Arachidonic Acid Drives Mini-glucagon Action in Cardiac Cells*

(Received for publication, July 8, 1996, and in revised form, February 27, 1997)

Anne Sauvadet , Troy Rohn , Françoise Pecker and Catherine Pavoine

From INSERM Unité 99, Hôpital Henri Mondor, 94010 Créteil, France

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES


ABSTRACT

Recent studies have shown that glucagon is processed by cardiac cells into its COOH-terminal (19-29) fragment, mini-glucagon, and that this metabolite is an essential component of the contractile positive inotropic effect of glucagon (Sauvadet, A., Rohn, T., Pecker, F. and Pavoine, C. (1996) Circ. Res. 78, 102-109). We now show that mini-glucagon triggers arachidonic acid (AA) release from [3H]AA-loaded embryonic chick ventricular myocytes via the activation of a phospholipase A2 sensitive to submicromolar Ca2+ concentrations. The phospholipase A2 inhibitor, AACOCF3, prevented mini-glucagon-induced [45Ca2+] accumulation into the sarcoplasmic reticulum, but inhibitors of lipoxygenase, cyclooxygenase, or epoxygenase pathways were ineffective. AA applied exogenously, at 0.3 µM, reproduced the effects of mini-glucagon on Ca2+ homeostasis and contraction. Thus AA: (i) caused [45Ca2+] accumulation into a sarcoplasmic reticulum compartment sensitive to caffeine; 2) potentiated caffeine-induced Ca2+ mobilization from cells loaded with Fura-2; 3) acted synergistically with glucagon or cAMP to increase both the amplitude of Ca2+ transients and contraction of electrically stimulated cells. AA action was dose-dependent and specific since it was mimicked by its non-hydrolyzable analog 5,8,11,14-eicosatetraynoic acid but not reproduced by other lipids such as, arachidic acid, linolenic acid, cis-5,8,11,14,17-eicosapentaenoic acid, cis-4,7,10,13,16,19-docosahexaenoic acid, or arachidonyl-CoA, even in the micromolar range. We conclude that AA drives mini-glucagon action in the heart and that the positive inotropic effect of glucagon on heart contraction relies on both second messengers, cAMP and AA.


INTRODUCTION

Data from the past few years have shed a new light on the physiological pathways of glucagon action (for review, see Ref. 1). Thus, the interaction of glucagon with cardiac tissue leads to its processing by a specific ectoendopeptidase and to the liberation of the COOH-terminal (19-29) fragment, mini-glucagon. This metabolite plays a key role in the positive inotropic effect of the hormone (2-5) since glucagon alone, under minimal degradation conditions, has no effect on heart cell contraction (5). We have shown that both peptides act synergistically on Ca2+ cycling in heart cells (6). Mini-glucagon and glucagon actions can be summarized as the ability of the former to accumulate Ca2+ into sarcoplasmic reticulum stores (SR stores)1 and that of the latter to induce Ca2+-induced-Ca2+-release from the same stores (6). Glucagon action is mediated by cAMP produced from either stimulation of adenylyl cyclase or inhibition of the cyclic GMP-inhibited phosphodiesterase (CGI-PDE or PDE III), depending on the species (6-8). In contrast, the action of mini-glucagon does not rely on classical transduction pathways. In fact, mini-glucagon does not evoke any detectable change in either cAMP or cGMP or inositol 1,4,5-trisphosphate production and its second messenger remained to be identified (5, 6).

Recently, considerable evidence has accumulated to suggest a role for arachidonic acid (AA) and/or its oxidized metabolites in signal transduction processes (9). AA release in response to cell receptor activation is considered as a key step in the positive inotropic response of angiotensin II, bradykinin, and endothelin (10-13).

Arachidonic acid is stored in esterified form in cell membrane phospholipids, from which it can be liberated through multiple enzymatic pathways (for review, see Ref. 14). PLA2 catalyzes the hydrolysis of phospholipids at the sn-2 position. Therefore, this enzyme can release arachidonate in a single step reaction. By contrast, PLC and PLD do not release free arachidonic acid directly, but generate lipid products containing arachidonate (diacylglycerol and phosphatidic acid, respectively), which can be released subsequently by diacylglycerol and monoacylglycerol lipases. Once released, free arachidonate may diffuse out of the cell, be reincorporated into phospholipids, or metabolized. In addition, several reports have focused on a signaling role for AA in heart cells. One of the most thoroughly characterized targets of AA is protein kinase C (15), and AA has also been reported to modulate sarcolemmal ion channels, including K+ channels (10, 16, 17)), and voltage-dependent Ca2+ channels (18), as well as to increase Ca2+ release from the sarcoplasmic reticulum in heart (19).

Several hormones which exert positive inotropic responses evoke release of AA. Thus, in neonatal rat ventricular myocytes, angiotensin II stimulates AA release via both PLC and PLA2 activation through AT1 and AT2 receptors, respectively (11). In isolated hearts, bradykinin and endothelin, which also activate PLC, stimulate release of AA (12, 13). Thus, in addition to representing an important signaling molecule under pathological circumstances, such as ischemia (20), AA, in the normal physiological setting may modulate important steps in excitation-contraction coupling.

The aim of the present study was to evaluate the role of AA in mediating the actions of mini-glucagon on Ca2+ homeostasis and cell contraction in embryonic chick heart cells. The main findings are that mini-glucagon triggers a Ca2+-dependent release of AA and that AA added exogenously, at a submicromolar concentration (0.3 µM), mimics mini-glucagon actions. Our data suggest that free AA, and not products of AA metabolism, is the primary mediator of mini-glucagon action in ventricular myocytes.


EXPERIMENTAL PROCEDURES

Materials

Mini-glucagon was obtained from ICN (Orsay, France), and glucagon from Novo Nordisk Laboratories (Bagsvaerd, Denmark). Penicillin-streptomycin antibiotic solution, trypsin, nucleotides, bovine serum albumin, arachidonic acid, cis-4,7,10,13,16,19-docosahexaenoic acid, and cis-5,8,11,14,17-eicosapentaenoic acid were purchased from Sigma (Saint Quentin Fallavier, France). AACOCF3 and ETYA were from Biomol (Plymouth Meeting, PA). Fura-2, Fura-2/AM were from Molecular Probes (Interchim, Montluçon, France). Fetal calf serum was from Life Technologies, Inc. (Cergy Pontoise, France). Phosphate-buffered saline 2040 and M199 media were obtained from Eurobio (Les Ulis, France). Digitonin from Merck has been recrystallized and stored as a 8 mg/ml solution in Me2SO. [45Ca2+] (10-40 mCi/mmol) was from ICN (Orsay, France). [5,6,8,9,11,12,14,15-3H]Arachidonic acid (180-240 Ci/mmol) was from DuPont NEN (Les Ulis, France). Antibodies from rabbit anti-mouse IgG-conjugated peroxidase were purchased from BIOSYS (Compiègne, France). Mouse Monoclonal antibodies against human cytoplasmic phospholipase A2 were from Santa Cruz.

Methods

Primary Culture of Chick Embryo Ventricular Cells

Fecundated eggs were obtained from the Haas farm (Kaltenhouse, France). Primary monolayer cultured heart cells were prepared from 13-day-old chick embryo ventricles as described previously (6, 21). Briefly, cells were dissociated by repeated cycles of trypsinization. The resulting cell suspension (5-7 × 105 cells/ml) was bubbled with 5% CO2, 95% air, at 4 °C, and kept in buffer A (M199 medium containing 0.1% (w/v) NaHCO3, 0.01% (w/v) L-glutamine, 0.1% penicillin-streptomycin antibiotic solution) until used, up to 5 days.

Rat Ventricular Myocytes

Ventricular myocytes were enzymatically dispersed from adult male Wistar rat (250-300 g) as described previously (7).

Measurements of [45Ca] Accumulation into Intracellular Compartments

Myocytes (5 × 105 cells/ml), suspended in buffer A and supplemented with 5% (v/v) fetal calf serum, were plated on glass coverslips in multiwell plates and kept at 37 °C in humidified 5% CO2, 95% air for 24 h, as described by Sauvadet et al. (6, 21). Cells were then washed in 2 × 1 ml of saline buffer B (10 mM glucose, 130 mM NaCl, 5 mM KCl, 10 mM Hepes buffered at pH 7.4 with Tris base, 1 mM MgCl2, 2 mM CaCl2). At time 0 of the experiment, the cells were immersed in 1 ml of saline buffer B, containing 2 mM [45Ca] (5 µCi/ml) in the presence or absence of various peptides and with or without enzymatic inhibitors. After various periods of incubation, the cells were washed two times for 10 s in 10 ml of Ca2+-free saline buffer B at 25 °C. As described previously in Ref. 6, to determine [45Ca] accumulation into intracellular stores, cells were then subjected to digitonin lysis, which selectively disrupts the sarcolemmal membranes. This procedure, previously described by Altschuld (22), consisted of an incubation for 45 s at 25 °C in Ca-free saline buffer B, supplemented with: 0.1 mM EGTA, 10 mM MgCl2, 10 mM ATP, 5 µM ruthenium red and digitonin (16 µg/mg of protein). Mg-ATP was added to protect against hypercontracture and ruthenium red to block Ca2+ efflux from the SR. Intracellular Ca2+ accumulation was estimated from [45Ca] recovered in digitonin-resistant structures attached to coverslips after the addition of 0.2 M NaOH for 2 h at room temperature. Samples were diluted in 10 ml of Beckman Ready Safe and counted in a scintillation counter. Data are expressed as mean ± S.E. To determine significant differences from control values, results were analyzed by employing the Student's t test.

Fura-2 Loading and Ca2+ Imaging

Cells were plated on plastic dishes, the bottom of which was replaced by a glass coverslip coated with laminin (1 µg/ml), and were incubated at 37 °C in humidified 5% CO2, 95% air for 17-24 h.

Cells, attached to laminin, were bathed in 2 ml of saline buffer B and incubated for 20 min at 25 °C with 1.5 µM Fura-2/AM (3 µl of 1 mM Fura-2/AM in Me2SO), in the presence of 1 mg/ml bovine serum albumin to improve Fura-2 dispersion and facilitate cell loading. Cells were then washed with saline buffer B (2 × 2 ml) and allowed to incubate in the same buffer for 15 min at 25 °C to facilitate hydrolysis of intracellular Fura-2/AM. The concentration of Fura-2 in myocytes was estimated as described previously (6, 21), according to the procedure of Donnadieu et al. (23). Under usual loading conditions, the average intracellular concentration of Fura-2 was 15 µM.

Ca2+ imaging, developed by A. Trautmann in collaboration with the IMSTAR CO (Paris, France), was essentially as described by Sauvadet et al. (6, 21). All tracings of fluorescence ratio are representative of at least 10 cells, and were performed on at least two different cell isolations. Imaging studies were performed on cells in which no spontaneous rise in [Ca2+]i was observed prior to experimental manipulation.

Field electrical stimulation (square waves, 10-ms duration, amplitude 20% above threshold, 0.5 Hz) was supplied through a pair of platinum electrodes connected to the output of a HAMEG stimulator (Paris, France). Cells were perifused with saline buffer B containing 1.27 mM CaCl2 and stimulated until a steady-state level of the Ca2+ transients was achieved, before each protocol, as described previously by Bassani (24). To evaluate the caffeine-releasable [Ca2+] pool, drugs and peptides were added to the perfusion medium a few seconds after interruption of electrical stimulation. In a second series of experiments, performed to measure variations in the amplitude of Ca2+ transients, electrical stimulation of the cells was maintained throughout the experiment. Drugs and peptides were added to the perfusion medium at time 0. To evaluate the effect of glucagon, mini-glucagon, 8-Br-cAMP, and/or arachidonic acid quantitatively, we used two parameters: the percentage increase in diastolic Fura-2 ratio (360:380) and the percentage increase in the amplitude of Ca2+ transient.

Contractility Measurements

Experiments were performed in conditions similar to Ca2+ imaging, but cells were illuminated with visible light and images transmitted through a solid-state camera (CCD, black and white, 0.847 cm high sensitivity) connected to the sideport of the microscope. Contractions of single stimulated (0.5 Hz) myocytes were displayed on a video monitor and the corresponding images (pixel × pixel) were recorded at a frequency of 9/s. Contractility measurements were determined by assessing changes in cell length using the Morphostar II software, developed by the IMSTAR CO (Paris, France). All tracings of cell length are representative of at least 5 cells and performed on at least two different cell isolations.

[3H]Arachidonic Acid Labeling

Embryonic chick ventricular myocytes (5 × 105 cells/ml), suspended in buffer A, were plated in multiwell plates and incubated with 1.5 µCi/ml [3H]AA (6.75 nM) in humidified 5% CO2, 95% air, at 37 °C. After 24 h, the cells were washed twice in saline buffer B containing 0.2% fatty acid-free bovine serum albumin and resuspended in saline buffer B.

Measurements of [3H]Arachidonic Acid Release in Intact Cells

At time 0 of the experiment, [3H]AA-labeled cells were exposed to various peptides and/or enzymatic inhibitors, and incubated for various periods of time at 37 °C. Incubation was terminated by the addition of ice-cold EGTA (2 mM final), and the media were immediately transferred to microcentrifuge tubes. Centrifugation at 17,600 × g for 20 min in a Sigma centrifuge (model 2K15) at 4 °C was performed to pellet any cells or debris inadvertently collected with the extracellular medium. The amount of radioactivity in the supernatant was quantitated by liquid scintillation counting.

Analysis of the lipids released in the incubation medium was performed as described in Ref. 25. At the end of the incubation period, the reaction mixture was acidified to pH 3.0 with HCl and the products were extracted twice with ethyl acetate. The dried extracts were dissolved in ethanol/chloroform (1:2, v/v) and chromatographed on silica gel thin layer plate (Whatman LK5) in ethyl acetate/isooctane/water/acetic acid (11:5:10:2, v/v) as the solvent system. Standard concentrations of AA, prostaglandin Es, and hydroxyeicosatetraenoic acids were co-chromatographed and visualized by exposing the plates to ultraviolet light. The area corresponding to each visualized spot was carefully extracted and the radioactivity was determined by liquid scintillation counting.

Measurements of [3H]Arachidonic Acid Release on Cells Permeabilized with Digitonin

[3H]AA-labeled cells were incubated for 10 min at 37 °C in the presence of 0.1 nM mini-glucagon or vehicle. Cells were next subjected to digitonin lysis for 45 s at 25 °C, in Ca2+-free saline buffer B, supplemented with 0.1 mM EGTA, 10 mM MgCl2, 10 mM ATP, 5 µM ruthenium red and digitonin (16 µg/mg of protein). At time 0 of the experiment, the cells were exposed to 0.1 nM mini-glucagon or vehicle and incubated for 30 min at 37 °C in saline buffer B containing 2 mM CaCl2, 10 mM MgCl2, 10 mM ATP, 5 µM ruthenium red and various concentrations of EGTA, ranging from 3.307 to 2.131 mM. These corresponded to [Ca2+]i ranging from 113 to 1193 nM, as calculated by using the EQUIV program (Thierry Capiod, INSERM Unité 274), and further determined in parallel experiments using cells permeabilized with digitonin and loaded with Fura-2. Incubation was terminated by the addition of ice-cold EGTA (2 mM final) and the media were transferred to microcentrifuge tubes. Centrifugation at 17,600 × g for 20 min in a Sigma centrifuge (model 2K15) at 4 °C was performed to pellet any cells or debris inadvertently collected with the extracellular medium. The amount of radioactivity in the supernatant was quantitated by standard liquid scintillation procedures.

Immunoblot Analysis of cPLA2

Embryonic chick ventricular cells or rat ventricular myocytes were disrupted by sonication in buffer C (40 mM Tris-HCl, pH 7.4, 1 mM EDTA, 0.25 M saccharose, 1 mM phenylmethylsulfonyl fluoride, 1 µg/ml leupeptin, 50 mM sodium fluoride, 100 µM sodium orthovanadate). After dilution (v/v) with sample buffer (250 mM Tris-HCl, pH 6.8, 8% SDS, 20% glycerol, 10% beta -mercaptoethanol), samples were boiled for 5 min, and the proteins (25 µg/lane) were separated on 7.5% SDS-polyacrylamide gel electrophoresis (25 mA/gel). The proteins were transferred to Hybond C super nitrocellulose membrane (Amersham), by electroblot using Tris glycine buffer containing 20% methanol. The transfer was performed at 90 mA for 60 min. Protein transfer was evaluated by staining the gel with Coomassie Blue. The nitrocellulose blots were agitated for 1 h at room temperature in TBST buffer (10 mM Tris-HCl, pH 8, 150 mM NaCl, 0.15% Tween 20) supplemented with 5% nonfat dry milk, washed three times with TBST, and incubated with primary monoclonal antibodies against cPLA2 (1:500 dilution in TBST supplemented with 5% nonfat dry milk) for 180 min. Membranes were washed three times with TBST and incubated with peroxidase-conjugated rabbit anti-mouse IgG (1:1000 dilution) for 90 min at room temperature in TBST containing 5% nonfat dry milk. Membranes were washed three times with TBST and the peroxidase activity was determined using the enhanced chemiluminescence Western blotting detection system (ECL (Amersham Corp.)).


RESULTS

Mini-glucagon Stimulates Arachidonic Acid Release from Embryonic Chick Ventricular Myocytes

The action of mini-glucagon on AA release was assessed on embryonic chick ventricular myocytes labeled for 24 h with [3H]AA before the addition of mini-glucagon, to allow steady state labeling of the cellular AA pool. As shown in Fig. 1A, 0.1 nM mini-glucagon evoked a sustained release of AA from cells over the 30-min period examined. Under similar experimental conditions, the level of AA release in the absence of mini-glucagon remained constant (Fig. 1A). The action of mini-glucagon was dose-dependent, with a maximal (164 ± 7%) increase of [3H]AA released after 30 min observed at 0.1 nM mini-glucagon and a half-maximal response occurring at 0.01 nM mini-glucagon (Fig. 1B). Under the same experimental conditions, angiotensin II, at 0.3 µM, evoked a similar increase (173 ± 14%) in [3H]AA release (Fig. 2). The action of mini-glucagon was specific since glucagon itself did not affect [3H]AA release significantly, even at micromolar concentrations (Fig. 1B, inset).


Fig. 1. Time course of mini-glucagon-stimulated [3H]AA release from embryonic chick ventricular cells (A) and dose-response and specificity of mini-glucagon action (B). Embryonic chick heart cells were labeled with 1.5 µCi/ml [3H]AA as described under "Experimental Procedures." Radiolabeled cells were washed twice in saline buffer containing 0.2% fatty acid-free bovine serum albumin and incubated for various periods of time in the presence or absence of 0.1 nM mini-glucagon (Fig. 1A) or for 30 min in the presence of increasing concentrations of mini-glucagon (Fig. 1B). The amount of [3H]AA released was expressed as percentage of control values which remained constant over the 30-min period that was examined (60 ± 6 disintegrations/min/µg of protein, A; 134 ± 9 disintegrations/min/µg of protein, B). The maximum [3H]AA released after 30 min never exceeded 10% of the total radioactivity taken up by the cells. Values are means ± S.E. of three (A), eight (B, mini-glucagon), and three (B, glucagon) different experiments, done in triplicate.
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Fig. 2. Mini-glucagon-induced [3H] AA release relies on PLA2 activation. Embryonic chick ventricular cells were labeled for 24 h with 1.5 µCi/ml [3H]AA as described under "Experimental Procedures." Radiolabeled cells were washed twice in saline buffer containing 0.2% fatty acid-free bovine serum albumin and incubated for 30 min in the presence or absence of various peptides (mini-glucagon, angiotensin, or endothelin) and with or without a PLA2 inhibitor, quinacrine or AACOCF3. The amount of [3H]AA released was expressed as percentage of control value (zero time: 148 ± 7 disintegrations/min/µg of protein). Values are means ± S.E. of two different experiments done in triplicate.
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It was important to determine if the 3H-labeled material that was released was authentic AA. Accordingly, cell supernatants were analyzed by extraction and resolution on thin layer chromatography (TLC). As shown in Table I, the 3H-labeled material in the supernatants of control and mini-glucagon-treated cells was mainly identified as [3H]AA (67 and 54%, respectively) but also consisted of [3H]lipoxygenase products (11 and 24%, respectively). No significant conversion of AA into cyclooxygenase or unidentified products could be detected in addition to the nonenzymatic degradation or contaminants of standard [3H]AA (Table I).

Table I. Identification of 3H-metabolites in supernatants of cells prelabeled with [3H]AA

Embryonic chick heart cells were labeled with 1.5 µCi/ml [3H]AA as described under "Experimental Procedures." After two washings in saline buffer containing 0.2% fatty acid free bovine serum albumin, [3H]AA-labeled cells were incubated for 30 min in the presence or absence of 0.1 nM mini-glucagon, and with or without a mixture of inhibitors of AA metabolism including: 30 µM NDGA, 1 µM indomethacin, and 100 µM SKF 250A. Analysis of the 3H-lipids released in the incubation medium was performed following extraction and chromatography on silica gel thin layer plate (TLC plate) as described under "Experimental Procedures." Results, corrected for yield of extraction, are expressed in disintegrations/min of 3H-product/104 cells and in % of total radioactivity recovered in the migration lane. Each sample represents the pool of quadruplicates. Data are from a typical experiment which has been repeated twice. Standard [3H]AA was migrated in parallel in order to determine the nonenzymatic breakdown of AA. Embryonic chick heart cells were labeled with 1.5 µCi/ml [3H]AA as described under "Experimental Procedures." After two washings in saline buffer containing 0.2% fatty acid free bovine serum albumin, [3H]AA-labeled cells were incubated for 30 min in the presence or absence of 0.1 nM mini-glucagon, and with or without a mixture of inhibitors of AA metabolism including: 30 µM NDGA, 1 µM indomethacin, and 100 µM SKF 250A. Analysis of the 3H-lipids released in the incubation medium was performed following extraction and chromatography on silica gel thin layer plate (TLC plate) as described under "Experimental Procedures." Results, corrected for yield of extraction, are expressed in disintegrations/min of 3H-product/104 cells and in % of total radioactivity recovered in the migration lane. Each sample represents the pool of quadruplicates. Data are from a typical experiment which has been repeated twice. Standard [3H]AA was migrated in parallel in order to determine the nonenzymatic breakdown of AA.
Metabolites Basal conditions
Inhibitors of AA metabolism
Migration of standard [3H]AA
Control Mini-glucagon Control Mini-glucagon

dpm (%) dpm (%) %
Arachidonic acid 2,950 (67) 4,754 (54) 8,879 (84) 20,388 (81) 81
Lipoxygenase products (HETE5; HETE13; HETE15) 484 (11) 2,113 (24) 423 (4) 1258 (5) 7
Cyclooxygenase products (PGE1; PGE2) 352 (8) 440 (5) 211 (2) 252 (1) 5
Nonidentified products 176 (4) 1,497 (17) 1,057 (10) 3,272 (9) 7

Evidence Supporting PLA2 Activation in Mini-glucagon Stimulated AA Release

In heart, AA formation is essentially due to PLA2 stimulation but may also occur upon activation of either PLC or PLD. The next series of experiments were performed to examine whether the release of AA elicited by mini-glucagon relied on PLA2 activation. Results shown in Fig. 2 indicated that the addition of PLA2 inhibitors, quinacrine, or the analogue of AA, AACOCF3, resulted in complete inhibition of mini-glucagon promoted release of AA (Fig. 2). Quinacrine and AACOCF3, also blocked angiotensin II-induced AA release. In contrast, these PLA2 inhibitors were partially effective against endothelin elicited AA release (Fig. 2), suggesting that part of release of AA derived from the PLC pathway (26).

The hormone-activated PLA2 (cPLA2) is considered to be a Ca2+-dependent enzyme. The Ca2+ dependence of mini-glucagon-stimulated AA release was tested by two ways. First, in intact cells, lowering the extracellular free Ca2+ concentration ([Ca2+]e), from 2 mM to 500 µM, considerably reduced mini-glucagon-stimulated AA release, while basal AA release remained unchanged (Fig. 3). The presence of Ca2+ in the extracellular medium was therefore a requirement for mini-glucagon action. Second, we examined AA release under controlled free Ca2+ concentration conditions. Myocytes were permeabilized with digitonin and incubated in Ca2+/EGTA buffers with varying free Ca2+. Those cells were still responsive to mini-glucagon, however, substantial variations in the degree of stimulation by mini-glucagon were observed. These variations could be overcome if cells were preincubated for 10 min with mini-glucagon before lysis with digitonin and if micromolar concentrations of GTP were added to the assay medium. This GTP requirement for the activation of AA release by mini-glucagon would suggest the involvement of a G protein. As shown in the Fig. 3B, under these conditions, a highly reproducible increase in arachidonic acid release was observed at 700 nM [Ca2+]i with 0.1 nM mini-glucagon in the presence of 10 µM GTP. The half-maximal stimulation occurred at 450 nM [Ca2+]i. It is noteworthy that in the absence of mini-glucagon, the basal release of AA had a similar Ca2+ dependence (Fig. 3B), but was not improved by the additional presence of 10 µM GTP in the incubation medium (not shown). Taken together these data are consistent with the mini-glucagon activation of a Ca2+-dependent PLA2.


Fig. 3. Ca2+-dependent stimulation of AA release by mini-glucagon: dependence on [Ca2+]e in intact cells (A) and dependence on [Ca2+]i in digitonin-permeabilized cells (B). Embryonic chick heart cells were labeled for 24 h with 1.5 µCi/ml [3H]AA as described under "Experimental Procedures." Radiolabeled cells were washed twice at 37 °C in saline buffer containing 0.2% fatty acid-free bovine serum albumin. In a first series of experiments, intact cells were incubated for 30 min in the absence or presence of 0.1 nM mini-glucagon, in a medium containing 0.5 or 2 mM Ca2+ (A). In a second series of experiments, cells were preincubated for 10 min with or without (control) 0.1 nM mini-glucagon, then permeabilized with digitonin, and incubated for 30 min with 10 µM GTP and increasing [Ca2+]i, in the absence or in the presence of 0.1 nM mini-glucagon (B). [Ca2+]i was determined both by the EQUIV program and the Fura-2 procedure as described under "Experimental Procedures." The amount of [3H]AA released was expressed as percentage of control value (139 ± 17 disintegrations/min/µg of protein, A; 62 ± 11 disintegrations/min/µg of protein, B). Values are means ± S.E. of triplicate determinations from two different experiments (A) or means ± S.E. of quadruplicate determinations from three to six different experiments (B).
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The presence of a cPLA2 in embryonic chick ventricular cells was evidenced by immunoblotting analysis using a mouse monoclonal antibody directed against the amino-terminal domain of human cPLA2 (amino acids 1-216) (Fig. 4). The major immunoreactive protein in embryonic chick ventricular cells had an apparent molecular mass of 80 kDa. Under the same experimental conditions, two bands were detected in rat ventricular myocytes, a major band of 105 kDa and an additional band of 85 kDa (Fig. 4).


Fig. 4. Immunochemical identification of cPLA2 in chick embryonic ventricular cells and in rat ventricular myocytes. Lysates (25 µg) of chick embryonic or rat ventricular myocytes were analyzed by SDS gel (7.5%) and probed with mouse monoclonal antibody against the amino-terminal part of the human cPLA2 (Santa Cruz), as described under "Experimental Procedures."
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Mini-glucagon Causes [45Ca] Accumulation into SR Stores via AA

We have previously shown that mini-glucagon increases [45Ca] accumulation into caffeine-sensitive SR stores. Our data raised the question as to the role of AA in mediating mini-glucagon-induced increase in SR Ca2+ content. Quiescent embryonic chick heart cells were incubated for 5 min at 37 °C, in a medium containing 2 mM [45Ca], with or without 0.1 nM mini-glucagon, and the influence of inhibitors of either PLA2 activity or AA metabolism was examined. After incubation, the cells were washed and subjected to digitonin lysis. [45Ca] accumulation was then evaluated, both in the digitonin-sensitive structures, taken as cytosolic fractions, and in digitonin-resistant structures. The SR Ca2+ pool was identified among digitonin-resistant structures on the basis of its caffeine sensitivity. As shown in Fig. 5, mini-glucagon elicited a 350 ± 58% increase in [45Ca] accumulation into the digitonin-resistant structures which was abolished in the presence of caffeine. PLA2 inhibitors markedly decreased (5 µM quinacrine) or completely prevented (10 µM AACOCF3) the [45Ca] accumulation caused by mini-glucagon. It should be noted that quinacrine and AACOCF3 had no significant effect on [45Ca] accumulation, when added alone.


Fig. 5. Mini-glucagon-induced [45Ca2+] accumulation into the SR stores is mediated by AA. Embryonic chick heart cells were incubated for 5 min with 2 mM [45Ca] (5 µCi/ml), as described under "Experimental Procedures," in the presence or absence of 0.1 nM mini-glucagon or 0.3 µM AA and with or without inhibitors of either PLA2 activity (5 µM quinacrine or 10 µM AACOCF3), or AA metabolism (30 µM NDGA + 1 µM indomethacin + 100 µM SKF 250A). At the end of the incubation, cells were subjected to digitonin lysis. The amount of [45Ca] accumulated into the digitonin-resistant structures was expressed as percentage of control values (38 ± 5 cpm [45Ca]/µg of protein). Values are means ± S.E. of quadruplicate determinations from two to eight different experiments. It has to be noted that, in each assay condition, the [45Ca] content in the cytosolic fraction was determined in parallel with the SR [45Ca] content, and that no significant change was detected (not shown).
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Exogenous application of AA, at a concentration of 0.3 µM, mimicked the effect of mini-glucagon, inducing a 270 ± 51% increase in [45Ca] accumulation into the caffeine-sensitive SR stores (Fig. 5). Taken together, these data strongly support the proposal that [45Ca] accumulation into SR stores induced by mini-glucagon relies on AA.

Next, we questioned whether AA action required its conversion into eicosanoids. Exposure of cells to a mixture including 30 µM NDGA, 1 µM indomethacin, and 100 µM SKF 250A (inhibitors of the lipoxygenase, cyclooxygenase, and epoxygenase pathways, respectively) potentiated [45Ca] accumulation into the SR stores in control conditions as well as in the presence of mini-glucagon (Fig. 5).

To check the efficiency of the mixture of AA metabolism inhibitors, the supernatants of [3H]AA-labeled cells were analyzed by TLC. As shown in Table I, in the absence or presence of mini-glucagon, the radioactivity of the supernatants recovered was 84-81% AA, 4-5% lipoxygenase products, and 2-1% cyclooxygenase products. This pattern was similar to that of standard [3H]AA (Table I), proving an efficient blockade by the inhibitors of the enzymatic degradation of AA. It has to be noted that exposure of the cells to inhibitors of AA metabolism induced a marked increase of [3H]AA release, both in control and mini-glucagon-treated cells. This denoted an active metabolism of AA into eicosanoids in embryonic chick heart cells. Taken together, these results suggest that AA, and not a product of its metabolism, triggers [45Ca] accumulation into the SR stores.

Arachidonic Acid Potentiates Caffeine-induced Ca2+ Mobilization

Previous imaging studies (6) have shown that mini-glucagon potentiates caffeine-induced Ca2+ mobilization from the SR. Thus, we next examined the effect of AA on Ca2+ transients triggered by caffeine in Fura-2 loaded cells. Fig. 6A shows Ca2+ transients during electrical stimulation, and after caffeine application under steady state conditions, i.e. within a few seconds after interruption of electrical stimulation, according to the protocol previously described (6). As previously reported (6), the application of 10 mM caffeine produced a unique [Ca2+]i transient, longer than those observed during electrical stimulation. AA added alone, at 0.3 µM, had no effect (Fig. 6B). In contrast, the application of 10 mM caffeine together with 0.3 µM AA resulted in a train of Ca2+ transients (Fig. 6C). These experiments confirmed that AA mimics mini-glucagon action and leads to the Ca2+ loading of caffeine-sensitive SR stores.


Fig. 6. AA potentiates Ca2+ release from the intracellular stores induced by caffeine. Embryonic chick ventricular cells were loaded with Fura-2 as described under "Experimental Procedures." Cells were electrically stimulated at 0.5 Hz and stimulation was discontinued a few seconds before the addition of 10 mM caffeine alone (A), 0.3 µM AA alone (B), or 10 mM caffeine plus 0.3 µM AA (C). These data were representative of at least 30 cells obtained from two different isolations.
[View Larger Version of this Image (18K GIF file)]

An additional series of experiments were performed to examine the dose dependence and the specificity of AA action on caffeine-induced Ca2+-mobilization (Fig. 7). The action of AA (C20:4) was dose-dependent, with a maximal (9.5 ± 0.1) number of Ca2+ spikes detected over a period of 40 s at 3 µM AA and a half-maximal response occurring at 100 nM AA (Fig. 7). ETYA, the C20 nonhydrolyzable analogue of AA, elicited a response identical to that of AA, a maximal effect (11.3 ± 0.1 spikes of Ca2+) being observed in the presence of 3 µM ETYA. In contrast, exposure of myocytes to arachidic acid (C20:0), linolenic acid (C18:3), cis-5,8,11,14,17-eicosapentaenoic acid (C20:5), cis-4,7,10,13,16,19-docosahexaenoic acid (C22:6), or arachidonyl-CoA (C20:4), in the concentration range of 0.3 to 3 µM, failed to mimic the action of AA on caffeine contractures (Fig. 7).


Fig. 7. Dose-dependent effects of AA and ETYA on caffeine-induced Ca2+ release from the intracellular stores. Embryonic chick ventricular cells were loaded with Fura-2 as described under "Experimental Procedures." Cells were electrically stimulated at 0.5 Hz and stimulation was discontinued a few seconds before the addition of 10 mM caffeine in the presence of increasing concentrations of either AA or ETYA. The number of Ca2+ spikes during the first 40 s following addition of caffeine and fatty acid is reported. Each point represents the mean ± S.E of at least 20 cells obtained from at least 2 different isolations. Arachidic acid (C20:0), linolenic acid (C18:3), cis-5,8,11,14,17-eicosapentaenoic acid (C20:5), cis-4,7,10,13,16,19-docosahexaenoic acid (C22:6), or arachidonyl-CoA (C20:4) were without effect.
[View Larger Version of this Image (27K GIF file)]

AA Mimics the Mini-glucagon Action on [Ca2+]i Cycling in Electrically Stimulated Cells

As previously reported (6) and as shown in Fig. 8A, perfusion of myocytes with 30 nM glucagon alone evoked a small increase in the amplitude of electrically stimulated Ca2+ transients (119 ± 2% of control amplitude, n = 58). Mini-glucagon added alone at 0.1 nM had no effect (n = 82, Fig. 8B). In contrast, when cells were perifused with 30 nM glucagon plus 0.1 nM mini-glucagon, a marked rise in the amplitude of Ca2+ transients was observed (200 ± 6% of control amplitude, n = 75) together with a 2-3-fold increase in diastolic [Ca2+]i, estimated as described previously in Refs. 6 and 21 (Fig. 8C).


Fig. 8. AA mimics mini-glucagon action on [Ca2+]i cycling in electrically stimulated cells. Embryonic chick ventricular cells were loaded with Fura-2 as described under "Experimental Procedures." Cells were electrically stimulated at 0.5 Hz and perfused in the presence of 30 nM glucagon alone (A), 0.1 nM mini-glucagon alone (B), 30 nM glucagon plus 0.1 nM mini-glucagon (C), 0.3 µM AA alone (D), 0.3 µM AA plus 30 nM glucagon (E), or 0.3 µM AA plus 75 µM 8-Br-cAMP (F). These data are typical tracings representative of at least 40 cells obtained from at least three different isolations.
[View Larger Version of this Image (66K GIF file)]

AA added alone had no effect on Ca2+ cycling in electrically stimulated cells (n = 75, Fig. 8D). However, when 0.3 µM AA was added in combination with 30 nM glucagon, marked increases in both the amplitude of Ca2+ transients (183 ± 7% of control amplitude, n = 69) and diastolic [Ca2+]i were observed (Fig. 8E). Similar results were obtained with 0.3 µM AA plus 75 µM 8 Br-cAMP (Fig. 8F). Under the same experimental conditions, 100 nM isoproterenol produced a 220 ± 9% increase over control in the amplitude of the Ca2+ transients (n = 40) along with a 3-fold increase in diastolic [Ca2+]i (not shown). These data clearly demonstrate the ability of AA to mimic the action of mini-glucagon.

The Synergistic Action of AA and 8-Br-cAMP on Cell [Ca2+]i Cycling Produces a Positive Inotropic Effect

Fig. 9A shows parallel measurements of Ca2+ and contraction in isolated myocytes stimulated at 0.5 Hz. Exposure of myocytes to 30 nM glucagon plus 0.1 nM mini-glucagon resulted in both an increase in the amplitude of Ca2+ transients (113% over control) and an increase in the amplitude of contraction (70% over control) (Fig. 9B). The addition of AA (0.3 µM) plus 8-Br-cAMP (75 µM) evoked 61 and 77% increases over control amplitude of both the contraction and Ca2+ transients, respectively (Fig. 9C), reproducing the effect of glucagon plus mini-glucagon. Perfusion of the cells with 0.3 µM AA was without effect on either [Ca2+]i transient or cell contraction (not shown) while 75 µM 8-Br-cAMP added alone evoked a 10% increase in the amplitude of the Ca2+ transients, and a small (4% over control) increase in contraction (not shown).


Fig. 9. Effects of AA added with 8-Br-cAMP, and mini-glucagon added with glucagon on Ca2+ transients and contraction. Embryonic chick ventricular cells were loaded with Fura-2 as described under "Experimental Procedures." Cells were electrically stimulated at 0.5 Hz. Each trace is an average of five steady-state beats in a single cell. Data are representative of at least 5 cells obtained from two different isolations. Control trace (A); 0.1 nM mini-glucagon plus 30 nM glucagon (B); 0.3 µM AA plus 75 µM 8-Br-cAMP (C).
[View Larger Version of this Image (22K GIF file)]


DISCUSSION

The goal of the present study was to examine the role of AA in mediating the actions of mini-glucagon in ventricular myocytes. Several lines of evidence support the conclusion that AA is indeed the second messenger of mini-glucagon: 1) mini-glucagon increases AA release from [3H]AA prelabeled myocytes, in a dose-dependent manner (0.001-0.1 nM), most likely via the activation of a Ca2+-dependent PLA2; 2) [Ca2+] accumulation into the SR stores of intact cells following exposure of myocytes to mini-glucagon (6) is prevented by PLA2 inhibitors, quinacrine, and AACOCF3; 3) the [Ca2+] accumulation into the SR stores is mimicked by AA as well as by its nonhydrolyzable analogue, ETYA; 4) AA acts synergistically with either glucagon or 8-Br-cAMP in increasing [Ca2+]i cycling in electrically stimulated myocytes; 5) finally, perfusion of ventricular myocytes with both AA and 8-Br-cAMP leads to a positive inotropic contractile response identical to that produced by the mixture, glucagon plus mini-glucagon.

It is known that the formation of AA may occur independently of PLA2 activity. For example, diacylglycerol can serve as a precursor for AA via activation of diacylglycerol lipase. Diacylglycerol can be produced upon activation of PLC or from phosphatidic acid, following PLD activation. However, in the present study, the cell permeant analogue of diacylglycerol, OAG, failed to reproduce mini-glucagon actions (not shown), making it unlikely that mini-glucagon stimulated AA release through a diacylglycerol pathway. In addition, mini-glucagon, in contrast with other hormones inducing AA release (i.e. endothelin, angiotensin or bradykinin (10-13, 19)), did not activate PLC (6). Taken together, these data support the proposal that mini-glucagon causes AA release via the activation of a PLA2.

Activation of PLA2 may lead to a PKC-dependent phosphorylation event (9, 20). However, such a mechanism is not likely here since ETYA, which is structurally similar to AA, but not a known activator of PKC, mimicked mini-glucagon action.

AA release evoked by mini-glucagon requires submicromolar concentrations of Ca2+. Angiotensin has been reported to stimulate the release of AA in heart cells through activation of a Ca2+-dependent PLA2 (11). However, the precise range of sensitivity of the enzyme toward Ca2+ has not been established. Three types of PLA2 have been identified in the cardiac tissue (20). A large part of the PLA2 activity in myocardium relies on a Ca2+-independent, 40-kDa PLA2 (27) responsible for the accelerated phospholipid catabolism during myocardial ischemia (27). Recently, a low molecular mass PLA2 (14 kDa), which is almost exclusively expressed in heart and placenta, has been cloned (28). This enzyme contains 16 cysteines, is sensitive to Ca2+, but only in the millimolar range and it remains to establish whether it is secreted or is located intracellularly. Finally, a PLA2 of high molecular mass, referred to as "cytosolic PLA2" (cPLA2), which has been purified (29) and cloned (30), is expressed in various tissues, including the heart (31, 32). The amino acid sequence inferred from the human cDNA encodes a 85.2 kDa protein, which, however, migrates as a 100-110 kDa protein on SDS-polyacrylamide gel electrophoresis (31). This reduced rate of migration has been suggested to be inherent to the sequence and not to glycosylation (31). Amino acid sequence of the chicken cPLA2 differs from the sequences of human and mouse enzymes by 20-30% (33). Therefore, the difference in the rates of migration (80 versus 105 kDa) that we observe between the chicken and the rat cPLA2 may be due to species variations in the amino acid sequence of the enzyme. It may be noted that a cPLA2 with an apparent molecular mass of ~80 kDa has recently been detected in rabbit vascular smooth muscle cells (34). Thus, our results indicate that cPLA2 is present in chick embryonic ventricular cells. As reviewed by Van Bilsen and Van der Vusse (20), the biochemical properties of this enzyme, including translocation to the membrane in a Ca2+-dependent manner and requisite for submicromolar concentrations of Ca2+ for optimal activity, make it an ideal candidate as a target for mini-glucagon action. Although the role of G proteins in the modulation of the cardiac PLA2 activity has not been subject of investigation, G proteins have been implicated in the regulation of the cytosolic PLA2 activity in other tissues (9, 35). We observed that the release of AA stimulated by mini-glucagon required GTP. Although further characterization is needed, these results suggest that mini-glucagon, through a G protein, activates a Ca2+-dependent PLA2.

The major role of AA as the precursor of the extended family of bioactive metabolites, the eicosanoids, is undisputed (15, 36). Our results suggest that the action of mini-glucagon is not dependent upon the production of eicosanoids from AA (Fig. 4 and Table I). More debated is the role of AA as a direct second messenger. Our data support the view that AA itself is an important signaling molecule in heart since, physiologically relevant (15), submicromolar concentrations of AA mimic mini-glucagon action. The most thoroughly characterized target of AA is protein kinase C (15). In heart cells, it has been shown that activation of protein kinase C by AA leads to the phosphorylation of troponin I and myosin light chain 2 (37). However, in our study, ETYA, which does not stimulate protein kinase C activity (19), mimics AA action and the target of AA remains to be identified.

In conclusion, we demonstrate that mini-glucagon action is mediated via AA and that AA together with 8-Br-cAMP triggers a positive inotropic response. Thus, the action of glucagon in heart does not only rely on a cAMP pathway but also requires the synergistic support of AA pathway. It is interesting to speculate whether such a synergism could also be involved in other positive inotropic responses to hormonal stimuli.


FOOTNOTES

*   This work was supported by the Institut National de la Santé et de la Recherche Médicale, the French Ministère de la Recherche et de la Technologie, and the Unité de Formation et de Recherche de Médecine, Créteil, Paris-Val de Marne.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.
1   The abbreviations used are: SR, sarcoplasmic reticulum; AA, arachidonic acid (cis-5,8,11,14-eicosatetraenoic acid); [Ca2+]i, intracellular free Ca2+ concentration; [Ca2+]e, extracellular free Ca2+ concentration; PLA2, phospholipase A2; ETYA, 5,8,11,14-eicosatetraynoic acid; 8-Br-cAMP, 8-bromo-cAMP; PLD, phospholipase D; PLC, phospholipase C; NDGA, nordihydroguaiaretic acid; OAG, 1-oleolyl-2acetyl-sn-glycerol.

ACKNOWLEDGEMENTS

We thank S. Lotersztajn, J. Masliah, and M. Berguerand for helpful discussion and J. Hanoune for his permanent support.


REFERENCES

  1. Pecker, F., and Pavoine, C. (1996) in Glucagon III: Handbook of Experimental Pharmacology (Lefèbvre, P. J., ed), pp. 75-104, Springer Verlag, Berlin
  2. Mallat, A., Pavoine, C., Dufour, M., Lotersztajn, S., Bataille, D., and Pecker, F. (1987) Nature 325, 620-622 [CrossRef][Medline] [Order article via Infotrieve]
  3. Blache, P., Kervran, A., Dufour, M., Martinez, J., Le-Nguyen, D., Lotersztajn, S., Pavoine, C., Pecker, F., and Bataille, D. (1990) J. Biol. Chem. 265, 21514-21519 [Abstract/Free Full Text]
  4. Blache, P., Kervran, A., Le-Nguyen, D., Dufour, M., Cohen-Solal, A., Duckworth, W., and Bataille, D. (1993) J. Biol. Chem. 268, 21748-21753 [Abstract/Free Full Text]
  5. Pavoine, C., Brechler, V., Kervran, A., Blache, P., Le-Nguyen, D., Laurent, S., Bataille, D., and Pecker, F. (1991) Am. J. Physiol. 260, C993-C999 [Abstract/Free Full Text]
  6. Sauvadet, A., Rohn, T., Pecker, F., and Pavoine, C. (1996) Circ. Res. 78, 102-109 [Abstract/Free Full Text]
  7. Méry, P. F., Brechler, V., Pavoine, C., Pecker, F., and Fischmeister, R. (1990) Nature 345, 158-161 [CrossRef][Medline] [Order article via Infotrieve]
  8. Brechler, V., Pavoine, C., Hanf, R., Garbarz, E., Fischmeister, R., and Pecker, F. (1992) J. Biol. Chem. 267, 15496-15501 [Abstract/Free Full Text]
  9. Mukherjee, A. B., Miele, L., and Pattabiraman, N. (1994) Biochem. Pharmacol. 48, 1-10 [CrossRef][Medline] [Order article via Infotrieve]
  10. Damron, D. S., Van Wagoner, D. R., Moravec, C. S., and Bond, M. (1993) J. Biol. Chem. 268, 27335-27344 [Abstract/Free Full Text]
  11. Lokuta, A. J., Cooper, C., Gaa, S. T., Wang, H. E., and Rogers, T. B. (1994) J. Biol. Chem. 269, 4832-4838 [Abstract/Free Full Text]
  12. Hsueh, W., Isakson, P., and Needleman, P. (1977) Prostaglandins 13, 1073-1091 [Medline] [Order article via Infotrieve]
  13. Prasad, M. (1991) Biochem. Biophys. Res. Commun. 174, 952-957 [Medline] [Order article via Infotrieve]
  14. Piomelli, D. (1995) in Psychopharmacology: the Fourth Generation of Progress (Bloom, F. E., and Kupfer, D. J., eds), pp. 595-607, Raven Press, New York
  15. Khan, W. A., Blobe, G. C., and Hannun, Y. A. (1995) Cell Signalling 7, 171-184 [CrossRef][Medline] [Order article via Infotrieve]
  16. Kim, D., and Clapham, D. E. (1989) Science 244, 1174-1176 [Medline] [Order article via Infotrieve]
  17. Honoré, H., Barhanin, J., Attali, B., Lesage, F., and Ladzunski, M. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 1937-1944 [Abstract]
  18. Huang, J. M. C., Xian, H., and Bacaner, M. (1992) Proc. Natl. Acad. Sci. U. S. A. 89, 6452-6456 [Abstract]
  19. Damron, D. S., and Bond, M. (1993) Circ. Res. 72, 376-386 [Abstract]
  20. Van Bilsen, M., and Van der Vusse, G. J. (1995) Cardiovasc. Res. 30, 518-529 [CrossRef][Medline] [Order article via Infotrieve]
  21. Sauvadet, A., Pecker, F., and Pavoine, C. (1995) Cell Calcium 18, 76-85 [Medline] [Order article via Infotrieve]
  22. Altschuld, R. A., Wenger, W. C., Lamka, K. G., Kindig, O. R., Capen, C. C., Mizuhira, V., Vander Heide, R. S., and Brierley, G. P. (1985) J. Biol. Chem. 260, 14325-14334 [Abstract/Free Full Text]
  23. Donnadieu, E., Bismuth, G., and Trautmann, A. (1992) J. Biol. Chem. 267, 25864-25872 [Abstract/Free Full Text]
  24. Bassani, R. A., Bassani, J. W. M., and Bers, D. M. (1992) J. Physiol. 453, 591-608 [Abstract]
  25. Matzuoka, I., and Nakanishi, H. (1985) Thromb. Res. 37, 185-193 [Medline] [Order article via Infotrieve]
  26. Galron, R. Y., Kloogy, A., Bdolah, A., and Sokolovsky, M. (1985) Biochem. Biophys. Res. Commun. 163, 936-943
  27. Hazen, S. L., Ford, D. A., and Gross, R. W. (1991) J. Biol. Chem. 266, 5629-5633 [Abstract/Free Full Text]
  28. Chen, J., Engle, S. J., Seilhamer, J. J., and Tischfield, J. A. (1994) J. Biol. Chem. 269, 23018-23024 [Abstract/Free Full Text]
  29. Clark, J. D., Milona, N., and Knopf, J. L. (1990) Proc. Natl. Acad. Sci. U. S. A. 87, 7708-7712 [Abstract]
  30. Clark, J. D., Lin, L. L., Kriz, R. W., Ramesha, C. S., Sultzman, L. A., Lin, A. Y., Milona, N., and Knopf, J. L. (1991) Cell 65, 1043-1051 [Medline] [Order article via Infotrieve]
  31. Clark, J. D., Schievella, A. R., Nalefski, E. A., and Lin, L. L. (1995) J. Lipid Mediators Cell Signalling 12, 83-117 [CrossRef][Medline] [Order article via Infotrieve]
  32. Sharp, J. D., and White, D. L. (1993) J. Lipid Mediators 8, 183-189 [Medline] [Order article via Infotrieve]
  33. Nalefski, E. A., Sultzman, L. A., Martin, D. M., Kriz, R. W., Towler, P. S., Knopf, J. L., and Clark, J. D. (1994) J. Biol. Chem. 269, 18239-18249 [Abstract/Free Full Text]
  34. Muthalif, M. M., Benter, I. F., Uddin, M. R., and Malik, K. U. (1996) J. Biol. Chem. 271, 30149-30157 [Abstract/Free Full Text]
  35. Axelrod, J. (1995) Trends Neurosci 18, 64-65 [CrossRef][Medline] [Order article via Infotrieve]
  36. Van der Vusse, G. J., Glatz, J. F. C., Stam, H. C. G., and Reneman, R. S. (1992) Physiol. Rev. 72, 881-940 [Free Full Text]
  37. Damron, D. S., Van Wagoner, D. R., Sweet, W., Moravec, C. S., and Bond, M. (1995) Circ. Res. 76, 1011-1019 [Abstract/Free Full Text]

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