Fig. 1B). Furthermore, when [³⁵S]methionine-labeled cPLA₂ and cPLA₂(1-522), which were generated by in vitro transcription/translation, were each mixed with the U937 cell lysate and then centrifuged, the former was detected predominantly in the supernatant with a minor portion being associated with the membrane, whereas the latter was again recovered exclusively in the membrane fraction (Fig. 1C). Thus, the removal of C-terminal one-third of cPLA₂ results in enhanced affinity for membranes.

View larger version (38K): [in this window] [in a new window]   Fig. 1.   cPLA2(1-522) is constitutively associated with cellular membranes. A, expression of cPLA2 and its truncated mutant, cPLA2(1-522), in HEK293 cells. Cell lysates (L) were centrifuged for 1 h at 100,000 × g to separate into the supernatant cytosolic (C) and pelleted membrane (M) fractions and were subjected to SDS-PAGE followed by immunoblotting using anti-cPLA2 antibody. B, lysates of U937 cells before () and after (+) treatment with anti-Fas antibody (Ab) for 12 h were centrifuged, and the resulting cytosolic and membrane fractions were subjected to immunoblotting using the same antibody. C, [35S]methionine-labeled cPLA2 and cPLA2(1-522), which had been prepared by in vitro transcription and translation, were incubated with U937 lysates (107 cells/ml equivalent) for 1 h, centrifuged, separated on SDS-PAGE, and visualized by autoradiography. WT, wild type.

We reasoned that if cPLA₂(1-522) is constitutively associated with the membrane site to which native cPLA₂ translocates from the cytosol only after an increase in cytoplasmic Ca²⁺ levels, this fragment would compete with the native enzyme for the site and eventually affect the function of the native enzyme in regulating stimulus-induced AA release. To explore this, we established double transfectants expressing both native cPLA₂ and cPLA₂(1-522); expression levels in these cells were verified by immunoblotting using anti-cPLA₂ antibody (Fig. 2A). When the cells expressing native cPLA₂ alone were stimulated with 10 µM A23187 for 30 min (immediate response) (Fig. 2B) or 1 ng/ml IL-1 for 4 h (delayed response) (Fig. 2C), there was a marked increase in [³H]AA release relative to control cells, which expressed endogenous cPLA₂ minimally and did not exhibit increased [³H]AA release after stimulation, as we have reported previously (16, 17). cPLA₂(1-522) failed to increase [³H]AA release (Fig. 2, B and C), in agreement with the fact that it lacks catalytic activity (34). Notably, the AA-releasing function of native cPLA₂, in both the A23187- and IL-1-dependent responses, was attenuated markedly when cPLA₂(1-522) was coexpressed (Fig. 2, B and C). This result raises the intriguing possibility that cPLA₂(1-522) behaves like a dominant-negative molecule, preventing the signaling function of the native form of cPLA₂.

View larger version (27K): [in this window] [in a new window]   Fig. 2.   cPLA2(1-522) inhibits the AA-releasing function of cPLA2. A, expression of wild-type (WT) cPLA2 and cPLA2(1-522) proteins in HEK293 transfectants as assessed by immunoblotting. B-D, [3H]AA-prelabeled cells were incubated for 30 min with 10 µM A23187 in the presence of 1% FCS (B) for 4 h with 1 ng/ml IL-1 in the presence of 10% FCS (C) and for 12 h with 10 µg/ml CHX in the presence of 1% FCS (D). The percentage release of [3H]AA was measured. Values are the means ± S.E. for more than three independent experiments (*p < 0.05 versus control).

Treatment of subconfluent HEK293 cells with CHX or etoposide, chemicals that induce apoptosis through the mitochondria-dependent pathway (43), alone caused significant cell death, reducing their viability to approximately 80 and 50% after 8 and 24 h of culture, respectively, in medium containing 1% FCS. This was accompanied by conversion of the overexpressed cPLA₂ to a 78-kDa fragment (see below), indicating endogenous caspase-3 activation during CHX- or etoposide-induced cell death. Cells transfected with either native cPLA₂ or cPLA₂(1-522) grew normally (see below), and their sensitivity to CHX- or etoposide-induced cell death was comparable to that of control cells (data not shown). Release of [³H]AA by transfectants expressing native cPLA₂ or cPLA₂(1-522) alone or in combination after CHX treatment did not significantly differ from that by control cells (Fig. 2D). Thus, neither native cPLA₂ nor cPLA₂(1-522) affected growth and death, as well as death-associated AA release, at least under the conditions employed here.

Requirement for cPLA₂ for the Function of Other PLA₂s-- The observation that cPLA₂(1-522) behaves like a dominant-negative inhibitor of cPLA₂-dependent AA release prompted us to utilize the cPLA₂(1-522) cotransfection system to assess the requirement for cPLA₂ for the function of other PLA₂ enzymes. Thus, we introduced cPLA₂(1-522) into 293 cells expressing sPLA₂-IIA, sPLA₂-X, or iPLA₂, which we had established previously (16-19), to investigate the effect on their AA-releasing functions. The expression of each PLA₂ and cPLA₂(1-522) in the established clones was confirmed by RNA blotting (Figs. 3, A and C, and 4A). As reported previously (16-19), sPLA₂-IIA increased [³H]AA release only after stimulation with A23187 (data not shown) or IL-1 (Fig. 3B). This signaling function of sPLA₂-IIA was abrogated almost completely by coexpression of cPLA₂(1-522) (Fig. 3B). sPLA₂-X is a unique sPLA₂ isozyme in terms of its ability to avidly hydrolyze phosphatidylcholine in the plasma membrane, causing spontaneous [³H]AA release during cell culture in medium containing FCS (19, 30). As shown in Fig. 3D, cPLA₂(1-522) coexpression did not affect this spontaneous [³H]AA release by sPLA₂-X. Therefore, the suppressive effect of cPLA₂(1-522) is limited to particular classes of signaling PLA₂ and is not a reflection of a nonspecific action. Thus, these results provide strong support for the hypotheses that cPLA₂ is a prerequisite for signaling sPLA₂s to function properly (13, 44, 45) and that sPLA₂-IIA and sPLA₂-X act on cells through different mechanisms (19).

View larger version (43K): [in this window] [in a new window]   Fig. 3.   cPLA2(1-522) inhibits sPLA2-IIA-, but not sPLA2-X-, mediated AA release. A and C, expression of transcripts for sPLA2-IIA (A) and sPLA2-X (C) with or without cPLA2(1-522) coexpression in HEK293 transfectants. B and D, [3H]AA release by cells transfected with sPLA2-IIA (B) and sPLA2-X (D) with or without cPLA2(1-522) coexpression. Cells were incubated for 4 h in medium containing 10% FCS in the presence (B) or absence (D) of 1 ng/ml IL-1. Values are the means ± S.E. for three independent experiments (*p < 0.05 versus control).

Overexpression of iPLA₂ in 293 cells led to increased FCS-dependent spontaneous [³H]AA release over the culture period (Fig. 4B, left) (16), the event likely to reflect its phospholipid remodeling function (21). Cotransfection of cPLA₂(1-522) did not significantly affect this remodeling activity of iPLA₂ (Fig. 4B, left), indicating that cPLA₂ and iPLA₂ are functionally segregated and that cPLA₂ does not play a role in phospholipid remodeling reactions. On the other hand, several studies have argued that iPLA₂ has the capacity to promote stimulus-induced AA release (22, 46-48). Indeed, when iPLA₂ transfectants were stimulated with A23187, immediate [³H]AA release was increased markedly (Fig. 4B, right). Surprisingly, this stimulus-induced [³H]AA release from iPLA₂-expressing cells was significantly reduced by cotransfection with cPLA₂(1-522) (Fig. 4B, right). Moreover, whereas the FCS-dependent phospholipid remodeling function of iPLA₂ did not show appreciable fatty acid selectivity (Fig. 4C, left) (16), A23187 stimulation of iPLA₂ transfectants caused release of [³H]AA in marked preference to [³H]OA (Fig. 4C, right). These results imply the existence of functional cooperation between cPLA₂ and iPLA₂ in the Ca²⁺-dependent cellular response.

View larger version (25K): [in this window] [in a new window]   Fig. 4.   Dual effects of cPLA2(1-522) on iPLA2-mediated AA release. A, expression of transcripts for iPLA2 and cPLA2(1-522) in HEK293 transfectants. B, [3H]AA release by cells expressing iPLA2 with or without cPLA2(1-522) coexpression after 4 h of culture with 10% FCS (left) or 30 min of stimulation with 10 µM A23187 (right). C, fatty acid selectivity of iPLA2. Cells prelabeled with [3H]AA or [3H]OA, with (+) or without () iPLA2 transfection, were cultured for 4 h with 10% FCS (left) or stimulated for 30 min with A23187 (right). Values are the means ± S.E. for three independent experiments (*p < 0.05 versus control).

iPLA₂ Is Cleaved by Caspase and Potentiates Cell Death-associated AA Release-- Treatment of U937 cells with anti-Fas antibody (34) or TNF/CHX (Fig. 5A) was accompanied by a time-dependent increase in [³H]AA release, which paralleled accumulation of apoptotic cells (see Fig. 8). Immunoblotting confirmed that cPLA₂, with an apparent molecular mass of 110 kDa, was entirely converted to a 78-kDa fragment in cells treated for 12 h with TNF plus CHX, as compared with those treated with TNF or CHX alone, in which cPLA₂ remained uncleaved (Fig. 5B). Thus, it is unlikely that the increased [³H]AA release in TNF/CHX-treated cells resulted from cPLA₂ activation. This increased [³H]AA release was markedly suppressed by the iPLA₂ inhibitors MAFP and BEL (Fig. 5C) and did not show appreciable fatty acid selectivity (data not shown) (34). Furthermore, the increased [³H]AA levels were accompanied by a reciprocal decrease in the radioactivity associated with the phospholipid fraction (34). These results suggest that the PLA₂ isozyme responsible for increased fatty acid release during apoptosis may be iPLA₂.

View larger version (16K): [in this window] [in a new window]   Fig. 5.   [3H]AA release from TNF/CHX-treated U937 cells is sensitive to iPLA2 inhibitors. A, time course of [3H]AA release from U937 cells cultured for the indicated periods with 100 units/ml TNF (solid circles), 10 µg/ml CHX (open squares), their combination (solid squares), or in their absence (open circles). Values are the means ± S.E. for more than three independent experiments (*, p < 0.05 versus control). B, proteolysis of cPLA2. Lysates of U937 cells treated for 12 h with various combination of TNF and CHX were subjected to immunoblotting using anti-cPLA2 antibody. C, [3H]AA-prelabeled U937 cells were cultured for 24 h with 100 units/ml TNF and 10 µg/ml CHX in the presence or absence of 10 µM MAFP or BEL. Values are the means ± S.E. for more than three independent experiments (*, p < 0.05 versus control; **, p < 0.05 versus cells treated with TNF/CHX without the inhibitors).

Immunoblot analysis of U937 cells with anti-iPLA₂ antibody revealed that, in addition to an intact 85-kDa iPLA₂ protein, another immunoreactive band with an estimated molecular mass of 70 kDa became visible 6-12 h after treatment with TNF/CHX (Fig. 6A), the time period during which TNF/CHX-mediated AA release (Fig. 5A) and caspase-3 activity (see Fig. 8) increased significantly. Similar results were also observed in TNF/CHX-treated L929 cells (data not shown). We noted that a potential caspase cleavage site, DXXDX (49), is present in iPLA₂ around Asp¹⁸³, which is located near the C-terminal end of the first ankyrin repeat (Fig. 6B). If iPLA₂ is cleaved at this site (DVTD¹⁸³Y), the predicted size of the resulting C-terminal fragment would be consistent with the size of the cleaved fragment observed in this study.

View larger version (29K): [in this window] [in a new window]   Fig. 6.   iPLA2 is cleaved by caspase-3. A, immunoblotting of U937 cells, which were cultured for the indicated periods with or without 100 units/ml TNF and 10 µg/ml CHX, using anti-iPLA2 antibody. B, the sequence DVTDY within the first ankyrin repeat of iPLA2 is a potential cleavage site for caspase-3. C, HEK293 cells transfected with iPLA2-FLAG or HA-iPLA2(184-C) (left) and cPLA2 wild-type (WT) or cPLA2(1-522) (right) were cultured for 24 h with or without 10 µg/ml CHX or etoposide and then taken for immunoblotting using the respective antibodies. D, 293 cells expressing iPLA2-FLAG were secondary transfected with caspase-3, cultured for 48 h, and then subjected to immunoblotting using anti-FLAG antibody.

To verify whether iPLA₂ is a substrate for caspase-3 and whether the resultant truncated iPLA₂ retains its activity to facilitate cell death-associated fatty acid release, we constructed an iPLA₂ mutant, iPLA₂(184-C), that would correspond to the truncated enzyme after cleavage at Asp¹⁸³ and transfected it into HEK293 cells. The full-length iPLA₂ and iPLA₂(184-C) constructs were tagged with FLAG or HA epitope, so that their expression could readily be visualized by immunoblotting using the respective anti-tag antibodies. The C-terminally FLAG-tagged iPLA₂ was expressed as the expected 85-kDa protein in HEK293 transfectants (Fig. 6C), but not in control cells (data not shown), when immunoblotted with anti-FLAG antibody. After culture of iPLA₂-FLAG-expressing 293 cells with CHX or etoposide for 24 h, there was a significant increase in a 70-kDa FLAG-immunoreactive protein band, which comigrated with N-terminally HA-tagged iPLA₂(184-C) that was visualized with anti-HA antibody (Fig. 6C, left). Under the same conditions, overexpressed cPLA₂ was almost completely cleaved to cPLA₂(1-522) in CHX- or etoposide-treated cells (Fig. 6C, right). Furthermore, transient transfection of caspase-3 into cells expressing iPLA₂-FLAG resulted in the significant appearance of a similar FLAG-immunoreactive 70-kDa protein (Fig. 6D), which was exactly the same size as the HA-tagged iPLA₂(184-C) (data not shown). These results suggest that iPLA₂ is indeed a substrate for caspase-3 and is cleaved at the consensus site Asp¹⁸³. However, only partial cleavage of endogenous (Fig. 6A) and overexpressed (Fig. 6C, left) iPLA₂ relative to cPLA₂, which was entirely converted to a truncated fragment (Figs. 5B and 6C, right), suggests that iPLA₂ is a poorer substrate for caspase-3 than cPLA₂ or that other caspase(s) is responsible for iPLA₂ cleavage.

We next investigated the functions of iPLA₂ and its truncated 70-kDa fragment using HEK293 transfectants. The expression levels of HA-tagged iPLA₂(184-C) and HA-tagged full-length iPLA₂ in the respective transfectants were comparable (Fig. 7A). When these cells were prelabeled with [³H]AA, washed, and then cultured in the presence of 10% FCS, HA-iPLA₂ transfectants released significantly more [³H]AA than parental cells (Fig. 7B), a result consistent with our previous report (16). Conjugating the N terminus of iPLA₂ with the HA epitope did not alter its fatty acid-releasing function at the cellular level, as spontaneous [³H]AA release by HA-iPLA₂ was similar to that by iPLA₂ without the tag (data not shown). Furthermore, HA-iPLA₂ transfectants released significantly more [³H]AA and [³H]OA in parallel than control cells over 24 h of culture with etoposide (Fig. 7C) or CHX (Fig. 7D), which induced cell death. Notably, we found that the fatty acid-releasing activity of HA-iPLA₂(184-C) was significantly higher than that of HA-iPLA₂. Thus, cells expressing HA-iPLA₂(184-C) released more [³H]AA than those expressing HA-iPLA₂ during culture (Fig. 7B); FCS-independent [³H]AA release by HA-iPLA₂(184-C) cells reached levels comparable to FCS-dependent release by replicate HA-iPLA₂-expressing cells. Moreover, cells expressing HA-iPLA₂(184-C) released more [³H]AA and [³H]OA than those expressing HA-iPLA₂ 6-24 h after treatment with etoposide (Fig. 7C) or CHX (Fig. 7D), a time that correlated with cell death. Release of both [³H]AA (Fig. 7E) and [³H]OA (data not shown) from cells expressing HA-iPLA₂(184-C) or HA-iPLA₂ after CHX treatment was suppressed markedly by MAFP or BEL in a dose-dependent manner. TLC analysis showed that more than 75% of the radioactivity released into the supernatant from the CHX-treated HA-iPLA₂(184-C) transfectants was associated with the free fatty acid fraction. Taken together, these results provide unequivocal evidence that iPLA₂ promotes cell death-associated fatty acid release.

View larger version (31K): [in this window] [in a new window]   Fig. 7.   Effects of native and truncated iPLA2 on fatty acid release. A, expression of iPLA2 and iPLA2(184-C) in HEK293 transfectants. B, [3H]AA release from the cells shown in A after culture for 24 h with or without 10% FCS. C, [3H]AA and [3H]OA release from the cells after culture for 24 h with or without 10 µg/ml etoposide in medium containing 1% FCS. D, time course of [3H]AA release from the cells cultured for the indicated periods with or without 10 µg/ml CHX. E, cells were cultured for 24 h with 10 µg/ml CHX in the presence of the indicated concentrations of MAFP or BEL. Values are the means ± S.E. for more than three independent experiments (*, p < 0.05 versus control; **, p < 0.05 versus HA-iPLA2-expressing cells).

iPLA₂ Affects Cell Death and Growth-- The appearance of trypan blue-positive U937 cells after treatment with anti-Fas antibody or TNF/CHX was retarded markedly by the addition of MAFP, the inhibitory effect of which was observed during culture for 3-6 h, but was not evident after 12 h (Fig. 8A). PS externalization, a hallmark of apoptosis (50), was also suppressed significantly by MAFP (Fig. 8B); the dose dependence of this effect was parallel to that of the inhibition of cell death and accumulation of released [³H]AA (data not shown). Similar effects were observed when BEL was added instead of MAFP (data not shown). In contrast, caspase-3 activation was not affected by MAFP (Fig. 8C), consistent with the results shown above that iPLA₂ activation by cleavage at Asp¹⁸³ occurs downstream of caspase-3. DNA fragmentation and cPLA₂ cleavage, both of which depend on caspase-3 (34, 51), were unaffected by MAFP (data not shown).

View larger version (31K): [in this window] [in a new window]   Fig. 8.   iPLA2 affects certain cellular apoptotic or growth processes. A-C, effects of MAFP on apoptotic processes in U937 cells undergoing anti-Fas antibody- or TNF/CHX-induced apoptosis. Cell viability was assessed by the trypan blue dye exclusion test (A), PS externalization was assessed using annexin V MicroBeads (B), and caspase-3 activity was assessed using a colorimetric protease assay kit (C). The viability (A) and caspase-3 activity (C) of cells that were cultured for the indicated periods with 50 ng/ml anti-Fas antibody (left) or 100 units/ml TNF plus 10 µg/ml CHX (right) in the presence (shaded bars) or absence (solid bars) of 10 µM MAFP were measured. Caspase-3 activity was expressed as fold increase relative to cells before anti-Fas antibody or TNF/CHX treatment. PS externalization was determined after 3 h of culture with the anti-Fas antibody or TNF/CHX in the presence of the indicated concentrations of MAFP; the culture without MAFP treatment was taken as 100%. D, proliferation rates of HEK293 cells transfected with native or truncated iPLA2 or cPLA2. Cells seeded at 5 × 104 cells/ml were cultured for 4 days, and viable cells were counted. The results were expressed as relative values, with the growth of control cells considered as 100%. Values are the means ± S.E. for more than three independent experiments (*, p < 0.05 versus respective controls). E, cellular morphology of parent 293 cells and cells stably expressing iPLA2 or iPLA2(184-C).

In contrast to the 293 cells transfected with native cPLA₂ or cPLA₂(1-522), which exhibited normal cell growth, those transfected with HA-iPLA₂ grew significantly more slowly than parental cells (Fig. 8D). This growth retardation was further pronounced in cells expressing HA-iPLA₂(184-C) (Fig. 8D). As shown in Fig. 8E, cells transfected with HA-iPLA₂ or HA-iPLA₂(184-C) displayed aberrant morphology, being shrunken and aggregated, as compared with the control cells, which exhibited a typical fibroblastic shape. Collectively, these observations suggest that iPLA₂ can participate in the modulation of cell proliferation and the apoptotic process.

Subcellular Distribution-- cPLA₂, iPLA₂, and their truncated products were each expressed as an enhanced green fluorescent (EGFP) fusion protein in 293 cells, and their subcellular distributions were analyzed by confocal laser microscopy. As expected, native cPLA₂ was located in the cytosol (Fig. 9A) and was translocated into the perinuclear region after cell activation, as reported previously (18, 52). Notably, cPLA₂(1-522) produced strong peri- and intra-nuclear fluorescence with no cytoplasmic signal (Fig. 9A). EGFP alone was distributed in the cytosol, implying that nuclear localization of cPLA₂(1-522) was not due to the EGFP fusion construct. In contrast, both signals for native iPLA₂ and iPLA₂(184-C) were detected throughout the cytoplasm (Fig. 9B).

View larger version (50K): [in this window] [in a new window]   Fig. 9.   Subcellular distribution. Subcellular localizations of EGFP-fused cPLA2, cPLA2(1-522) (A), iPLA2, and iPLA2(184-C) (B), which were transiently transfected into 293 cells, were assessed by confocal microscopy (left). Phase-contrast photographs are also shown (right).

	DISCUSSION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

In the present study, we have shown that the two intracellular PLA₂s expressed ubiquitously in a wide variety of cells, cPLA₂ and iPLA₂, undergo caspase-directed cleavage at specific sites and may play distinctive roles during the process of apoptosis. The N-terminal cPLA₂ fragment, which is produced after cleavage at Asp⁵²², has no catalytic function, is constitutively associated with membranes, and suppresses stimulus-induced AA release by uncleaved cPLA₂ as well as by other signaling PLA₂ enzymes, most likely through its dominant-inhibitory action (Fig. 10A). This implies that, on the way to apoptotic death, cells lose the ability to produce eicosanoids in response to extracellular stimuli. On the other hand, the C-terminal iPLA₂ fragment, which is produced after cleavage at Asp¹⁸³, is functionally more active than intact iPLA₂ and accelerates cell death-associated fatty acid release, which may be linked to certain apoptotic membranous changes (Fig. 10B).

View larger version (29K): [in this window] [in a new window]   Fig. 10.   Schematic model for the roles of cPLA2 and iPLA2 in the apoptotic pathway. A, cPLA2 is cleaved by caspase-3 at Asp522 and is catalytically inactivated. The resultant N-terminal fragment, cPLA2(1-522), constitutively translocates to the nuclear membrane, where it may prevent the remaining uncleaved cPLA2 from interacting with the cPLA2-binding site in a Ca2+-dependent fashion, thereby inhibiting the AA-releasing function of cPLA2. As a result, apoptotic cells lose the ability to produce lipid mediators so as to die silently. The molecular mechanisms and physiological significance of the intranuclear transport of cPLA2(1-522) remains to be elucidated. B, iPLA2 is cleaved by caspase-3 at Asp183 and is further activated. Both intact and cleaved iPLA2 act on the membranes to accelerate membrane turnover, which modifies the process of apoptosis.

cPLA₂ Cleavage-- Several previous studies have suggested that cPLA₂ is involved in apoptosis or cytotoxicity (31-33). However, our present results suggest that cPLA₂ is unnecessary for the apoptotic pathway and that the involvement of cPLA₂ in apoptosis observed in these studies may reflect secondary or cell type-specific events. It is more likely that cPLA₂ participates indirectly in the cell death pathway only in particular cell types in which endogenous eicosanoids or related lipid mediators influence their survival or growth. Adam-Klages and coworkers (35, 36) have recently reached the same conclusion that cPLA₂ is proteolytically inactivated by caspase-3, and they additionally showed cleavage by caspase-1, which cleaves cPLA₂ at Asp⁴⁵⁹. The latter proteolysis was also observed in our experimental system at a relatively late phase of Fas-mediated apoptosis.²

An important finding is that the N-terminal fragment of cPLA₂, cPLA₂(1-522), acts as a dominant-negative inhibitor, preventing cPLA₂-mediated stimulus-induced AA release (Fig. 2) that is linked to downstream eicosanoid biosynthesis (16, 17). Because cPLA₂(1-522) is predominantly associated with membranes even in unstimulated cells (Fig. 1), the likely explanation for the dominant-negative effect is that this truncated form competes with native cPLA₂ for binding to a membrane site that is necessary for the activation process. Although the mechanism for constitutive association of cPLA₂(1-522) with the membrane is unclear, deletion of the C terminus may lead to exposure of a putative Ca²⁺-independent membrane binding domain, apart from the C2 domain, within the cPLA₂(1-522) fragment (53). Alternatively, cPLA₂(1-522) may compete for phosphorylation at Ser⁵⁰⁵ by mitogen-activated protein kinase, which is required for optimal activation of cPLA₂ (6).

In support of membrane distribution of cPLA₂(1-522) as assessed by subcellular fractionation (Fig. 1), immunofluorescent microscopic analysis showed that cPLA₂(1-522) is located in the nuclear, not in the cytosolic, compartment in HEK293 transfectants (Fig. 9A). Intranuclear location of cPLA₂(1-522) is noteworthy, because there is a putative bipartite nuclear localization sequence (residues 273-284, KRYVESSLWKKK) within cPLA₂(1-522). In addition, cPLA₂ has a potential nuclear export motif (residues 556-564, LPYPLILRP), which would be eliminated after caspase-3-directed cleavage at Asp⁵²². Thus, exposure of the nuclear targeted sequence after the cleavage may contribute to intranuclear transport of cPLA₂(1-522), and this possibility is now under investigation. Furthermore, intranuclear function of cPLA₂(1-522) during the apoptotic process needs to be evaluated in the future.

cPLA₂(1-522) also markedly reduced stimulus-induced AA release by sPLA₂-IIA (Fig. 3B), a signaling sPLA₂ isozyme that plays an augmentative role in eicosanoid production in autocrine, paracrine, and juxtacrine fashions (13, 15-19, 55). This suppressive effect of cPLA₂(1-522) is rather selective, because spontaneous AA release by sPLA₂-X (Fig. 3D), which has been shown to act on cells in a manner different from sPLA₂-IIA (19, 20), and iPLA₂ (Fig. 4B), which has been implicated in phospholipid remodeling (21), was unaffected by cPLA₂(1-522). Thus, our present results give additional support to the hypothesis that cPLA₂ is required for signaling sPLA₂s to act properly (13, 44, 45). According to the proposed model for this, certain cPLA₂ reaction products (fatty acid or lysophospholipid derivatives) may perturb membrane microdomains in which signaling sPLA₂s are functioning.

Although several studies (46-48) have argued for a possible contribution of iPLA₂ to stimulus-induced AA release, this interpretation has been suggested by others to need reevaluation (21), because most of the results were based solely on the pharmacological inhibitory effect of BEL. Akiba et al. (22) have recently reported that antisense oligonucleotide for iPLA₂ suppressed zymosan-induced AA release in macrophage-like cells, even though there is general recognition that cPLA₂ is an absolute requirement for this process (56). These discrepancies may be reconciled by our present observation that A23187-induced AA release from iPLA₂-expressing cells was markedly reduced by cPLA₂(1-522) (Fig. 4B), which suggests that endogenous cPLA₂ and overexpressed iPLA₂ functionally cooperated in this setting. As fatty acid release from iPLA₂-expressing cells after A23187 stimulation was relatively AA-selective (Fig. 4C), we presume that iPLA₂ may potentiate the function of endogenous cPLA₂ by modifying substrate susceptibility, or that iPLA₂, in conjunction with cPLA₂, acts on AA-rich phospholipid pools in microdomains, during intracellular Ca²⁺ signaling.

iPLA₂ Cleavage-- We previously reported that the increased fatty acid release observed in U937 cells undergoing Fas-mediated apoptosis was sensitive to general iPLA₂ inhibitors (34); however, studies using inhibitors alone might be insufficient to define the contribution of iPLA₂ to this event. Here we have substantiated the involvement of iPLA₂ in cell death-associated fatty acid release. We have shown that iPLA₂ is a substrate for caspase-3 (or other caspases), being cleaved during the apoptotic process at the consensus Asp¹⁸³ within the first ankyrin repeat (Fig. 6). This cleavage led to production of a truncated enzyme devoid of most of the first ankyrin repeat. Importantly, the produced fragment, iPLA₂(184-C), which possesses the entire catalytic domain and seven of eight ankyrin repeats, is functionally more active than intact iPLA₂ in cells (Fig. 7). Since the activities of the intact and truncated forms of iPLA₂ were similar when measured in the in vitro PLA₂ assay² and the intracellular location of iPLA₂ before and after cleavage was similar (Fig. 9B), marked differences in their fatty acid-releasing capacities in vivo may indicate the existence of a putative intracellular iPLA₂-regulatory factor, which binds and negatively regulates iPLA₂. Removal of the first ankyrin repeat may release iPLA₂ from the inhibitory factor, leading to increased accessibility to endogenous substrates. Conversely, it is also possible that the truncated form of iPLA₂ might interact with the putative iPLA₂ activator more efficiently than the intact enzyme. These speculations are consistent with the suggestion that iPLA₂ forms a high molecular weight complex, probably through the ankyrin repeat motif, in cells (8, 57, 58).

Our data suggest that, at least in certain cell types, iPLA₂, and iPLA₂(184-C) even more, plays a modifying role in the cell death pathway (Fig. 8). iPLA₂ appears to have a significant effect on transbilayer movement of PS, an apoptotic membranous event, at the early stage of cell death. The delay in appearance of dead cells following inhibition of iPLA₂ suggests that perturbed membrane turnover significantly affects some apoptotic changes, although it depends on cell types. Furthermore, the apparently slow growth and abnormal morphology of iPLA₂-transfected 293 cells in comparison with control cells implies that unbalanced membrane phospholipid homeostasis often affects the cell proliferation machinery. Consistent with this, overloading of cell with non-hydrolyzable alkyl-phospholipid analogs disturbs the balance between iPLA₂-mediated deacylation and subsequent reacylation and inhibits cell proliferation (54). Thus, caspase-directed cleavage of iPLA₂ leads to accelerated phospholipid remodeling, which perturbs the structure, dynamics, integrity, or asymmetry of bilayer membranes, thereby influencing growth and/or the apoptotic state of the cell.

	ACKNOWLEDGEMENTS

We thank Drs. S. Jones and K. Takahashi for providing hamster iPLA₂ and human caspase-3 cDNAs, respectively.

	FOOTNOTES

^* This work was supported by Grants-in-aid for Scientific Research from the Ministry of Education, Science and Culture of Japan and Special Coordination Funds for Promoting Science and Technology from the Science and Technology Agency.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Both authors contributed equally to this work.

^§ To whom correspondence should be addressed. Tel.: 81-3-3784-8196; Fax: 81-3-3784-8245; E-mail: kudo@pharm.showa-u.ac.jp.

Published, JBC Papers in Press, March 19, 2000, DOI 10.1074/jbc.M000271200

² G. Atsumi, M. Murakami, K. Kojima, and I. Kudo, unpublished observations.

	ABBREVIATIONS

The abbreviations used are: PLA₂, phospholipase A₂; cPLA₂, cytosolic phospholipase A₂; sPLA₂, secretory PLA₂; iPLA₂, Ca²⁺-independent PLA₂; TNF, tumor necrosis factor ; IL-1, interleukin-1; AA, arachidonic acid; OA, oleic acid; PBS, phosphate-buffered saline; MAFP, methylarachidonyl fluorophosphate; BEL, bromoenol lactone; CHX, cycloheximide; HEK, human embryonic kidney; FCS, fetal calf serum; PS, phosphatidylserine; PCR, polymerase chain reaction; PAGE, polyacrylamide gel electrophoresis; EGFP, enhanced green fluorescent protein; HA, hemagglutinin.

	REFERENCES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES