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Am J Physiol Cell Physiol 290: C1350-C1354, 2006. First published December 7, 2005; doi:10.1152/ajpcell.00487.2005
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RECEPTORS AND SIGNAL TRANSDUCTION

Activation of MAPKs in thrombin-stimulated ventricular myocytes is dependent on Ca2+-independent PLA2

Caroline S. Beckett, Karin Pennington, and Jane McHowat

Department of Pathology, St. Louis University School of Medicine, St. Louis, Missouri

Submitted 27 September 2005 ; accepted in final form 1 December 2005


   ABSTRACT
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Thrombin stimulation of isolated rabbit ventricular myocytes activates a membrane-associated, Ca2+-independent PLA2 (iPLA2) that selectively hydrolyzes plasmalogen phospholipids and results in increased production of arachidonic acid and lysoplasmenylcholine. To determine whether MAPK regulates myocardial iPLA2 activity, we isolated ventricular myocytes from rabbit heart by collagenase digestion and pretreated them with MAPK inhibitors before stimulating them with thrombin. Pretreatment with PD-98059 to inhibit p42/44 MAPK or SB-203580 to inhibit p38 MAPK had no significant effect on thrombin-stimulated, membrane-associated iPLA2 activity. Thrombin stimulation resulted in significant increases in both p42/44 and p38 MAPK activity after 2 min. Pretreatment with the iPLA2-selective inhibitor bromoenol lactone completely inhibited thrombin-stimulated MAPK activity, suggesting that activation of MAPKs was dependent on iPLA2 activation. Ventricular myocyte MAPK activity was increased by incubation of the myocytes with lysoplasmenylcholine, a metabolite produced by iPLA2-catalyzed membrane plasmalogen phospholipid hydrolysis. Altogether, these data suggest that activation of MAPKs occurs downstream of and is dependent on iPLA2 activation in thrombin-stimulated rabbit ventricular myocytes.

lysoplasmenylcholine; cell signaling; protease-activated receptors

PLA2 CATALYZES THE HYDROLYSIS of sn-2 fatty acids from membrane phospholipids, resulting in the production of several biologically active phospholipid metabolites, such as lysophospholipids, arachidonic acid, eicosanoids, and platelet-activating factor (for review, see Refs. 14, 16). In previous studies, we have demonstrated that thrombin stimulation of isolated rabbit ventricular myocytes results in activation of a membrane-associated, Ca2+-independent PLA2 (iPLA2) that selectively hydrolyzes plasmalogen phospholipids, resulting in increased production of plasmenylcholine (LPlsCho) as well as arachidonic acid and prostacyclin release (17).

Studies regarding the regulation of iPLA2 activity suggest that multiple mediators may be involved, including phosphofructokinase (6, 7), CaM (28), ATP (5), and PKC (15, 21, 24). We have demonstrated previously that isolated membrane fractions of rabbit ventricular myocytes that were incubated with either thrombin or PMA to activate PKC demonstrated increased iPLA2 activity, suggesting that a membrane-associated PKC was involved in the activation of iPLA2 (24). Because the membrane fraction was incubated in Ca2+-free medium, we concluded that a novel PKC isoform (Ca2+ independent and diacylglycerol dependent) that is resident in the membrane fraction was responsible for the activation of iPLA2 (24).

Several studies have demonstrated activation of MAPKs after thrombin stimulation (for review, see Ref. 4). Sabri et al. (23) showed that in neonatal rat ventricular myocytes, activation of the protease-activated receptor (PAR)-1 by thrombin results in stimulation of several kinases, including p42/44 and p38 MAPK and JNK. Phosphorylation of the cytosolic, Ca2+-dependent PLA2 (cPLA2) by MAPKs has been demonstrated in a variety of cells (for review, see Refs. 8, 22); however, a similar mechanism for iPLA2 activation has not been explored. Because previous studies have established that activation of both iPLA2 and MAPKs is a consequence of PAR-1 cleavage, the present study was conducted to determine whether the two events occur in the same intracellular signaling pathway in thrombin-stimulated rabbit ventricular myocytes.


   METHODS
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 METHODS
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Cell isolation and culture. Adult rabbits of either sex weighing 2–3 kg were anesthetized with intravenous pentobarbitone sodium (50 mg/kg), and their hearts were removed rapidly. The heart was mounted on a Langendorff apparatus and perfused for 5 min with Tyrode solution containing (in mM) 118 NaCl, 4.8 KCl, 1.2 CaCl2, 1.2 MgCl2, 24 NaHCO3, 1.2 KH2PO4, and 11 glucose. The Tyrode solution was saturated with 95% O2-5% CO2 to yield pH 7.4. This procedure was followed by 4-min perfusion with a Ca2+-free Tyrode solution containing EGTA (100 µM) and a final perfusion for 20 min with Tyrode solution containing 100 µM Ca2+ and 0.033% collagenase. The ventricles were cut into small pieces and shaken in fresh enzyme solution. Individual myocytes were washed with a HEPES buffer containing (in mM) 133.5 NaCl, 4.8 KCl, 1.2 MgCl2, 0.3 CaCl2, 1.2 KH2PO4, 10 glucose, and 10 HEPES, pH 7.4. Extracellular Ca2+ was increased to 1.2 mM in three stages at intervals of 20 min. Myocytes were incubated overnight in medium-199 with 10% FCS at 37°C and then washed three times with 1.2 mM Ca2+-HEPES solution.

Measurement of MAPK activity. p44/42 MAPK (ERK1/2) and p38 MAPK activities were assayed using nonradioactive kits obtained from New England BioLabs (Beverly, MA). Assay of active p44/42 MAPK involves immunoprecipitation of cytosolic activated kinase with an immobilized phospho-p44/42 MAPK MAb, measurement of activity by phosphorylation of the transcription factor Elk-1, and detection of phosphorylated Elk-1 by immunoblotting using an anti-phospho-Elk-1 antibody and densitometric analysis. Assay of active p38 MAPK involves immunoprecipitation with immobilized phospho-p38 MAPK MAb, measurement of activity by phosphorylation of activating transcription factor 2 (ATF2) and detection of phosphorylated ATF2 by immunoblotting using anti-phospho-ATF2 and densitometric analysis.

Immunoblot analysis of p42/44 and p38 MAPKs. Myocytes were suspended in lysis buffer containing (in mM) 20 HEPES (pH 7.6), 250 sucrose, 2 DTT, 2 EDTA, 2 EGTA, 10 beta-glycerophosphate, 1 sodium orthovanadate, 2 PMSF, 20 µg/ml leupeptin, 10 µg/ml aprotinin, and 5 µg/ml pepstatin A (buffer 2). Cells were sonicated on ice for six bursts of 10 s each and centrifuged at 20,000 g at 4°C for 20 min to remove cellular debris and nuclei. Cytosolic and membrane fractions were separated by centrifuging the supernatant at 100,000 g for 60 min. The pellet was resuspended in lysis buffer, and the suspension was centrifuged twice at 100,000 g for 60 min to minimize contamination of the membrane fraction with cytosolic protein. The final pellet was resuspended in lysis buffer containing 0.1% Triton X-100. Protein (cytosol or membrane) was mixed with an equal volume of SDS sample buffer and heated at 95°C for 5 min before being loaded onto a 10% PAGE gel. Protein was separated by SDS-PAGE at 200 V for 35 min and transferred electrophoretically onto polyvinylidene difluoride (PVDF) membranes (Bio-Rad Laboratories, Hercules, CA) at 100 V for 1 h. Nonspecific sites were blocked with Tris-buffered solution containing 0.05% (vol/vol) Tween 20 and 5% (wt/vol) nonfat milk for 1 h at room temperature. The blocked PVDF membrane was incubated with primary antibodies to MAPKs (1 in 1,000 dilution, Cell Signaling Technology, Beverly, MA) horseradish peroxidase-conjugated secondary antibodies. Regions of antibody binding were detected using ECL (Amersham, Arlington Heights, IL) after exposure to Hyperfilm (Amersham). Multiple exposures of the blots to film were developed.

PLA2 activity. Myocytes were suspended in 1 ml of buffer containing (in mM) 250 sucrose, 10 KCl, 10 imidazole, 5 EDTA, 2 DTT, with 10% glycerol, pH 7.8 (buffer 1). The suspension was sonicated on ice six times for 10 s, and the sonicate was centrifuged at 20,000 g for 20 min to remove cellular debris and nuclei. The supernatant was then centrifuged at 100,000 g for 60 min to separate the membrane fraction (i.e., the pellet) from the cytosolic fraction (i.e., the supernatant). The pellet was washed twice to minimize contamination of the membrane fraction with cytosolic protein by resuspension in buffer 1 and centrifugation at 100,000 g for 60 min. The final pellet was resuspended in buffer 1. PLA2 activity in cytosolic and membrane fractions was assessed using an incubating enzyme (8 µg of membrane protein or 200 µg of cytosolic protein) with 100 µM (16:0 dilution; 18:1 dilution of [3H]) plasmenylcholine substrate in assay buffer containing (in mM) 10 Tris, 4 EGTA, 10% glycerol, pH 7.0, at 37°C for 5 min in a total volume of 200 µl. Reactions were terminated by the addition of 100 µl of butanol and released radiolabeled fatty acid was isolated by application of 25 µl of the butanol phase to channeled Silica Gel G plates, development in petroleum ether-diethyl ether-acetic acid (70:30:1 dilution, vol/vol) and subsequent quantification by liquid scintillation spectrometry. Protein content of each sample was determined using the Lowry method with freeze-dried BSA as the protein standard (13).

Statistical analysis. Statistical comparison between multiple values was performed using ANOVA. All results are expressed as means ± SE. Statistical significance was set at P < 0.05.


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Because we demonstrated previously that thrombin stimulation of isolated membrane fractions from isolated rabbit ventricular myocytes results in iPLA2 activation, we concluded that the signaling sequence from activation of PAR-1 by thrombin to activation of iPLA2 was contained entirely within the membrane (24). To determine whether MAPKs are involved in iPLA2 activation after thrombin stimulation, we first identified the subcellular localization of MAPKs in ventricular myocytes. Immunoblot analysis of myocyte cytosolic and membrane fractions demonstrated the presence of both total and phosphorylated p44/42 and p38 MAPKs almost exclusively in the cytosolic fraction (Fig. 1), with little signal present in the membrane fractions. The cytosolic localization suggests that MAPKs may not be required for membrane-associated iPLA2 activation. Stimulation of isolated ventricular myocytes with thrombin (0.05 IU/ml for 2 min) did not result in detectable translocation of MAPKs from the cytosolic to the membrane fraction (Fig. 1).


Figure 1
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  		Fig. 1. Immunoblot analysis of the localization of total and phosphorylated (circled P) p44/42 MAPK and p38 MAPK in the cytosolic (cyt) and membrane (mem) subcellular fractions isolated from rabbit ventricular myocytes under unstimulated (A) or thrombin-stimulated conditions (0.05 IU/ml, 2 min) (B). Proteins were separated by SDS-PAGE and transferred onto polyvinylidene difluoride membranes. Membranes were probed with anti-p44/42 or anti-p38 MAPK antibodies (1:1,000 dilution) and incubated with horseradish peroxidase-linked secondary antibodies (1:50,000 dilution). Immunoblots were detected using ECL and exposure to film for 5 min. Shown are immunoblots representative of 6 separate myocyte isolations. Phosphorylated forms of corresponding kinases: p44, total p44 MAPK; p42, total p42 MAPK, p38, total p38 MAPK.

 
To determine whether MAPKs are activated in thrombin-stimulated ventricular myocytes, we measured MAPK activity after thrombin stimulation with or without SB-203580 pretreatment to inhibit p38 MAPK and with or without PD-98059 pretreatment to inhibit p44/42 MAPK (Fig. 2). Thrombin stimulation of isolated ventricular myocytes resulted in significant increases in both p44/42 MAPK (Fig. 2A) and p38 MAPK (Fig. 2B) activities that were sustained for 20 min after 2 min of stimulation. Because activation of iPLA2 is apparent after 1 min of thrombin stimulation (17), the time course of thrombin-induced MAPK activation appears to lag behind that of iPLA2. Pretreatment with PD-98059 significantly inhibited thrombin-stimulated p44/42 MAPK activity (Fig. 2A) but did not have any effect on p38 MAPK activity (Fig. 2B). Conversely, pretreatment with SB-203580 completely inhibited the increase in p38 MAPK activity observed after thrombin stimulation but did not affect p44/42 MAPK activity.


Figure 2
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  		Fig. 2. p44/42 MAPK (A) and p38 MAPK (B) activities measured after stimulation of isolated ventricular myocytes by thrombin (0.05 IU/ml; bullet). Pretreatment of ventricular myocytes with PD-98059 (5 µM, 10 min; {square}) completely inhibited thrombin-stimulated p44/42 MAPK activity, whereas pretreatment with SB-203580 (1 µM, 10 min; {triangleup}) had no effect (A). Conversely, pretreatment with SB-203580 completely inhibited thrombin-stimulated p38 MAPK activity, whereas pretreatment with PD-98059 had no effect (B). Data shown represent means ± SE of results from 6 different myocyte isolations. *P < 0.05, **P < 0.01 vs. unstimulated values.

 
To test whether active MAPK is necessary for iPLA2 activation, we pretreated isolated ventricular myocytes with SB-203580 or PD-98059 before thrombin stimulation and measured iPLA2 activity. As shown in Fig. 3, neither SB-203580 nor PD-98059 had a significant inhibitory effect. Thus MAPKs do not appear to regulate thrombin-stimulated, membrane-associated iPLA2 activity in isolated ventricular myocytes. Furthermore, pretreatment with bromoenol lactone (BEL), a selective inhibitor of iPLA2, resulted in complete inhibition of thrombin-stimulated MAPK activation, suggesting that iPLA2 is an upstream regulator of MAPK activation in ventricular myocytes (Fig. 4).


Figure 3
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  		Fig. 3. Pretreatment of ventricular myocytes with PD-98059 (5 µM, 10 min; hatched bars) or SB-203580 (1 µM, 10 min; open bars) has no significant effect on thrombin (0.05 IU/ml, 2 min; solid bars)-stimulated, membrane-associated, Ca2+-independent PLA2 (iPLA2) activity (4 mM EGTA) using (16:0 dilution; [3H], 18:1 dilution) plasmenylcholine substrate. Data shown represent the mean ± SE for results from 6 different myocytes isolations. **P < 0.05, thrombin-stimulated values vs. corresponding controls.

 

Figure 4
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  		Fig. 4. p44/42 MAPK (A) and p38 MAPK (B) activities measured after stimulation of isolated ventricular myocytes by thrombin (0.05 IU/ml; bullet) are completely inhibited by 5 µM bromoenol lactone (BEL) pretreatment (10 min; x). Data shown represent means ± SE of results from 6 different myocytes isolations. *P < 0.05, **P < 0.01 vs. unstimulated values.

 
To determine whether activation of MAPKs in thrombin-stimulated ventricular myocytes is mediated by one or both of the metabolites of iPLA2-catalyzed membrane plasmalogen hydrolysis, we incubated isolated ventricular myocytes with LPlsCho or arachidonic acid. Figure 5 shows that a significant increase in both p44/42 MAPK and p38 MAPK activities was observed after 5-min incubation with LPlsCho and returned to normal after 30 min. We did not observe a similar activation of MAPKs when ventricular myocytes were incubated with 5 µM arachidonic acid for up to 30 min (data not shown).


Figure 5
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  		Fig. 5. Incubation of ventricular myocytes with lysoplasmenylcholine (LPlsCho, 5 µM) results in significant activation of p44/42 MAPK ({square}) and p38 MAPK (bullet). Data shown represent means ± SE of results from 6 different myocyte isolations. *P < 0.05, **P < 0.01 vs. control values.

 
Altogether, these data suggest that activation of MAPKs is dependent on the production of LPlsCho and does not regulate membrane-associated iPLA2 activity in thrombin-stimulated isolated rabbit ventricular myocytes. Our results also suggest the intriguing possibility that in thrombin-stimulated rabbit ventricular myocytes, activation of MAPKs occurs downstream of, and is dependent on, iPLA2 activation.


   DISCUSSION
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 DISCUSSION
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PLA2-catalyzed hydrolysis of membrane phospholipids results in the stoichiometric production of a free fatty acid (most notably arachidonic acid) and a lysophospholipid. Both of these metabolites exert profound effects on membrane properties directly or can serve as precursors for biologically active metabolites such as eicosanoids or platelet-activating factor. PLA2 isoforms are classified into three main types: secretory (sPLA2), cytosolic Ca2+-activated (cPLA2), and iPLA2. All three types have been identified in the myocardium (14, 16), but several studies have suggested that the majority of myocardial PLA2 activity is Ca2+ independent and membrane associated.

To date, the iPLA2 isoforms that contribute to membrane-associated iPLA2 activity in isolated ventricular myocytes have not been identified; however, there are several possible candidates. The iPLA2 that has been isolated from the cytosol of P388D1 macrophages (iPLA2-beta) is present almost exclusively in membrane factions isolated from the heart (10, 18–20). In addition, a novel, membrane-associated iPLA2 (iPLA2-{gamma}) has been identified in the heart (12). This isoform is inhibited by BEL and is tightly bound to the membrane fraction (12). Sequence homology between the two iPLA2 isoforms is confined to the ATP binding motif, the serine lipase site, and a region of nine amino acids that have no known functional significance.

Studies regarding the regulation of myocardial iPLA2 suggest that multiple mechanisms are involved. To date, little information has been produced regarding signal transduction pathways and intracellular changes that occur in isolated cardiac myocytes linking stimulation of thrombin receptors to the activation of iPLA2. Analysis of the amino acid sequences for rabbit ventricular myocyte iPLA2-beta and iPLA2-{gamma} (GenBank accession nos. AY-744674 and AY-738591, respectively) reveals several putative MAPK consensus phosphorylation sites (SP/TP). We previously determined (24) that membrane-associated iPLA2 activity is dependent on a membrane-associated novel PKC isoform and that the entire signaling pathway between PAR-1 and iPLA2 resides in the membrane fraction. This effectively eliminates the possibility of a requirement for cytosolic kinases in the regulation of myocardial membrane-associated iPLA2. Our data support this observation and indicate that activation of MAPKs is a consequence of iPLA2 activity in thrombin-stimulated ventricular myocytes.

MAPKs have been shown to regulate both cPLA2 and sPLA2 in the myocardium. For example, Magne et al. (11) demonstrated that beta2-adrenergic receptor stimulation is coupled to myocardial cPLA2 by a MAPK-dependent pathway. Activation of myocardial cPLA2 occurs via phosphorylation of the enzyme together with an increase in intracellular Ca2+, which drive its translocation from cytosol to membrane localization (11). In addition, p38 MAPK activation is necessary for interleukin-1beta-induced synthesis and release of Group IIa sPLA2 in cardiac myocytes (3). Clearly, there is evidence of MAPK activation of sPLA2 and cPLA2 in cardiac myocytes, but there is no indication of its involvement in the activation of iPLA2. Because MAPKs are localized in the cytosol, it would be expected that they might regulate the activity of other cytosolic enzymes such as cPLA2 and sPLA2. However, the majority of iPLA2 activity in ventricular myocytes is membrane associated and therefore may not be accessible for phosphorylation by a cytosolic kinase. Activation of iPLA2 and MAPKs in thrombin-stimulated myocytes occurs within 2 min. It is possible that an event downstream from MAPK activation is the sequential activation of sPLA2 or cPLA2 in the myocardium, representing a dependence of these PLA2 isoforms on iPLA2 activation in thrombin-stimulated cardiac myocytes.

We (17) have previously shown that thrombin activates a membrane-associated iPLA2 that results in accelerated membrane plasmalogen hydrolysis, leading to increased arachidonic acid and LPlsCho production. Accumulation of LPlsCho in ischemic cardiac myocytes may initiate arrhythmogenesis because it has been demonstrated to alter the function of integral membrane proteins directly at concentrations lower than those required for structurally similar amphiphilic metabolites (9). We have also shown that LPlsCho increases the L-type Ca2+ channel current, intracellular free Ca2+, and myofilament Ca2+ sensitivity in normoxic cardiac myocytes (9). These changes lead to prolonged action potential duration and augmented cardiac myocyte contractility and are often followed by early or delayed afterdepolarization and sustained membrane depolarization (9).

In addition to its potential arrhythmogenic properties, LPlsCho has also been shown to activate myocardial cAMP-dependent protein kinase (26) and to increase cAMP response element binding protein phosphorylation and c-fos expression (27). Activation of MAPKs by LPlsCho has not been demonstrated previously. However, activation of MAPK pathways occurs in myocardial ischemia (for review, see Refs. 1 and 2) and may be a result, at least in part, of iPLA2 activation and LPlsCho accumulation. Activation of MAPKs plays a key role in the pathogenesis of several cardiovascular diseases, including ischemic and reperfusion injury, myocardial hypertrophy, and heart failure. A previous study using rat neonatal ventricular myocytes demonstrated that PAR-1 and PAR-2 activated JNK, p38 MAPK, and AKT and promoted hypertrophy (23). In addition, the isoform specificity of two p38 MAPKs has been described after their overexpression, with p38-beta mediating cardiac hypertrophy and p38-{alpha} mediating apoptotic cell death (25). In this study, we did not use p38 antibodies that were isoform specific; therefore, no conclusion can be drawn regarding which ventricular myocytes isoforms are activated by thrombin. Altogether, these data suggest that activation of MAPKs may be important in cardiac myocyte cell survival and growth in the border regions of the ischemic myocardium.

Altogether, these results demonstrate that thrombin stimulation of rabbit ventricular myocyte iPLA2 may participate in the progression of long-term cardiovascular disease or remodeling via activation of MAPKs.


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This work was funded by National Heart, Lung, and Blood Institute Grant HL-68588.


   FOOTNOTES
 

Address for reprint requests and other correspondence: J. McHowat, Dept. of Pathology, St. Louis Univ. School of Medicine, 1402 S. Grand Blvd., St. Louis, MO 63104 (e-mail: jane.mchowat{at}tenethealth.com)

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.


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