Role of Nitric-Oxide Synthase, Free Radicals, and Protein Kinase C in Opioid-Induced Cardioprotection

Opioids generate free radicals that mediate protection in isolated cultured cardiomyocytes. We hypothesize that the nature of these radicals is nitric oxide, and that nitric oxide activates the protein kinase C (PKC) isoform. Through this signal transduction pathway, opiates protect cardiomyocytes during hypoxia and reoxygenation. Cell viability was quantified in chick embryonic ventricular myocytes with propidium iodide. Oxygen radicals were quantified using a molecular probe, 2 ,7 -dichlorofluorescin diacetate (DCFH-DA). After a 10-min infusion of the opioid receptor agonist BW373U86 (BW; 2 or 20 pM) and a 10-min drug-free period, cells were subjected to hypoxia for 1 h followed by reoxygenation for 3 h. BW produced a concentration-dependent reduction in cardiomyocyte death (2 pM, 35.3 3.9%, n 5; 20 pM, 21.5 4.0%, n 8, p 0.05 versus controls) and attenuated oxidant stress compared with controls (43.3 4.2%, n 8). The increase in DCFH-DA oxidation with BW before hypoxia was abolished by the specific nitric-oxide synthase inhibitors nitro-L-arginine methyl ester (LNAME) or N-monomethyl-L-arginine (L-NMMA) (100 M each). L-NAME or L-NMMA blocked the protective effects of BW. BW selectively increased the activity of PKC isoform in the particulate fraction, and its protection was abolished by the selective PKC inhibitor rottlerin (1 M). Similar to BW, infusion with 5 M of the nitric oxide donor S-nitroso-N-acetylpenicillamine (SNAP) reduced cardiomyocyte death (24.6 3.7, n 8), and this protection was blocked by chelerythrine or rottlerin. Chelerythrine and rottlerin had no effect on BW-generated oxygen radicals before hypoxia, but they abolished the protection of SNAP. The nature of DCFH oxidation produced by opioid receptor stimulation is nitric oxide. Nitric oxide mediates cardioprotection via activating PKC in isolated myocytes. Opioids protect against ischemia-reperfusion injury in vivo (Schultz et al., 1998a) and in vitro (Liang and Gross, 1999; Huh et al., 2001). Intravascular administration of opioids affects the coronary endothelium, circulating blood elements, and activates a signal transduction cascade in cardiomyocytes. Which effect (on endothelium, blood cells, or cardiomyocytes) accounts for cardioprotection remains unclear. Opioids increase nitric oxide synthesis from vascular endothelial cells and monocytes (Fimiani et al., 1999). In anesthetized rats, intravenous infusion of opioids reduces myocardial infarct size (Fryer et al., 2000). The role of nitric oxide in opioid-induced cardioprotection has not been defined. Several recent studies strongly suggest that nitric oxide from vascular endothelium mediates the cardioprotection of early and late preconditioning (Bolli et al., 1998, 2000). Since there are many confounding factors present in in vivo settings, we chose isolated cultured cardiomyocytes to determine whether nitric oxide, which originates from cardiomyocytes, mediates opioid-induced cardioprotection. Stimulation of opioid 1 receptors causes mitochondria to release oxygen radicals in cardiomyocytes, and this effect correlates with cardioprotection (McPherson and Yao, 2001a,b). These radicals are thought to activate protein kinase C (PKC) (Gopalakrishna and Anderson, 1989) and mediate cardioprotection (Simkhovich et al., 1998). The goal of this study is to determine the nature of these radicals produced by opioids (H2O2, nitric oxide, or both). Translocation of activated PKC , , and from cytosol to membranes has been detected in preconditioned hearts (Ping et al., 1997; Kawamura et al., 1998). Ping and colleagues (1999) have shown that nitric oxide induces translocation of the PKC isoform and mediates the late phase of preconditioning in a conscious rabbit model of cardiac ischemia-reperfusion (Bolli, 2000). To determine whether this signaling pathway mediates the cardioprotection of opioids, we studied the effects of the selective opioid receptor agonist BW373U86 (Chang et al., 1993) on the enzyme activity of Supported by National Heart, Lung, and Blood Institute U.S. Public Health Service Grants HL03881, HL70324, and HL70325. ABBREVIATIONS: PKC, protein kinase C; DCFH, 2 ,7 -dichlorofluorescin; DCFH-DA, DCFH diacetate; DCF, oxided DCFH; BW, BW373U86; SNAP, S-nitroso-N-acetylpenicillamine; BSS, balanced salt solution; PI, propidium iodide; PBS, phosphate-buffered saline; L-NMMA, Nmonomethyl-L-arginine; L-NAME, nitro-L-arginine methyl ester; BNTX, benzylidenenaltrexone; Chel, chelerythrine. 0022-3565/02/3013-1012–1019$7.00 THE JOURNAL OF PHARMACOLOGY AND EXPERIMENTAL THERAPEUTICS Vol. 301, No. 3 Copyright © 2002 by The American Society for Pharmacology and Experimental Therapeutics 4567/986097 JPET 301:1012–1019, 2002 Printed in U.S.A. 1012 at A PE T Jornals on A uust 0, 2017 jpet.asjournals.org D ow nladed from total PKC, and the and isoforms, in cytosol and particulate fractions. Materials and Methods Cardiomyocyte Isolation and Culture. Ventricular myocytes from 10-day-old chick embryos were prepared according to a method described previously (McPherson and Yao, 2001a,b). Briefly, hearts were harvested and placed in Hanks’ balanced salt solution lacking magnesium and calcium (Invitrogen, Carlsbad, CA). Ventricles were minced, and myocytes were dissociated by use of four to six repeated exposures to trypsin degradation (0.025%; Invitrogen) at 37°C with gentle agitation. Then, isolated cells were transferred to a solution with a trypsin inhibitor for 8 min, filtered through a 100m mesh filter, centrifuged for 5 min at 1200 rpm at 4°C, and finally resuspended in a nutritive medium described previously (McPherson and Yao, 2001b). Resuspended cells were placed in a Petri dish in a humidified incubator (5% CO2, 95% air at 37°C) for 45 min to promote early adherence of fibroblasts. Nonadherent cells were counted with a hemocytometer, and viability was measured with trypan blue (0.4%). Approximately 1 10 cells in nutritive medium were pipetted onto coverslips (25-mm) and incubated for 3 to 4 days, after which synchronous contractions of the monolayer were noted. Experiments were performed on spontaneously contracting cells at day 3 or 4 after isolation. The myocyte culture system was checked for nonmuscle cell contamination by staining with anti-myosin heavy chain monoclonal antibodies (CCM-52) labeled with horseradish peroxidase. More than 95% of plated cells stained for myosin. The remaining cells, less than 5% of total cells, consisted mainly of fibroblasts. This experiment was reported in 1996 (Vanden Hoek et al., 1996) and routinely performed to ensure purity of cardiomyocyte isolation and culture. Endothelial cell contamination was minimal. Hypoxia System. Glass coverslips containing spontaneously beating chick myocytes were placed in a stainless steel, 1-ml, flowthrough chamber (Penn Century Co., Philadelphia, PA). The chamber was sealed with Kynar film (McMaster-Carr, Elmhurst, IL) placed between the coverslip and the metal hypoxic chamber to minimize oxygen exchange between the chamber wall and the perfusate and then mounted on a temperature-controlled platform (37°C) on an inverted microscope. A water-jacketed glass equilibration column mounted above the microscope stage was used to equilibrate the perfusate to known oxygen tensions (PO2). The standard perfusion medium was equilibrated for 1 h before the experiment by bubbling with a gas mixture of 21% oxygen, 5% carbon dioxide, and 74% nitrogen. A hypoxic solution, composed of balanced salt solution (BSS) containing no glucose with 2-deoxyglucose (20 mM) added to inhibit glycolysis, was bubbled with a gas mixture of 20% carbon dioxide and 80% nitrogen for 1 h before the experiments. The pH of the perfusion solution was routinely verified (normoxic BSS, 7.4; hypoxic BSS, 6.8). Stainless steel or polymer tubing with low oxygen solubility connected the equilibration column to the flow-through chamber to minimize ambient oxygen transfer into the perfusate. PO2 in our hypoxic chamber was routinely monitored by Oxyspot (Medical Systems Inc., Greenvale, NY) under conditions identical to those of experiments using an optical phosphorescence quenching method (Wilson et al., 1988; Lo et al., 1996). PO2 in the chamber was 5.33 0.71 mm Hg (n 6) during hypoxia and 136 3.65 mm Hg (n 6) during normoxia perfusion. Necrosis Assay. Fluorescent cell images were obtained with an X10 objective lens (Nikon Fluor; Nikon, Tokyo, Japan). Data were acquired and analyzed with Metamorph software (Universal Imaging Corp., Downingtown, PA). There were approximately 600 cardiomyocytes under the selected field for each experiment. Multiple fields were examined and compared before each study; the field with normal synchronous contraction was chosen and monitored throughout experiments. Cell viability was quantified with the nuclear stain propidium iodide (PI; 5 M) (Molecular Probes, Eugene, OR), an exclusion fluorescent dye that binds to chromatin upon loss of membrane integrity (Altman et al., 1993). PI is not toxic to cells over a course of 8 h, permitting its addition to the perfusate throughout the experiments. At the completion of each experiment, digitonin (300 M) was added to the perfusate for 1 h. Digitonin disrupts the membrane integrity of all cells allowing PI to enter. Percent loss of viability (cell death) was expressed relative to the maximum value after 1 h of digitonin exposure (100%). Quantification of Oxygen Radicals. Oxygen radicals generated in cells were assessed with the probe 2 ,7 -dichlorofluorescin (DCFH). The membrane-permeable diacetate form of DCFH (DCFHDA) was added to the perfusate at a final concentration of 5 M. Within the cell, esterases cleave the acetate groups on DCFH-DA, thus trapping DCFH intracellularly (Sawada et al., 1996). Oxygen radicals in the cells lead to oxidation of DCFH, yielding the fluorescent product DCF (Vanden Hoek et al., 1996). DCFH in cardiomyocytes is readily oxidized by H2O2 or hydroxyl radical but is relatively insensitive to superoxide (Vanden Hoek et al., 1996). Fluorescence was measured with an excitation wavelength of 480 nm, dichroic 505-nm long pass and emitter bandpass of 535 nm (Chroma Technology, Brattleboro, VT) with neutral density filters to attenuate the excitation light intensity. Fluorescence intensity was assessed in clusters of several cells identified as regions of interest. The background was identified as an area without cells or with minimal cellular fluorescence. Intensity is reported as the percentage of initial value after subtraction of the background value. Permeablization of Cardiac Myocytes. We used a technique described by Gray et al. (1997) to permeabilize cardiomyocytes to allow a peptide ( V1–2 in this study) to enter cardiomyocytes before experiments. The temperature of isolated and cultured myocytes was slowly reduced by two sequential 2-min incubations, each with 2 ml (for 35-mm culture dishes) of fresh phosphate buffer solution. The first incubation with phosphate buffer solution was carried out at room temperature; the second, with chilled PBS in an ice bath. The PBS was discarded, and the cells were incubated with 1 ml of freshly prepared permeabilization buffer [20 mM HEPES, pH 7.4, 10 mM EGTA, 140 mM KCl, 50 g/ml saponin (Sigma-Aldrich, St. Louis, MO), 5 mM NaN3, and 5 mM oxalic acid dipotassium salt] containing the desired peptides for 10 min in an ice bath. ATP was added just before adding the permeabilization buffer to cells (i.e., 30 l of 200 mM ATP, pH 7.4, per milliliter of permeabilization buffer). The cells were then gently washed four times on ice with 2 ml of chilled PBS. Then, an additional 2 ml of chilled PBS was added to the cells for a 20-min recovery on ice. After the chilled PBS was removed, 2 ml of room temperature PBS was added, and the cells were placed at room temperature for 2 min. This step was repeated with PBS at 37°C, after which the original cell media were added back to the cells at 37°C. The cells were further incubated for 30 min at 37°C before interventions. PKC Enzyme Assay. Enzyme activity of total PKC and its and isoforms was measured by a method described previously (Ping et al., 1997, 1999). For each experiment, 5 million cells were collected in sample buffer (50 mM Tris-HCl, pH 7.5, 5 mM EDTA, 10 mM each EGTA and benzamidine, 50 g/ml phenylmethylsulfonyl fluoride, 10 g/ml each of aprotinin, leupeptin, and pepstatin A, and 0.3% -mercaptoethanol) (Sigma-Aldrich). The collection of cells was centrifuged at 45,000g for 30 min and separated into cytosol and particulate fractions. The particulate pellet was dissolved ultrasonically in sample buffer. Enzyme protein concentration was determined according to the Bradford method (Bradford, 1976). Each fraction, 50 to 100 g, was assayed for activity of total PKC and its isoforms (assay kit; Amersham Biosciences, Piscataway, NJ). The activity of total PKC in the pellet (particulate) and the supernatant (cytosolic) was assayed separately. For PKC and assay, proteins were immunoprecipitated overnight by PKC and monoclonal antibody (BD Biosciences PharMingen, San Diego, CA) in immunoprecipitation buffer (pH 7.4) (150 mM NaCl, 50 mM Tris, 1 mM EGTA, 1 mM EDTA, 1% Nonidet P-40, 1 mM sodium orthovanadate, 1 mM phenylmethylsulfonyl Nitric Oxide and PKC Signal Opioid Cardioprotection 1013 at A PE T Jornals on A uust 0, 2017 jpet.asjournals.org D ow nladed from fluoride, 16 g/ml benzamidine-HCl, and 10 g/ml each for phenanthroline, aprotinin, leupeptin, and pepstatin A) (Sigma-Aldrich) with protein A/G beads (Santa Cruz Biotechnology, Inc., Santa Cruz, CA). PKC -specific substrate (ERMRPRKRQGSVRRRV) (BIOMOL Research Laboratories, Plymouth Meeting, PA) was used for the phosphorylation reaction with [P]ATP (Amersham Biosciences). Since there is no specific substrate available for PKC , the same substrate for total PKC was used for the phosphorylation reaction with [P]ATP with the proteins that were immunoprecipitated overnight by PKC monoclonal antibody (BD Biosciences PharMingen). Additionally, we used rottlerin (1 M) to block PKC and V1–2 (100 M) to block PKC phosphorylation to ensure the specificity of the increased enzyme activity. Chemicals. BW373U86, S-nitroso-N-acetylpenicillamine (SNAP), N-monomethyl-L-arginine (L-NMMA), and nitro-L-arginine methyl ester (L-NAME) were purchased from Sigma-Aldrich. Chelerythrine was purchased from Calbiochem-Novabiochem Corp. (San Diego, CA). BW373U86, L-NMMA, L-NAME, or chelerythrine was dissolved in BSS buffer before administration. Rottlerin and BNTX were purchased from BIOMOL Research Laboratories and dissolved in a 1:5 cocktail of ethanol/saline. PI and DCFH-DA were purchased from Molecular Probes. Experimental Protocol. Figure 1 shows the experimental protocol. Fifteen groups of cardiomyocytes [control, BW (2 pM), BW (20 pM), BNTX (0.1 M), BNTX BW, L-NMMA (100 M), L-NMMA BW, L-NAME (100 M), L-NAME BW, rottlerin (1 M), rottlerin BW, SNAP (5 M), rottlerin SNAP, chelerythrine (4 M), and SNAP Chel] were studied. Cardiocytes were subjected to 1 h of hypoxia followed by 3 h of reoxygenation. Ethanol/saline (1:5) (control series) or BW (2 or 20 pM) was added to the perfusate for 10 min followed by 10 min of a drug-free period before the cells were subjected to hypoxia and reoxygenation. For the corresponding series, rottlerin (1 M), L-NMMA (100 M), or L-NAME (100 M) was added to the perfusate during baseline (1 h) before 60 min of hypoxia. Nine additional series of experiments were used to determine the effects of the above interventions on production of oxygen radicals before and during hypoxia and reoxygenation. An additional six groups of cardiomyocytes were used to examine whether the nitric oxide donor SNAP (5 M) activates PKC and mediates cardioprotection [control, SNAP, chelerythrine (4 M), SNAP Chel, rottlerin (1 M), and SNAP rottlerin]. For the PKC enzyme activity assay, BW (20 pM) was administered for 10 min followed by a 10-min drug-free period, then cardiocytes were collected for the assay. In the control group, vehicle (ethanol/ saline 1:5) was given for 10 min instead of BW administration. Statistical Analysis. Data are expressed as mean S.E.M. Differences between groups for cell death and oxygen radical production were compared by a two-factor analysis of variance with repeated measures and Fisher’s least significant difference test. Differences between groups were considered significant at values of P 0.05.