Am J Physiol Cell Physiol Journal of Applied Physiology
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Am J Physiol Cell Physiol 293: C1605-C1615, 2007. First published September 5, 2007; doi:10.1152/ajpcell.00012.2007
0363-6143/07 $8.00
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
293/5/C1605    most recent
00012.2007v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Poulsen, K. A.
Right arrow Articles by Lambert, I. H.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Poulsen, K. A.
Right arrow Articles by Lambert, I. H.

MUSCLE CELL BIOLOGY AND CELL MOTILITY

Induction of group VIA phospholipase A2 activity during in vitro ischemia in C2C12 myotubes is associated with changes in the level of its splice variants

K. A. Poulsen,1 S. F. Pedersen,1 M. Kolko,2 and I. H. Lambert1

1Department of Molecular Biology, University of Copenhagen, and 2Eye Pathology Institute, Copenhagen, Denmark

Submitted 10 January 2007 ; accepted in final form 2 September 2007


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
The involvement of group VI Ca2+-independent PLA2s (iPLA2-VI) in in vitro ischemia [oxygen and glucose deprivation (OGD)] in mouse C2C12 myotubes was investigated. OGD induced a time-dependent (0–6 h) increase in bromoenol lactone (BEL)-sensitive iPLA2 activity, which was suppressed by specific short interfering (si)RNA knockdown of iPLA2-VIA. OGD was associated with an increase in iPLA2-VIA protein levels, whereas mRNA levels were unchanged. The levels of iPLA2-VIB mRNA and protein were not increased by OGD. RT-PCR and Western blot analysis identified a mouse iPLA2-VIA homolog to catalytically inactive 50-kDa iPLA2-VIA-ankyrin variants previously identified in humans. Both the mRNA and protein levels of this ~50-kDa variant were reduced significantly within 1 h following OGD. In C2C12 myoblasts, iPLA2-VIA seemed to predominantly reside at the endoplasmatic reticulum, where it accumulated further during OGD. A time-dependent reduction in cell viability during the early OGD period (3 h) was partially prevented by iPLA2-VIA knockdown or pharmacological inhibition (10 µM BEL), whereas iPLA2-VIA overexpression had no effect on cell viability. Taken together, these data demonstrate that OGD in C2C12 myotubes is associated with an increase in iPLA2-VIA activity that decreases cell viability. iPLA2-VIA activation may be modulated by changes in the levels of active and inactive iPLA2-VIA isoforms.

hypoxia; Ca2+-independent phospholipase A2-β; Ca2+-independent phospholipase A2-{gamma}

ISCHEMIA is a common and clinically important cause of injury in many tissues. In skeletal muscle, ischemia/hypoxia arises during circulatory and respiratory insufficiency and during surgery and trauma and is associated with a pronounced loss of function (14).

Alterations in cell membrane structure and organization during ischemia are considered as important factors in determining the degree of irreversible damage (22). Cell membrane composition and organization are partially regulated by a deacylation-reacylation cycle during which the sn-2 ester bonds of glycerophospholipids are hydrolyzed by PLA2 followed by immediate reincorporation of new fatty acids by CoA-dependent acyltransferases maintaining a low cellular concentration of free fatty acids (FFA) and lysophospholipids (6). This balance may become disturbed as a result of increased PLA2 activity (20), resulting in elevated levels of FFAs [e.g., arachidonic acid (AA)] and lysophopholipids [e.g., lysophosphatidylcholine (LPC)] (17, 41, 47, 50). AA mediates a variety of pathophysiological responses either directly or indirectly when metabolized by cycloxygenases, lipoxygenases, or cytochrome P-450 (13), whereas LPC is toxic in high concentrations due to its detergent-like properties, i.e., LPC increases membrane fluidity and permeability (20). On the other hand, the activity of certain PLA2 subtypes may have membrane repair functions and potentially provide protection during cellular stress (17, 41). Thus, both protective and detrimental effects of PLA2 activity in ischemia may be envisaged.

PLA2s are divided into four major families based on their nucleotide sequence: secretory low-molecular-weight PLA2s (sPLA2s), which require millimolar concentrations of Ca2+ for catalytic activity; intracellular high-molecular-weight Ca2+-dependent PLA2s (cPLA2s), which require micromolar concentrations of Ca2+; and Ca2+-independent PLA2s (iPLA2s) and platelet-activating factor acetylhydrolases (PAF-AHs) (28).

Whereas the involvement of PLA2 in the release of AA during ischemia/hypoxia has been established in some cells and tissues (41, 47, 50), the level of PLA2 activation and the regulation of the specific PLA2 subtypes involved are incompletely understood. In a number of cell types, including cardiomyocytes (41), vascular smooth muscle cells (43), and pancreatic β-cells (58), the majority of PLA2 activity appears to be of the iPLA2 type and sensitive to bromoenol lactone (BEL). Induction of iPLA2 activity during ischemia has been demonstrated in ischemic rabbit hearts (21, 25) and proximal tubules (48). Furthermore, in the myocardium, iPLA2 activation is reported to be implicated in the induction of lethal malignant tachyarrhythmias due to increased phospholipid hydrolysis (36, 61).

Two mammalian group VI iPLA2s, designated iPLA2-VIA and iPLA2-VIB, have been cloned (28). iPLA2-VIA, also known as iPLA2-β, has been implicated in diverse cellular processes including membrane remodeling (7), cell proliferation (39, 49), cell volume regulation (see Refs. 29 and 31), insulin secretion (8), programmed cell death (see, e.g., Refs. 4, 46, and 52), and intracellular calcium mobilization (53). iPLA2-VIA contains eight NH2-terminal ankyrin repeats plus a lipase consensus site, which appears to constitute a homotetramer in its functional form and possess PLA2, PLA1, and lysophospholipase as well as transacylase activity (1, 57). The human iPLA2-VIA mRNA transcript is found in five different splice variants, of which two, termed iPLA2-VIA-1 and iPLA2-VIA-2, encode catalytically active proteins. iPLA2-VIA-2 is an extended version of iPLA2-VIA-1 that contains an additional 54-amino acid insert encoded by exon 9, which interrupts the last ankyrin repeat (32). Two other splice variants, iPLA2-VIA-ankyrin-1 and iPLA2-VIA-ankyrin-2, are inactive forms of ~50 kDa that have maintained the ankyrin repeats but lack the COOH-terminal part containing the lipase motif (32). The iPLA2-VIA-ankyrin variants have been proposed to regulate iPLA2-VIA activity in a dominant negative manner through interaction with the iPLA2-VIA functional unit (33, 39). IPLA2-VIB, also designated iPLA2-{gamma}, exhibits several splice variants (38) and different translational products originating from different in-frame start codons resulting in multiple expressed protein products (37). iPLA2-VIB has been assigned roles in organelle membrane remodeling (44) and protection against oxidative stress (16).

To date, little information is available on the expression and regulation of PLA2 subtypes in skeletal muscle. In mouse skeletal muscle-derived myotubes (C2C12), we have previously demonstrated that simulated ischemia [oxygen and glucose deprivation (OGD)] increased the release of AA to the extracellular medium and that this was blocked by the iPLA2 inhibitors BEL and arachidonyl trifluoromethyl ketone (31). The present study addresses the role of iPLA2 in AA accumulation during OGD in C2C12 myotubes. Here, we demonstrate that OGD-induced cell death is associated with an increase in iPLA2-VIA activity and involves changes in the levels of mRNA and protein for different iPLA2-VIA isoforms.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Reagents and media. Unless otherwise indicated, reagents were of the highest available grade and obtained from Sigma-Aldrich (Brondby, Denmark). Alexa fluor 568-conjugated anti-mouse antibody was from Molecular Probes (Leyden, The Netherlands), and FITC-conjugated anti-rabbit antibody was from Jackson Immuno Research (Suffock, UK). Paraformaldehyde was prepared fresh regularly as a 20% stock solution in double-distilled (dd)H2O, filtered, and kept at 4°C. The standard isotonic medium contained (in mM) 143 NaCl, 5 KCl, 1 Na2HPO4, 1 CaCl2, 0.1 MgSO4, and 10 HEPES (pH 7.4). Tris-buffered saline (TBS) contained (in mM) 150 NaCl, 10 Tris·HCl, 1 MgCl2, and 1 EGTA (pH 7.4).

Cell culture and ischemic simulation. The muscle cell culture used is a clone of the C2C12 myoblast cell line that was isolated and cultured as previously described (30). Myoblasts were grown in 10 ml DMEM (catalog no. 32430, Invitrogen, Brondby, Denmark) supplemented with 10% FCS, 100 IU/ml penicillin, 100 µg/ml streptomycin sulfate, 20 µg/ml gentamicin, and 3 µg/ml amphotericin B (Invitrogen). At confluence, the medium was changed to DMEM containing 2% FCS, and myoblasts were left to differentiate into multinucleated myotubes for 2–4 days. Cells were kept in an atmosphere of 95% air-5% CO2 at 37°C. C2C12 myoblasts from passages 3 to 8 were used for experiments. The in vitro ischemia model was set up to combine the stresses of anoxia, glucose deprivation, and stagnant incubation medium (OGD). To create the ischemic environment for protein and mRNA extracts, cells were sealed in either 25- or 75-cm2 culture flasks. Prior to the sealing step, media in the flasks were exchanged with a deoxygenated normal isotonic ringer and gassed with 100% N2 for 2 min to eliminate oxygen from the remaining air space. Subsequently, the flasks were tightly sealed, preventing gas from entering. Cell lysates were isolated as described below but under a constant flow of N2.

Preparation of cell lysates. C2C12 myotubes grown in 75-cm2 culture flasks were washed in ice-cold PBS and lysed in ice-cold lysis buffer [for activity measurements: 50 mM HEPES, 1 mM EDTA, 1 mM Na-ortho-vanadate, and 1:100 (vol/vol) protease inhibitors containing AEBSF, pepstatin A, E-64, bestatin, leupeptin, and aprotinin (Sigma-Aldrich); and for Western blots: 20 mM HEPES, 150 mM NaCl, 1 mM EDTA, 10% glycerol, 1% Triton X-100, 1 mM Na-ortho-vanadate, and 1:100 (vol/vol) protease inhibitors]. Cells were scraped off and placed on ice. The resulting cell lysate was sonicated two times for 10 s on ice, the homogenate was centrifuged at 10,000 g at 4°C for 10 min, and the supernatant was transferred for protein determination. Cytosolic and mitochondrial fractions were separated using a mitochondria isolation kit from Pierce (catalog no. 89874, Rockford, IL) and the protocol supplied by the manufacturer. Membrane fractions was isolated as described by Haustetter et al. (24). Lysates were stored at –80°C until use.

PLA2 activity in cell lysates. C2C12 cell lysates were spun through 30-kDa cutoff filters (Millipore Microcon YM-30, 12 min, 14,000 g). Activity was measured using the PLA2 substrate arachidonoyl thio-phosphatidylcholine and the protocol recommended by the manufacturer (Cayman Chemical, Ann Harbor, MI). In brief, aliquots of 10 µl of supernatants containing 60–80 µg protein were added to 96-well microtiter plates together with 5 µl of assay buffer (80 mM HEPES, 150 mM NaCl, 10 mM CaCl2, 4 mM Triton X-100, 30% glycerol, and 1 mg/ml BSA). Arachidonoyl thio-phosphatidylcholine was dissolved in 160 mM HEPES, 300 mM NaCl, 20 mM CaCl2, 8 mM Triton X-100, 60% glycerol, and 2 mg/ml BSA and diluted 1:1 in ddH2O. For measurements of Ca2+-independent activity, 10 mM CaCl2 was replaced with 5 mM EDTA. The reaction was initiated by the addition of 200 µl of substrate to each well, allowed to run for 60 min, and stopped by the addition of 10 µl of 25 mM 5,5'-dinitrobis-(2-dinitrobenzoic azid) (DTNB) plus 475 mM EGTA in 0.5 M Tris·HCl (pH 8.0). The absorbance was measured at 405 nm in a Fluostar Optima plate reader (BMG LabTechnologies, Offenburg, Germany). PLA2 activity was expressed in micromoles per milligram of protein per minute determined from the extinction coefficient of DNTB and the protein content in the supernatants using Lambert-Beer's law. BEL-sensitive PLA2 activity was estimated as the total activity minus activity in the presence of 10 µM BEL.

Isolation of RNA, RT-PCR, and quantitative real-time PCR. Total RNA was isolated from C2C12 myotubes using the RNeasy Mini Kit (Qiagen, Ballerup, Denmark) according to the manufacturer's protocol. RNA quality was evaluated by gel electrophoresis. cDNA was prepared in a total volume of 40 µl by hybridization of 500 ng random primers to 4 µg RNA at 65°C for 5 min followed by extension at 42°C for 50 min in the presence of 200 units Superscript II reverse transcriptase (Invitrogen), 500 µM dNTP, 10 µM DTT, 50 mM Tris·HCl (pH 8.3), 75 mM KCl, and 3 mM MgCl2. Finally, the reverse transcriptase was inactivated at 70°C for 15 min. Conventional PCR was performed in a total volume of 20 µl containing 1 µl of RT reaction cDNA, 0.5 mM dNTPs, 0.5 µM of each primer, 2 mM MgCl2, and 2 units Taq polymerase in PCR buffer [50 mM KCl and 10 mM Tris·HCl (pH 9.0)] and run for 35 cycles. Primers used for standard PCR were iPLA2-VIA, forward exon 8: 5'-TGGGGAGACTCCTGCATTGA-3', forward exon 9: 5'-AGCTGCAACTCACCCCCTCT-3', forward exon 12: 5'-GGAGTTCCTGAAGCGGGAGT-3', reverse exon 11: 5'-TCTGTCATCTTGGTGTGCTC-3', and reverse exon 13: 5'-TGAGTCGGTGGCTTCAGGTT-3'; iPLA2-VIB, forward: 5'-GAATAACCCTTCGGCCTTGG-3' and reverse: 5'-AGAAGGCAACAGGCCATCAA-3'; iPLA2-{delta}, forward: 5'-TCACTGGGGAACCTCTCATC-3' and reverse: 5'-CTCCACAGCTGTCCAGTCAA-3'; iPLA2-{varepsilon}, forward: 5' ATGGCAAACTTGTGGGAGAC-3' and reverse: 5'-GAGCATTCGATGGTCATCCT-3'; iPLA2-{zeta}, forward: 5'-TCCGAGAGATGTGCAAACAG-3' and reverse: 5'-CTCCAGCGGCAGAGTATAGG-3'; cPLA2-IVA, forward: 5'-GTGGGCGAAAATGAACAAGC-3' and reverse: 5'-CGATTCGGGGTCATCAAAAA-3'; cPLA2-IVB, forward: 5'-GGCACTGGCCAACCTCTATG-3' and reverse: 5'-ATTGCTCCAGATGCCTTCCA-3'; cPLA2-IVC, forward: 5'-ACCCTGCACTTGGGGCTTAT-3' and reverse: 5'-TCCTTGATGCTGGGGTCATT-3'; and PAF-AH(II), forward: 5'-GGTTCAGGGCGTCATACTCG-3' and reverse: 5'-AGCCGTAGCTCCTCCAAAGG-3'. PCR products was confirmed by PCR cloning and sequencing. Real-time PCR was performed in triplicate using Stratagene MX4000 Real-Time PCR equipment and Taqman Universal PCR Master Mix (ABI, Foster City, CA) in a total volume of 20 µl containing 1 µl of the RT reaction, 200 nM of probe and primers, and 10 µl of 2x Mastermix. Probe and primers for real-time PCR were designed to avoid amplification of genomic DNA using Primer Express (ABI, Foster City, CA) or MWG homepage software (MWG Biotech, Martinsried, Germany). Probe and primer sequences were iPLA2-VIA, probe: 5'-CGGCATCCAGTACTTCAGACTGAACCCC-3', forward exon 16: 5'-GGGCCTGGTGCGAGATG-3', and reverse exon 17: 5'-GCGTTGACCAGCACTGCAT-3'; iPLA2-VIA, exon 9 probe: 5'-CTCAGCCCCCAGCGATCAGCTTAAACAA-3', forward exon 9: 5'-GACCACCACTTTCCAATCATC-3', and reverse exon 10: 5'-TGGGCATGAGATCCTGAAG-3'; iPLA2-VIA, exon 9A probe: 5'-TAGGCAGGCAGTCACTCAAACCAG-3', forward exon 9A: 5'-CCCAGCGATCAGCTTAAAC-3', reverse exon 10: 5'-TGGGCATGAGATCCTGAAG-3'; iPLA2-VIB, probe: 5'-TCAAGATGGAGGTCTGCTTCTGAATAACCCT-3', forward exon 7: 5' GCTCCAGGCTACTTTGCAGAGT-3', and reverse exon 8: 5'-CCAGATGCATTTACATTCGTGAA-3'; and 18S rRNA, probe: 5'-TACCGCGGCTGCTGGCACC-3', forward: 5'-TTTAATATACGCTATTGGAGCTGGAA-3', and reverse: 5'-GGATCCATTGGAGGGCAAGT-3'. Probes were labeled with FAM in the 5'-end and TAMRA in the 3'-end. The signal for mRNA expression was normalized to the reference gene level (18S rRNA), and the relative expression ratio was calculated using the following equation: Formula, where Etarget and Eref are the PCR amplification efficiencies for the target and reference gene (18S) and {Delta}Ct target and {Delta}Ct ref are the changes in threshold cycle (Ct) values for target and reference gene transcripts.

PCR cloning for sequencing. PCR products were cloned using TOPO TA cloning according to the manufacturer's instructions (Invitrogen). In brief, PCR products were extracted (Qiagen), ligated into the PCR-II TOPO vector, and transformed into Oneshot competent Escherichia coli. After a 30-min incubation on ice, cells were heat shocked for 30 s at 42°C, and 250 µl SOC medium (2%) was added; 50 µl of each transformation were plated on LB plates containing X-gal and ampicillin and left to incubate overnight. Positive colonies were picked and cultured in ampicillin containing LB medium for 18 h, and DNA was isolated (Perfectprep mini kit, Eppendorf, Horsholm, Denmark). Plasmids were analyzed by restriction analysis and sequenced (MWG Biotech).

Preparation of antibodies to iPLA2-VIB. The generation of rabbit polyclonal antiserum was performed at the peptide synthesis and antibody production facilities at the Louisiana State University Health Sciences Center. A peptide corresponding to an epitope in the COOH-terminal part of mouse and human iPLA2-VIB (SHYLGGCQYK) was synthesized and conjugated to keyhole limpet hemocyanin carrier protein. Rabbits were immunized on day 0 and boosted again on days 14 and 21 with the conjugated peptide. Immunized serum was tested by ELISA before bleeding of the rabbits, the serum was prepared, and 0.1% sodium azide was added as a preservative. The antiserum was affinity purified with the use of the nonconjugated synthetic peptide and a Pierce AminoLink Plus Immobilization Kit, and the affinity purified antibody was desalted and concentrated using Millipore centricon YM50 columns according to the manufacturer's instructions.

Western blot analysis. For Western blot analysis, equivalent amounts of C2C12 protein (20–30 µg) were separated by SDS-PAGE on precast NuPAGE 10% bis-Tris gels (Invitrogen). Proteins were transferred to nitrocellulose membranes, blocked in 5% nonfat dry milk in TBS for 1 h, and then incubated with primary antibody overnight {cPLA2-IVA, 1:500 (sc-454, Santa Cruz Biotechnology, Santa Cruz, CA); iPLA2-VIA, 1:500 (sc-25504, Santa Cruz Biotechnology); iPLA2-VIA, 1:500 (CAY-160507, Cayman Chemical); iPLA2-VIB, 1:100 (Cell Signaling Technology, Danvers, MA); voltage-dependent anion channel (VDAC), 1:400 (no. 4866, Cell Signaling Technology); and TMX3, 1:250 [a generous gift from Dr. Ellgaard, University of Copenhagen (see Ref. 24)]}. Membranes were washed in TBS and incubated with alkaline phosphatase-conjugated anti-rabbit or anti-mouse secondary antibodies (1:600, Jackson Immuno Research). Immunospecific staining was developed on the membranes using 5-bromo-4-chloro-3-indolyl phosphate/nitroblue tetrazolium (KPL, Gaitherburg, MD). Densities of immunoreactive proteins were quantified using UN-SCAN-IT gel automatic digitizing software (version 5.1).

Short interfering RNA knockdown of iPLA2-VIA. The sequence (5'-AACAGCACAGAGAAUGAGGAG-3') for the short interfering (si)RNA used was originally published by Su et al. (54). Double-stranded RNA was subsequently synthesized by MGW Biotech. C2C12 myotubes were transfected with 20 nM siRNA complexed to GeneEraser transfection reagent according to the manufacturer's instructions (Stratagene, La Jolla, CA) and allowed to differentiate in 2% growth medium containing no antibiotics for another 48 h. The GeneEraser/siRNA-containing medium was removed prior to the launch of the experiments. Protein knockdown was evaluated by Western blot analysis and/or RT-PCR, and a scrambled siRNA (Santa Cruz Biotechnology) was used the negative control.

Transfection with iPLA2-VIA vectors. C2C12 myoblasts or myotubes were transfected with control vector (pcDNA3), iPLA2-VIA-1 vector, or iPLA2-VIA-2 vector [a kind gift from Dr. Jackowski (15)] using Lipofectamine 2000 according to the manufacturer's instructions (Invitrogen). One microgram vector and 5 µl Lipofectamine 2000 were used per well in 6- and 96-well dishes; 0.1 µg vector and 0.5 µl Lipofectamine 2000 were used in 24-well dishes. Transfection was performed 24 h prior to the initiation of experiments.

Immunocytochemistry. C2C12 myoblasts were seeded at a density of 4,000 cells/well in eight-chamber polystyrene vessel glass slides (Becton Dickinson, Brondby, Denmark) and grown until 50–60% confluence. Following stimulation, cells were fixed directly in the chambers with 2% paraformaldehyde, washed in PBS, and permeabilized for 15 min in 0.1% Triton X-100 in PBS. Permeabilized cells were washed, blocked in 2.5% BSA for 30 min, and labeled with rabbit iPLA2-VIA (1:100, Cayman Chemical), mouse cytochrome c clone 6H2.B4 (1.250, BD Pharmingen, Erembodegem, Belgium), and mouse protein disulphide-isomerase (PDI; 1:250, Affinity Bioreagents, Golden, CO) in PBS with 1% BSA overnight. Cells were washed three times in PBS with 1% BSA and incubated with FITC-conjugated anti-rabbit and Alexa fluor 568-conjugated anti-mouse secondary antibodies (1:400) for 2 h. After three washes in PBS, the chamber vessel was removed, and slides were mounted [90% glycerol, 10% of 10x PBS, and 2% (wt/vol) N-propylgalleate]. For OGD experiments, cells were incubated for 3–6 h in deoxygenated experimental solution in a controlled atmosphere incubator at 37°C with 0.1% CO2 and 0.5% O2 (Jouan IG 750, Winchester, VA). Following OGD, cells were fixed with 2% paraformaldehyde under hypoxic conditions and treated as described above. Cells were visualized using the x40/1.25 numerical aperture plan apochromat objective of a Leica DM IRB/E microscope with a Leica TSC NT confocal laser scanning unit (Leica Lasertechnik, Heidelberg, Germany). For all experiments, settings were carefully adjusted to avoid bleed through between the FITC and TRIC channels. No or negligible labeling was seen in the absence of primary antibodies (not shown). The optical slice thickness was 1 µm. Images are frame averaged and shown in RGB pseudocolor.

Cell viability assay. Cell viability was determined using the MTT assay. C2C12 myoblasts were seeded in 96-well plates at a density of 2,000 cells/well, grown to confluence, and allowed to differentiate for 2–4 days. Transfections were performed in the last part of the differentiation period as described above (24 h for expression or 48 h for siRNA). Cells transfected with siRNA control (Santa Cruz Biotechnology) or control vector (pcDNA3, Invitrogen) were included and used to correct absorbance values in each experiment. Each condition was performed in at least triplicate. At the start of the experiments, the medium was replaced with 100 µl experimental solution and incubated under normoxic or OGD conditions for 3–24 h. Two hours before the end of the incubation period 10 µl of 2 mM MTT were added to the wells. For OGD, MTT was deoxygenated and added under hypoxic conditions. Experiments were timed such that equal cell growth/diffentiation times were ensured. Following incubation, cells were lysed in 100 µl of 10 mM HCl containing 10% SDS and incubated for 18 h. Absorbance was read at 570 nm in a Fluostar Optima plate reader (BMG LabTechnologies), and cell death was expressed as the reduction in cell viability relative to normoxic controls.

Statistics and data presentation. Data are presented as means ± SE. Statistical significance was estimated by paired Student's t-test and/or one-way ANOVA, with 0.05 as the level of significance.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
iPLA2 activity in C2C12 myotubes was examined by assessing the BEL and Ca2+ sensitivity of total PLA2 activity toward a synthetic AA-containing substrate. As shown in Fig. 1A, total PLA2 activity under control conditions was reduced by 40% following 20 min of preincubation with 10 µM BEL. The omission of Ca2+ from the activity buffer did not affect PLA2 activity significantly, nor did it change the magnitude of BEL inhibition (Fig. 1A). This indicates that 1) cPLA2 activity constitutes a negligible fraction of the total PLA2 activity and 2) ~60% of iPLA2 activity implicates iPLA2s other that iPLA2-VIs. These may include BEL-insensitive iPLA2-IVC or iPLA2-VIIB [PAF-AH (II)], which was found to be expressed in C2C12 myotubes (Fig. 1B). mRNAs for BEL-sensitive iPLA2-VIA, iPLA2-VIB, iPLA2-VIC (neuropathy target esterase), and iPLA2-VIE (TTS-2.2) were also expressed in C2C12 myotubes (Fig. 1B). Subsequently, the BEL sensitivity of the total activity was used as a measure of iPLA2-VI activity. The BEL-sensitive fraction of PLA2 activity was estimated at ~15 nmol·mg protein–1·min–1 under control conditions (normoxic) and gradually increased with time during OGD to a peak at ~35 nmol·mg protein–1·min–1 after 2–3 h (Fig. 2). To investigate the specific involvement of iPLA2-VIA in OGD-induced PLA2 activity, siRNA was employed. As shown in Fig. 3A, iPLA2-VIA expression was significantly reduced by iPLA2-VIA siRNA knockdown. Under normoxic conditions, iPLA2-VIA siRNA knockdown had only a minor effect on BEL-sensitive PLA2 activity (Fig. 3B). In contrast, OGD-induced BEL-sensitive PLA2 activity was completely abolished by iPLA2-VIA siRNA knockdown, whereas it was unaffected in untreated or control siRNA-treated cells (Fig. 3B). Thus, OGD-induced BEL-sensitive PLA2 activity appears to be primarily mediated by iPLA2-VIA.


Figure 1
View larger version (38K):
[in this window]
[in a new window]

  		Fig. 1. PLA2 activity and subtype expression in C2C12 myotubes. A: PLA2 activity under control (Ctrl) conditions and in the presence and absence of Ca2+ and bromoenol lactone (BEL). The synthetic PLA2 substrate arachidonoyl thio-phosphatidylcholine was used to estimate PLA2 activity directed at arachidonic acid-containing phospholipids. PLA2 activity was estimated in the presence of Ca2+ (10 mM) or absence (Ca2+-free buffer chelated with 5 mM EDTA). BEL was used in a final concentration of 10 µM. PLA2 activity is shown in micromoles per milligram of protein per minute. Values are means ± SE of 6 experiments. *Significantly different (P < 0.05) relative to Ca2+ control; #significantly different relative to Ca2+-free control. B: PLA2 mRNA expression in C2C12 myotubes. Lane 1, Ca2+-dependent PLA2 (cPLA2)-IVA; lane 2, cPLA2-IVB; lane 3, cPLA2-IVC; lane 4, Ca2+-independent PLA2 (iPLA2)-VIA (primers used were forward exon 12 and reverse exon 13; see MATERIALS AND METHODS); lane 5, iPLA2-VIB; lane 6, platelet-activating factor acetylhydrolase II [PAF-AH(II)]; lane 7, iPLA2-{delta}; lane 8, iPLA2-{varepsilon}; lane 9, iPLA2-{zeta}; lane 10, iPLA2-{varepsilon} from mouse HL-1 cells (PCR positive control). Bp markers are indicated.

 

Figure 2
View larger version (15K):
[in this window]
[in a new window]

  		Fig. 2. Oxygen and glucose deprivation (OGD) stimulates BEL-sensitive PLA2 activity in C2C12 myotubes. BEL-sensitive PLA2 activity in protein extracts from C2C12 myotubes was estimated throughout a 6-h OGD period. Cells were incubated under control (normoxic, 0 h) conditions or exposed to OGD for various time periods. Protein was extracted and assayed for PLA2 activity as described in MATERIALS AND METHODS in the absence and presence of 10 µM BEL, and the difference was used to calculate BEL-sensitive activity. Protein samples taken under control conditions (0 h) and after 3 h of normoxia (ctrl) were used as controls. Values are mean ± SE of 4 experiments. *Significantly different (P < 0.05) from 0 h normoxic control.

 

Figure 3
View larger version (19K):
[in this window]
[in a new window]

  		Fig. 3. Short interfering (si)RNA knockdown of iPLA2-VIA blocks OGD-induced, BEL-sensitive PLA2 activity. C2C12 myotubes transfected with 20 nM iPLA2-VIA (VIA-Si), control siRNA (Ctrl-Si), or untreated (Ctrl) were exposed to OGD for 3 h. C2C12 myotubes were also incubated under normoxic conditions as indicated. A: representative Western blot showing the reduction in iPLA2-VIA protein levels by siRNA targeting (top) and load control (Ponceau staining of polyvinylidene difluoride membrane; bottom). B: BEL-sensitive PLA2 activity levels under normoxia and after 3 h of OGD in nontransfected, iPLA2-VIA siRNA-transfected, and control siRNA-transfected myotubes. Values are means ± SE of 4 experiments. *Significantly different (P < 0.05) from nontransfected normoxic myotubes; #significantly different from OGD-treated myotubes.

 
During OGD, the mRNA level of iPLA2-VIA did not change significantly, whereas that of iPLA2-VIB declined following 2 h of OGD (Fig. 4). At the protein level, iPLA2-VIA abundance increased moderately within the initial 1–3 h of OGD and remained elevated for the 6 h investigated (Fig. 5A). In contrast, no changes in protein expression for cPLA2-IVA (Fig. 5A) or in 63-, 77-, and 88-kDa iPLA2-VIB proteins (Fig. 5B) were detected.


Figure 4
View larger version (9K):
[in this window]
[in a new window]

  		Fig. 4. Quantification of OGD-induced changes in mRNA levels for iPLA2-VIA and iPLA2-VIB. Total RNA was isolated from C2C12 myotubes exposed to OGD for 0–6 h. RNA was reverse transcribed and cDNA was real-time PCR amplified using the Taqman Real-time PCR probe system. Primers and probes are described in MATERIALS AND METHODS. mRNA levels were normalized to the 18S rRNA housekeeping gene and shown relative to the normoxic control sample (0 h). Values are means ± SE of 6 experiments. *Significantly different (P < 0.05) from the 0 h normoxic control.

 

Figure 5
View larger version (42K):
[in this window]
[in a new window]

  		Fig. 5. Changes in iPLA2-VIA, iPLA2-VIB, and cPLA2-IVA protein mass during OGD. Protein was isolated from C2C12 myotubes exposed to OGD for 0–6 h and subjected to Western blot analysis. A: Western blots and quantification of the changes in the levels of iPLA2-VIA and cPLA2-IVA. B, right: Western blot showing expression of iPLA2-VIB isoforms during OGD in C2C12 myotubes. Left, quantification of the 63-kDa iPLA2-VIB isoform. All iPLA2-VIB isoforms were similar with respect to variations in intensity. Antibodies used were iPLA2-VIA (CAY-160507, 1:500), cPLA2-IVA (sc-454, 1:500), and iPLA2-VIB. Values are means ± SE of 4 experiments for iPLA2-VIA and cPLA2-IVA and 3 experiments for iPLA2-VIB. *Significantly different (P < 0.05) from the 0 h normoxic control.

 
iPLA2-VIA catalytic activities are thought to be regulated through protein-protein interactions between active and inactive iPLA2-VIA isoforms, which are the product of alternative mRNA splicing. The inactive variants, known as iPLA2-VIA-ankyrins (32, 33), were initially identified from human sources, and their presence in mice has not yet been confirmed. Using primers designed to anneal on both sides of mouse exon 9 and 10 splice sites (forward exon 8 and reverse exon 11), two fragments were identified from C2C12 cDNA (Fig. 6A, lane 1). DNA sequencing confirmed these to represent iPLA2-VIA-1 and VIA-2 transcripts (i.e., without and with 165-bp exon 9, respectively). Exchanging the forward primer with one that annealed inside exon 9 (forward exon 9) resulted in amplification of the two fragments (Fig. 6A, lane 2). Both of these cDNAs contained the expected sequence of exon 9 with the longer form containing an extra insert of 29 bp inserted between the exon 9 and 10 splice sites (Fig. 6B). The presence of this insert resulted in a frame shift inducing a stop codon inside exon 10. This transcript encodes a putative truncated iPLA2-VIA protein of ~53 kDa that lacks its COOH-terminal lipase consensus site.


Figure 6
View larger version (23K):
[in this window]
[in a new window]

  		Fig. 6. Identification of a mouse iPLA2-VIA-ankyrin splice variant. A: amplification of C2C12 cDNA using specific primers. Lane 1, amplification of C2C12 cDNA using primers forward exon 8 and reverse exon 11 (see MATERIALS AND METHODS). Gel fragments 3 and 4 in the gel were sequenced (sequence not shown) and shown to represent iPLA2-VIA-1 and iPLA2-VIA-2, respectively. Lane 2, amplification using primers forward exon 9 and reverse exon 11 (see MATERIALS AND METHODS). Both fragments 1 and 2 contain part of exon 9 (underlined) with fragment 2 containing the extra 29-bp insert between exon 9 and exon 10 (bold; designated here as exon 9A). B: sequence alignment of the two sequences obtained from fragments 1 and 2. C: DNA sequence alignment of mouse and human exon 9A and the mouse genomic sequence located at chromosome 15. Putative splice acceptor and donor sites are marked in bold.

 
The possible involvement of the iPLA2-VIA-ankyrin variant in regulating iPLA2-VIA activity was evaluated by tracking the expression of exon 9A-containing iPLA2-VIA mRNA transcripts through 6 h of OGD using quantitative real-time PCR. As the transcripts containing exon 9A also contain exon 9, a separate amplification of exon 9 was required to distinguish changes in exon 9A from changes in exon 9-containing mRNA. Using the exon-specific primers, the basal levels of exon 9- and exon 9A-containing iPLA2-VIA transcripts were estimated to be ~8- and 16-fold lower compared with the total iPLA2-VIA transcript level (data not shown, n = 4). In contrast to the unchanged total iPLA2-VIA mRNA level throughout OGD (Fig. 4), the level of exon 9A-containing transcript was significantly reduced within 30 min, whereas the level of exon 9-containing transcript was significantly reduced at 4 h of OGD (Fig. 7A). An iPLA2-VIA antibody that has been shown to react with human iPLA2-VIA-ankyrin (39) recognized a protein of the expected 50 kDa in C2C12 protein lysates (Fig. 7B). The level of the iPLA2-VIA-ankyrin protein was markedly reduced within the first hour of OGD (Fig. 7B), indicating that the drop in iPLA2-VIA exon 9A mRNA is paralleled by a reduction in iPLA2-VIA-ankyrin protein levels. In resting C2C12 myoblasts, iPLA2-VIA seemed to localize throughout the cell but predominantly in the perinuclear region (Fig. 8A). Following 3 and 6 h of OGD, iPLA2-VIA staining seemed to concentrate in a granular manner in the perinuclear region (Fig. 8A). Substantial colocalization between iPLA2-VIA and the endoplasmic reticulum (ER) luminal protein PDI was observed both under control conditions and after 3 and 6 h of OGD. In contrast, colocalization between iPLA2-VIA and the mitochondrial marker cytochrome c was not detected under any condition (Fig. 8A). Subcellular fractionation and Western blot analysis using VDAC as a mitochondrial marker showed that iPLA2-VIA was predominantly found in cytosolic and membrane fractions and verified that no, or very low levels of, iPLA2-VIA was present in isolated mitochondria (Fig. 8B). No substantial changes in the subcellular distribution of iPLA2-VIA could be detected following 3 h of OGD (Fig. 8B). While addressing this issue and its possible relevance to OGD-induced cell death is beyond the scope of the study, it may be noted that mitochondrial integrity appeared to be unperturbed in the time period studied, in contrast to our findings in cultured cardiomyocytes (2).


Figure 7
View larger version (12K):
[in this window]
[in a new window]

  		Fig. 7. Regulation of iPLA2-VIA-ankyrin mRNA and protein during OGD in C2C12 myotubes. A: quantitative real-time PCR showing the expression of exon 9- and exon 9A-containing mRNA transcripts normalized to 18S rRNA levels (relative expression). Data represent means ± SE of 4 experiments. *Significantly different (P < 0.05) from the 0 h normoxic control. B: equal amounts of cell lysate from C2C12 myotubes were subjected to Western blot analysis using the iPLA2-VIA antibody (sc-25504), which recognizes the iPLA2-VIA-ankyrin variant. One of three representative blots is shown. iPLA2-VIA-ankyrin variant migrated very close to the 50-kDa marker. C: ratio of mRNA and protein of full-length iPLA2-VIA versus iPLA2-VIA-ankyrin following 0, 1, 2, 4, and 6 h of OGD stimulation. Data for the mRNA ratio were calculated as relative mRNA levels of total iPLA2-VIA mRNA (Fig. 5) over relative iPLA2-VIA exon 9A mRNA levels (A). The protein ratio was calculated from relative levels of full-length 85-kDa iPLA2-VIA protein (Fig. 6) and 50-kDa iPLA2-VIA-ankyrin from 3 blots (B).

 

Figure 8
View larger version (53K):
[in this window]
[in a new window]

  		Fig. 8. Subcellular localization of iPLA2-VIA. A: C2C12 myoblasts were seeded in 8-chamber polystyrene vessel glass slides, exposed to OGD for the times indicated, fixed with paraformaldehyde, permeabilized, blocked for unspecific protein binding, and labeled with primary antibody [iPLA2-VIA, cytochrome c (Cyt C), and protein disulphide-isomerase (PDI)] in PBS with 1% BSA overnight. Following incubation, cells were washed, incubated with FITC-conjugated anti-rabbit and Alexa568 anti-mouse secondary antibodies (1:400) for 2 h, and mounted (for details, see MATERIALS AND METHODS). For ODG, cells were incubated for 3 and 6 h in deoxygenated experimental solution in a controlled atmosphere incubator at 37°C with 0.1% CO2-0.5% O2 followed by paraformaldehyde fixation under hypoxic conditions and subsequent treatments as above. Cells were visualized using the x40/1.25 numerical aperture plan apochromat objective of a Leica DM IRB/E microscope with a Leica TSC NT confocal laser scanning unit. Settings were carefully adjusted to avoid bleed through between the FITC and TRIC channels. No or negligible labeling was seen in the absence of primary antibodies (not shown). The optical slice thickness was 1 µm, and images were frame averaged and are shown in RGB pseudocolor. The yellow signal in the merge images indicates the colocalization of iPLA2-VIA and the endoplasmic reticulum (ER) marker PDI. B: Western blots showing subcellular localization of iPLA2-VIA under control conditions and following 3 h of OGD. C, cytosol; Mi, mitochondria; Me, crude membrane. Voltage-dependent anion channel (VDAC) was used as a marker for mitochondria. Membrane fractions were enriched in the ER resident proteins TMX3 and ERp57 (data not shown). Different antibodies had to be used for the Western blots as the Cyt C and PDI antibodies used for the confocal imaging were not recommended for Western blotting by the manufacturers.

 
Finally, we evaluated the involvement of iPLA2-VIA in OGD-induced cell death in C2C12 myotubes. As shown in Fig. 9, exposure to OGD was associated with a time-dependent loss of cell viability. Reduction of iPLA2-VIA protein level by siRNA-mediated knockdown or pharmacological inhibition of iPLA2 using BEL (10 µM) significantly attenuated cell death at 3 h of OGD but not after 6 h of OGD. In contrast, after 24 h of OGD, cell death was augmented by both BEL and iPLA2-VIA knockdown (Fig. 9A). Conversely, overexpression of recombinant mouse iPLA2-VIA-1 or iPLA2-VIA-2 had no significant effect on ODG-induced cell death (Fig. 9B).


Figure 9
View larger version (14K):
[in this window]
[in a new window]

  		Fig. 9. Involvement of iPLA2-VIA in OGD-induced cell death. C2C12 myotubes were cultured in 96-well plates and exposed to normoxia or OGD for 3, 6, and 24 h. A: effect of iPLA2-VIA siRNA knockdown and iPLA2 inhibition (10 µM BEL) on cell viability. Cells were either pretreated with siRNA targeting iPLA2-VIA for 48 h (20 nM) or BEL was added to the incubation medium prior to the start of the experiments. A control siRNA (Ctrl) was included and used to correct iPLA2-VIA siRNA values. B: effect of iPLA2-VIA expression on cell viability. Expressions of iPLA2-VIA-1 or iPLA2-VIA-2 in C2C12 myotubes following 24 h of transfection were verified in Western blots. Mock, nontransfected cells; VIA-1, cells transfected with iPLA2-VIA-1; VIA-2, cells transfected with iPLA2-VIA-2. For cell viability measurements, C2C12 myotubes were transfected iPLA2-VIA-1 or iPLA2-VIA-2 24 h before the start of the experiments. The effect of transfection with the control vector (pcDNA3) on cell viability was used to correct the values of iPLA2-VIA-transfected cells. Cell viability was estimated using the MTT activity assay. Values are means ± SE of 5 experiments in 2 separate series of experiments (A and B) and presented as the relative reduction in cell viability (cell death). *Significantly different from normoxia (0 h); #significantly different from 3 and 24 h of OGD.

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
Skeletal muscle ischemia/hypoxia is a clinical important phenomenon (14). Although studies from other cell types have pointed to an important role for iPLA2 in ischemic/hypoxic-induced cell damage (36, 48, 52, 61, 62), this is, to our knowledge, the first study addressing the role of iPLA2s in skeletal muscle ischemia. The PLA2 activity in C2C12 myotubes toward an AA-containing substrate was found to be mainly Ca2+ independent and inhibited ~40% by BEL, implicating iPLA2-IVs in this activity. As chelation of Ca2+ alone does not separate various iPLA2 activities, we used BEL and/or siRNA to isolate iPLA2-VI activity. The finding that iPLA2-VIA siRNA knockdown had a relative limited effect on BEL-sensitive PLA2 activity under normoxic conditions suggests that other BEL-sensitive PLA2s are active under basal conditions, e.g., iPLA2-VIB. mRNA for iPLA2-{delta} and iPLA2-{zeta} was also expressed in C2C12 myotubes; however, the activity of these PLA2s toward AA-containing phospholipids has been suggested to be low (26, 59). Conversely, the BEL-sensitive PLA2 activity induced by OGD was completely prevented in iPLA2-VIA siRNA-treated C2C12 myotubes, strongly indicating that iPLA2-VIA is activated by OGD. In congruence with this, hypoxia-induced iPLA2 activity also appears to be mediated by iPLA2-VIA in PC12 cells (52).

The changes in OGD-induced activity correlated with the time scale for changes in iPLA2-VIA protein, indicating that increased activity, at least in part, reflects increased protein mass. The absence of a change in iPLA2-VIA mRNA compared with that of the protein suggests that long-term regulation of iPLA2-VIA activity in response to OGD predominantly occurs at the translational and/or posttranslational levels. The OGD-induced induction appears to be specific for iPLA2-VIA, since neither the mRNA nor protein level of iPLA2-VIB was affected by OGD.

The activity of the iPLA2-VIA enzyme complex may be modulated by changes in the ratio of active versus inactive iPLA2-VIA isoforms. The iPLA2-VIA-ankyrin splice variants are characterized by additional short sequences, encoded by exon 9A and 10A, spliced into the mRNA transcript after exon 9 and 10, resulting in frame shifts, the introduction of premature stop codons, and truncated protein products. Larsson and coworkers (33) initially demonstrated that coexpression of full-length and truncated iPLA2-VIA-ankyrin variants suppressed iPLA2 activity, and later findings in CHO cells transfected with hamster iPLA2-VIA and human iPLA2-VIA-ankyrin-1 indicated that it was due to a direct interaction between iPLA2-VIA variants (39). The variant of the mouse iPLA2-VIA transcript identified in the present study appears to be a homolog to human iPLA2-VIA-ankyrins. The 29-bp insert is found in the mouse genome at chromosome 15 in the intron spaced between iPLA2-VIA exon 9 and 10. Although 24 bp shorter, it has ~90% homology to the first part of human exon 9A (Fig. 6C). Notably, the mRNA level of this iPLA2-VIA-ankyrin transcript variant was reduced within 30 min of OGD, whereas the total iPLA2-VIA mRNA level was not significantly altered. iPLA2-VIA-ankyrin protein mass was likewise found to be reduced within 1 h during OGD. Interestingly, although the basal level of iPLA2-VIA-ankyrin mRNA was fairly low, the protein level was high compared with full-length active forms. Thus, if the scenario of an inhibitory interaction between the enzymatically active iPLA2-VIA and iPLA2-VIA-ankyrin is correct, a significant increase in the ratio of active to inactive protein variants during OGD would be required to lift this inhibition. Consistent with this notion, the mRNA ratio increased ~2.5-fold and the protein ratio increased ~8-fold during OGD (Fig. 7C). Hence, the increase in BEL-sensitive activity during OGD may, at least in part, be controlled by a combination of upregulation of active iPLA2-VIA isoforms and downregulation of inactive isoforms. Whether these changes involve increased synthesis or reduced degradation or a combination of both was not investigated.

Localization of iPLA2-VIA in specific cellular compartments (9, 12, 19, 45, 51, 52, 61) and subcellular relocation of the lipase upon activation have previously been demonstrated (45, 60). However, it is unclear whether iPLA2-VIA relocates in response to ischemic stress. OGD in C2C12 was [in agreement with findings in human colon cancer (35), kidney cells (18) and β-cells (9)] associated with increased iPLA2-VIA intensity in the perinuclear region in organelles, which appeared to be the ER (Fig. 8A). This intensity increase does apparently not reflect iPLA2-VIA translocation from the cytosol to the ER as subcellular fractionation did not shown major changes in the subcellular distribution pattern following 3 h of OGD (Fig. 8B). iPLA2-VIA was, in contrast to findings in other cell lines (11, 12, 23, 35, 51, 61), absent from, or very weakly expressed in, mitochondria. In PC12 cells, iPLA2-VIA has been shown to translocate into the nucleus during hypoxia (52); however, we did not find evidence for nuclear translocation in C2C12 myoblasts. Thus, these findings are most consistent with the interpretation that iPLA2-VIA is primarily targeted to the ER in C2C12 cells, where it is concentrated during OGD.

A number of studies have reported that iPLA2 has protective functions in certain cell types, primarily under oxidative stress conditions (16, 18, 37, 42, 55, 56). Conversely, PLA2 may also mediate cell damage following various death-inducing stimuli in other cell systems (3, 17, 61), including during severe oxidative stress (5, 10, 40). In C2C12 myotubes, siRNA knockdown or pharmacological inhibition of iPLA2-VIA was found to moderately counteract OGD-induced cell death in the early OGD phase, whereas in the late phase (24 h), cell death was slightly but significantly increased. In contrast to the effect of inhibition/knockdown, overexpression of iPLA2-VIA did not affect cell viability in C2C12 cells. Thus, although these data indicate that iPLA2-VIA activity may have both damaging and protective functions depending on the degree of exposure to OGD, the moderate effect of its inactivation on cell viability suggests that iPLA2-VIA is not an indispensable mediator of the cell death response under these conditions. It is possible that the role of iPLA2-VIA may vary with the specific death pathways activated, a question not addressed in the present study. The physiological function of iPLA2-VIA in the ER during OGD is unclear; however, studies on ER iPLA2s in oxidant-induced cell death have indicated that ER PLA2 activity may protect against oxidative stress-induced lipid peroxidation (18, 27). On the other hand, iPLA2-VIA has also been proposed to participate in ceramide generation during ER stress-induced apoptosis in INS-1 cells (34). The damaging effects of iPLA2-VIA in the early OGD period might otherwise be mediated by release of FFAs that affect cell signaling or mitochondrial permeability/function (12), by direct disruption of phospholipid membranes (17), or by altering nuclear membrane architecture (52).

In conclusion, deprivation of C2C12 myotubes of oxygen and glycose (OGD) induced specific activation of iPLA2-VIA and reciprocal changes in the expression levels of active and inactive iPLA2-VIA splice variants. iPLA2-VIA accumulated in the perinuclear area, apparently the ER, during OGD. Although not essential for loss of cell viability during OGD, increased iPLA2-VIA activity contributes to the reduced cell viability in the early OGD phase. Together, these findings indicate that iPLA2-VIA activity is an important element of ischemic cell injury in skeletal muscle.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported by Danish Natural Sciences Research Council Grants 21-02-0358 and 21-04-0535, Fonden af 1870, and the FØTEK 3 program/Directorate for Food, Fisheries and Agri Business.


   ACKNOWLEDGMENTS
 
The technical assistance of Dorthe Nielsen is gratefully acknowledged.


   FOOTNOTES
 

Address for reprint requests and other correspondence: K. A. Poulsen. Dept. of Molecular Biology, Univ. of Copenhagen, Universitetsparken 13, Copenhagen Ø DK-2100, Denmark (e-mail: kapoulsen{at}aki.ku.dk)

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.


   REFERENCES
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
1. Ackermann EJ, Kempner ES, Dennis EA. Ca2+-independent cytosolic phospholipase A2 from macrophage-like P388D1 cells. Isolation and characterization. J Biol Chem 269: 9227–9233, 1994.[Abstract/Free Full Text]

2. Andersen AD, Poulsen KA, Lambert IH, Pedersen SF. Roles of NHE1 and iPLA2 in ischemia/reperfusion-induced damage in HL-1 mouse cardiomyocytes (Abstract). FASEB J 21: A684, 2007.[Web of Science]

3. Arai K, Ikegaya Y, Nakatani Y, Kudo I, Nishiyama N, Matsuki N. Phospholipase A2 mediates ischemic injury in the hippocampus: a regional difference of neuronal vulnerability. Eur J Neurosci 13: 2319–2323, 2001.[CrossRef][Web of Science][Medline]

4. Atsumi G, Tajima M, Hadano A, Nakatani Y, Murakami M, Kudo I. Fas-induced arachidonic acid release is mediated by Ca2+-independent phospholipase A2 but not cytosolic phospholipase A2, which undergoes proteolytic inactivation. J Biol Chem 273: 13870–13877, 1998.[Abstract/Free Full Text]

5. Balboa MA, Balsinde J. Involvement of calcium-independent phospholipase A2 in hydrogen peroxide-induced accumulation of free fatty acids in human U937 cells. J Biol Chem 277: 40384–40389, 2002.[Abstract/Free Full Text]

6. Balsinde J. Roles of various phospholipases A2 in providing lysophospholipid acceptors for fatty acid phospholipid incorporation and remodelling. Biochem J 364: 695–702, 2002.[CrossRef][Web of Science][Medline]

7. Balsinde J, Bianco ID, Ackermann EJ, Conde-Frieboes K, Dennis EA. Inhibition of calcium-independent phospholipase A2 prevents arachidonic acid incorporation and phospholipid remodeling in P388D1 macrophages. Proc Natl Acad Sci USA 92: 8527–8531, 1995.[Abstract/Free Full Text]

8. Bao S, Bohrer A, Ramanadham S, Jin W, Zhang S, Turk J. Effects of stable suppression of group VIA phospholipase A2 expression on phospholipid content and composition, insulin secretion, and proliferation of INS-1 insulinoma cells. J Biol Chem 281: 187–198, 2006.[Abstract/Free Full Text]

9. Bao S, Jin C, Zhang S, Turk J, Ma Z, Ramanadham S. β-Cell calcium-independent group VIA phospholipase A2 (iPLA2β): tracking iPLA2β movements in response to stimulation with insulin secretagogues in INS-1 cells. Diabetes 53, Suppl 1: S186–S189, 2004.[Abstract/Free Full Text]

10. Birbes H, Gothie E, Pageaux JF, Lagarde M, Laugier C. Hydrogen peroxide activation of Ca2+-independent phospholipase A2 in uterine stromal cells. Biochem Biophys Res Commun 276: 613–618, 2000.[CrossRef][Web of Science][Medline]

11. Broekemeier KM, Iben JR, LeVan EG, Crouser ED, Pfeiffer DR. Pore formation and uncoupling initiate a Ca2+-independent degradation of mitochondrial phospholipids. Biochemistry 41: 7771–7780, 2002.[CrossRef][Medline]

12. Brustovetsky T, Antonsson B, Jemmerson R, Dubinsky JM, Brustovetsky N. Activation of calcium-independent phospholipase A (iPLA) in brain mitochondria and release of apoptogenic factors by BAX and truncated BID. J Neurochem 94: 980–994, 2005.[CrossRef][Web of Science][Medline]

13. Caro AA, Cederbaum AI. Role of cytochrome P450 in phospholipase A2- and arachidonic acid-mediated cytotoxicity. Free Radic Biol Med 40: 364–375, 2006.[CrossRef][Web of Science][Medline]

14. Carvalho AJ, McKee NH, Green HJ. Metabolic and contractile responses of fast and slow twitch rat skeletal muscles to ischemia and reperfusion. Plast Reconstr Surg 99: 163–171, 1997.[Web of Science][Medline]

15. Chiu CH, Jackowski S. Role of calcium-independent phospholipases (iPLA2) in phosphatidylcholine metabolism. Biochem Biophys Res Commun 287: 600–606, 2001.[CrossRef][Web of Science][Medline]

16. Cummings BS, Gelasco AK, Kinsey GR, McHowat J, Schnellmann RG. Inactivation of endoplasmic reticulum bound Ca2+-independent phospholipase A2 in renal cells during oxidative stress. J Am Soc Nephrol 15: 1441–1451, 2004.[Abstract/Free Full Text]

17. Cummings BS, McHowat J, Schnellmann RG. Phospholipase A2s in cell injury and death. J Pharmacol Exp Ther 294: 793–799, 2000.[Abstract/Free Full Text]

18. Cummings BS, McHowat J, Schnellmann RG. Role of an endoplasmic reticulum Ca2+-independent phospholipase A2 in oxidant-induced renal cell death. Am J Physiol Renal Physiol 283: F492–F498, 2002.[Abstract/Free Full Text]

19. Cummings BS, McHowat J, Schnellmann RG. Role of an endoplasmic reticulum Ca2+-independent phospholipase A2 in cisplatin-induced renal cell apoptosis. J Pharmacol Exp Ther 308: 921–928, 2004.[Abstract/Free Full Text]

20. Farooqui AA, Horrocks LA, Farooqui T. Deacylation and reacylation of neural membrane glycerophospholipids. J Mol Neurosci 14: 123–135, 2000.[CrossRef][Web of Science][Medline]

21. Ford DA, Hazen SL, Saffitz JE, Gross RW. The rapid and reversible activation of a calcium-independent plasmalogen-selective phospholipase A2 during myocardial ischemia. J Clin Invest 88: 331–335, 1991.[Web of Science][Medline]

22. Fredsted A, Mikkelsen UR, Gissel H, Clausen T. Anoxia induces Ca2+ influx and loss of cell membrane integrity in rat extensor digitorum longus muscle. Exp Physiol 90: 703–714, 2005.[Abstract/Free Full Text]

23. Gadd ME, Broekemeier KM, Crouser ED, Kumar J, Graff G, Pfeiffer DR. Mitochondrial iPLA2 activity modulates the release of cytochrome c from mitochondria and influences the permeability transition. J Biol Chem 281: 6931–6939, 2006.[Abstract/Free Full Text]

24. Haugstetter J, Blicher T, Ellgaard L. Identification and characterization of a novel thioredoxin-related transmembrane protein of the endoplasmic reticulum. J Biol Chem 280: 8371–8380, 2005.[Abstract/Free Full Text]

25. Hazen SL, Ford DA, Gross RW. Activation of a membrane-associated phospholipase A2 during rabbit myocardial ischemia which is highly selective for plasmalogen substrate. J Biol Chem 266: 5629–5633, 1991.[Abstract/Free Full Text]

26. Jenkins CM, Mancuso DJ, Yan W, Sims HF, Gibson B, Gross RW. Identification, cloning, expression, and purification of three novel human calcium-independent phospholipase A2 family members possessing triacylglycerol lipase and acylglycerol transacylase activities. J Biol Chem 279: 48968–48975, 2004.[Abstract/Free Full Text]

27. Kinsey GR, McHowat J, Beckett CS, Schnellmann RG. Identification of calcium-independent phospholipase A2{gamma} in mitochondria and its role in mitochondrial oxidative stress. Am J Physiol Renal Physiol 292: F853–F860, 2007.[Abstract/Free Full Text]

28. Kudo I, Murakami M. Phospholipase A2 enzymes. Prostaglandins Other Lipid Mediat 68–69: 3–58, 2002.

29. Lambert IH. Reactive oxygen species regulate swelling-induced taurine efflux in NIH3T3 mouse fibroblasts. J Membr Biol 192: 19–32, 2003.[CrossRef][Web of Science][Medline]

30. Lambert IH, Nielsen JH, Andersen HJ, Ortenblad N. Cellular model for induction of drip loss in meat. J Agric Food Chem 49: 4876–4883, 2001.[CrossRef][Web of Science][Medline]

31. Lambert IH, Pedersen SF, Poulsen KA. Activation of PLA2 isoforms by cell swelling and ischaemia/hypoxia. Acta Physiol (Oxf) 187: 75–85, 2006.[CrossRef][Medline]

32. Larsson Forsell PK, Kennedy BP, Claesson HE. The human calcium-independent phospholipase A2 gene multiple enzymes with distinct properties from a single gene. Eur J Biochem 262: 575–585, 1999.[Web of Science][Medline]

33. Larsson PK, Claesson HE, Kennedy BP. Multiple splice variants of the human calcium-independent phospholipase A2 and their effect on enzyme activity. J Biol Chem 273: 207–214, 1998.[Abstract/Free Full Text]

34. Lei X, Zhang S, Bohrer A, Bao S, Song H, Ramanadham S. The group VIA calcium-independent phospholipase A2 participates in ER stress-induced INS-1 insulinoma cell apoptosis by promoting ceramide generation via hydrolysis of sphingomyelins by neutral sphingomyelinase. Biochemistry 46: 10170–10185, 2007.[CrossRef][Medline]

35. Liou JY, Aleksic N, Chen SF, Han TJ, Shyue SK, Wu KK. Mitochondrial localization of cyclooxygenase-2 and calcium-independent phospholipase A2 in human cancer cells: implication in apoptosis resistance. Exp Cell Res 306: 75–84, 2005.[CrossRef][Web of Science][Medline]

36. Mancuso DJ, Abendschein DR, Jenkins CM, Han X, Saffitz JE, Schuessler RB, Gross RW. Cardiac ischemia activates calcium-independent phospholipase A2beta, precipitating ventricular tachyarrhythmias in transgenic mice: rescue of the lethal electrophysiologic phenotype by mechanism-based inhibition. J Biol Chem 278: 22231–22236, 2003.[Abstract/Free Full Text]

37. Mancuso DJ, Jenkins CM, Gross RW. The genomic organization, complete mRNA sequence, cloning, and expression of a novel human intracellular membrane-associated calcium-independent phospholipase A2. J Biol Chem 275: 9937–9945, 2000.[Abstract/Free Full Text]

38. Mancuso DJ, Jenkins CM, Sims HF, Cohen JM, Yang J, Gross RW. Complex transcriptional and translational regulation of iPLAgamma resulting in multiple gene products containing dual competing sites for mitochondrial or peroxisomal localization. Eur J Biochem 271: 4709–4724, 2004.[Web of Science][Medline]

39. Manguikian AD, Barbour SE. Cell cycle dependence of group VIA calcium-independent phospholipase A2 activity. J Biol Chem 279: 52881–52892, 2004.[Abstract/Free Full Text]

40. Martinez J, Moreno JJ. Role of Ca2+-independent phospholipase A2 on arachidonic acid release induced by reactive oxygen species. Arch Biochem Biophys 392: 257–262, 2001.[CrossRef][Web of Science][Medline]

41. McHowat J, Creer MH. Catalytic features, regulation and function of myocardial phospholipase A2. Curr Med Chem Cardiovasc Hematol Agents 2: 209–218, 2004.[CrossRef][Medline]

42. McHowat J, Swift LM, Arutunyan A, Sarvazyan N. Clinical concentrations of doxorubicin inhibit activity of myocardial membrane-associated, calcium-independent phospholipase A2. Cancer Res 61: 4024–4029, 2001.[Abstract/Free Full Text]

43. Miyake R, Gross RW. Multiple phospholipase A2 activities in canine vascular smooth muscle. Biochim Biophys Acta 1165: 167–176, 1992.[Medline]

44. Murakami M, Masuda S, Ueda-Semmyo K, Yoda E, Kuwata H, Takanezawa Y, Aoki J, Arai H, Sumimoto H, Ishikawa Y, Ishii T, Nakatani Y, Kudo I. Group VIB Ca2+-independent phospholipase A2gamma promotes cellular membrane hydrolysis and prostaglandin production in a manner distinct from other intracellular phospholipases A2. J Biol Chem 280: 14028–14041, 2005.[Abstract/Free Full Text]

45. Pedersen SF, Poulsen KA, Lambert IH. Roles of phospholipase A2 isoforms in the swelling- and melittin-induced arachidonic acid release and taurine efflux in NIH3T3 fibroblasts. Am J Physiol Cell Physiol 291: C1286–C1296, 2006.[Abstract/Free Full Text]

46. Perez R, Melero R, Balboa MA, Balsinde J. Role of group VIA calcium-independent phospholipase A2 in arachidonic acid release, phospholipid fatty acid incorporation, and apoptosis in U937 cells responding to hydrogen peroxide. J Biol Chem 279: 40385–40391, 2004.[Abstract/Free Full Text]

47. Phillis JW, O'Regan MH. A potentially critical role of phospholipases in central nervous system ischemic, traumatic, and neurodegenerative disorders. Brain Res Brain Res Rev 44: 13–47, 2004.[CrossRef][Medline]

48. Portilla D, Shah SV, Lehman PA, Creer MH. Role of cytosolic calcium-independent plasmalogen-selective phospholipase A2 in hypoxic injury to rabbit proximal tubules. J Clin Invest 93: 1609–1615, 1994.[Web of Science][Medline]

49. Sanchez T, Moreno JJ. Calcium-independent phospholipase A2 through arachidonic acid mobilization is involved in Caco-2 cell growth. J Cell Physiol 193: 293–298, 2002.[CrossRef][Web of Science][Medline]

50. Sapirstein A, Bonventre JV. Phospholipases A2 in ischemic and toxic brain injury. Neurochem Res 25: 745–753, 2000.[CrossRef][Web of Science][Medline]

51. Seleznev K, Zhao C, Zhang XH, Song K, Ma ZA. Calcium-independent phospholipase A2 localizes in and protects mitochondria during apoptotic induction by staurosporine. J Biol Chem 281: 22275–22288, 2006.[Abstract/Free Full Text]

52. Shinzawa K, Tsujimoto Y. PLA2 activity is required for nuclear shrinkage in caspase-independent cell death. J Cell Biol 163: 1219–1230, 2003.[Abstract/Free Full Text]

53. Smani T, Zakharov SI, Csutora P, Leno E, Trepakova ES, Bolotina VM. A novel mechanism for the store-operated calcium influx pathway. Nat Cell Biol 6: 113–120, 2004.[CrossRef][Web of Science][Medline]

54. Su X, Mancuso DJ, Bickel PE, Jenkins CM, Gross RW. Small interfering RNA knockdown of calcium-independent phospholipases A2 beta or gamma inhibits the hormone-induced differentiation of 3T3-L1 preadipocytes. J Biol Chem 279: 21740–21748, 2004.[Abstract/Free Full Text]

55. Swift L, McHowat J, Sarvazyan N. Inhibition of membrane-associated calcium-independent phospholipase A2 as a potential culprit of anthracycline cardiotoxicity. Cancer Res 63: 5992–5998, 2003.[Abstract/Free Full Text]

56. Tanaka H, Takeya R, Sumimoto H. A novel intracellular membrane-bound calcium-independent phospholipase A2. Biochem Biophys Res Commun 272: 320–326, 2000.[CrossRef][Web of Science][Medline]

57. Tang J, Kriz RW, Wolfman N, Shaffer M, Seehra J, Jones SS. A novel cytosolic calcium-independent phospholipase A2 contains eight ankyrin motifs. J Biol Chem 272: 8567–8575, 1997.[Abstract/Free Full Text]

58. Turk J, Ramanadham S. The expression and function of a group VIA calcium-independent phospholipase A2 (iPLA2beta) in beta-cells. Can J Physiol Pharmacol 82: 824–832, 2004.[CrossRef][Web of Science][Medline]

59. van Tienhoven M, Atkins J, Li Y, Glynn P. Human neuropathy target esterase catalyzes hydrolysis of membrane lipids. J Biol Chem 277: 20942–20948, 2002.[Abstract/Free Full Text]

60. Wang Z, Ramanadham S, Ma ZA, Bao S, Mancuso DJ, Gross RW, Turk J. Group VIA phospholipase A2 forms a signaling complex with the calcium/calmodulin-dependent protein kinase IIbeta expressed in pancreatic islet beta-cells. J Biol Chem 280: 6840–6849, 2005.[Abstract/Free Full Text]

61. Williams SD, Gottlieb RA. Inhibition of mitochondrial calcium-independent phospholipase A2 (iPLA2) attenuates mitochondrial phospholipid loss and is cardioprotective. Biochem J 362: 23–32, 2002.[CrossRef][Web of Science][Medline]

62. Williams SD, Hsu FF, Ford DA. Electrospray ionization mass spectrometry analyses of nuclear membrane phospholipid loss after reperfusion of ischemic myocardium. J Lipid Res 41: 1585–1595, 2000.[Abstract/Free Full Text]




This article has been cited by other articles:


Home page
 Am. J. Physiol. Cell Physiol.Home page
A.-D. Andersen, K. A. Poulsen, I. H. Lambert, and S. F. Pedersen
HL-1 mouse cardiomyocyte injury and death after simulated ischemia and reperfusion: roles of pH, Ca2+-independent phospholipase A2, and Na+/H+ exchange
Am J Physiol Cell Physiol, May 1, 2009; 296(5): C1227 - C1242.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
293/5/C1605    most recent
00012.2007v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Poulsen, K. A.
Right arrow Articles by Lambert, I. H.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Poulsen, K. A.
Right arrow Articles by Lambert, I. H.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Visit Other APS Journals Online
Copyright © 2007 by the American Physiological Society.