Endothelin-Mediated Positive Inotropic Effect Induced by Reactive Oxygen Species in Isolated Cardiac Muscle
Abstract Cardiac endothelium, both coronary and endocardial, produces a number of inotropic molecules. Changes in cardiac endothelial function by substances in the superfusing blood may thus participate in the control of muscle-pump performance of the heart. Reactive oxygen species (ROS) have been implicated in normal and pathological vascular physiology by influencing vascular endothelial function. Therefore, we examined the influence of ROS on endocardial endothelial modulation of myocardial performance. Right ventricular cat papillary muscles were briefly (15 s) exposed to electrolysis-generated ROS. Peak total isometric twitch tension and peak rate of tension development increased by 7.8±0.7% (P<.05) and 9.7±1.5% (P<.05), respectively (n=12). Isometric twitch duration was slightly increased (time from stimulus to half isometric relaxation, +2.7±0.6%; P<.05). ROS scavengers such as ascorbic acid (n=6), superoxide dismutase and catalase (n=8), or catalase alone (n=6), but not superoxide dismutase alone (n=6), blocked the inotropic effect. Interestingly, the positive inotropic effect was completely blocked by selectively damaging endocardial endothelium (Triton X-100, 0.5%, 1-s immersion, n=7) before ROS generation and by preincubating the muscles with the endothelin-A receptor antagonist BQ 123 (n=11). Preincubation with NG-nitro-l-arginine methyl ester and indomethacin (n=5) or with atenolol (n=6) did not influence the inotropic effect. Confocal scanning laser microscopic observations of muscles stained with viability tracers (n=9) revealed that significantly more but not all endocardial endothelial cells were damaged in electrolysis-treated muscles than in control muscles (42±5% versus 14±4%, P<.05). Accordingly, brief exposure of isolated cardiac muscle to electrolysis-generated ROS damaged the endocardial surface in part and increased contractile performance by stimulating endothelin release from endocardial endothelium. Hence, ROS-induced endothelin release from endocardial endothelium may be involved in normal and/or disturbed regulation of cardiac function.
Cardiac endothelial cells, both coronary (micro)vascular and endocardial, produce a number of molecules that may influence the contractile function of the underlying myocardium, such as endothelin-1,1 nitric oxide (NO),2 and prostaglandins.3 Therefore, changes in cardiac endothelial function induced by physical or humoral stimuli may constitute an important control of cardiac performance.4 5 The net effect of the release of cardiac endothelium-derived cardioactive substances in basal in vitro and in vivo conditions seems to be a tonic positive inotropic and contraction-prolonging effect. This has been concluded from experiments with isolated cardiac muscle,6 7 isolated intact hearts,8 and in situ hearts from open-chest dogs,9 in which a selective destruction of endocardial or coronary (micro)vascular endothelial cells resulted in an immediate and irreversible negative inotropic effect and an abbreviation of the contraction-relaxation cycle. Whether cardiac endothelium-derived NO, prostaglandins, and endothelin-1 cooperate or whether another, yet-unidentified, factor(s) or mechanism(s) participates in this positive inotropic and contraction-prolonging effect is still under debate.4 10
Endothelial dysfunction has been defined as an imbalance between endothelium-derived vascular relaxing and contracting factors, between anticoagulant and procoagulant mediators, and between growth-inhibiting and growth-promoting factors and is thought to participate in the initiation and/or progression of numerous cardiovascular diseases, including hypertension, hyperlipidemia, atherosclerosis, microvascular angina, diabetic angiopathies, reperfusion injury, and heart failure (for review see Reference 1111 ). To what extent (endo)cardial endothelial dysfunction results in a disturbed control of cardiac performance, cell growth, or other cardiac functions by an imbalanced release of endothelin-1, NO, prostaglandins, or other factors has not yet been established.
Reactive oxygen species (ROS) are continuously formed in vivo. ROS participate in the pathogenesis of various diseases, such as hypertension,12 atherosclerosis,13 diabetes mellitus,14 ischemia/reperfusion injury,15 and heart failure,16 probably by modulating or impairing endothelial cell function. In addition, ROS have also been ascribed a role in physiological endothelium-mediated regulation of vascular tone.17 Since endothelial cells can be both a target and a source for ROS,18 19 interaction between ROS and the endothelium thus appears to play an important role in normal and abnormal cardiovascular homeostasis.
In the present study, we investigated the influence of ROS on the action of endocardial endothelium (EE) on myocardial performance. Therefore, the contractile response of isolated papillary muscles to electrolysis-generated ROS was examined in the presence or absence of EE and/or inhibitors of endothelium-derived cardioactive substances. We preferred electrolysis above pharmacological or enzymatic ROS-producing procedures mainly because it offered the possibility of an on/off oxidative stress, short enough to avoid myocardial damage. Possible endothelial and myocardial cell damage was assessed biochemically by measurement of lipid peroxidation products and morphologically by confocal scanning laser microscopic observations of muscles stained with viability tracers.
Materials and Methods
Cardiac Muscle Preparation
Papillary muscles (n=76) were isolated from the right ventricles of cats, which had been anesthetized with pentobarbitone sodium, and mounted vertically in an organ bath filled with Krebs-Ringer solution containing (mmol/L) NaCl 118, KCl 4.7, MgSO4 · 7H2O 1.2, NaHCO3 20, KH2PO4 1.1, glucose 4.5, and CaCl2 · 2H2O 1.25 (pH 7.4 and temperature of 35°C) and bubbled with a gas mixture of 95% O2/5% CO2. The lower end of the muscle was held by a phosphor-bronze clip, and the upper tendinous end was attached to an electromagnetic force-length transducer6 with a Tevdek 7-0 braided thread. Muscles were stimulated electrically at 0.2 Hz and at a voltage ≈10% above threshold by rectangular pulses of 5-ms duration through two longitudinally arranged platinum electrodes.
Electrolysis-Induced Oxidative Stress
ROS were produced by electrolyzing the bathing solution through two additional platinum electrodes (1×2 cm), flanking the contracting muscle perpendicularly to the two stimulation electrodes. A constant current (DC) generated by an RP 216 current unit (Eagle Products) was applied through these electrodes into the bathing solution. Generation of ROS by electrolyzing a physiological buffer solution has been described by Jackson et al20 and has been used frequently to investigate the influence of ROS in the cardiovascular system. In the present study, electrolysis was continued for only 15 s by use of a current intensity of 30 mA, which was far below the level (ie, 60 mA or more) at which fibrillatory movements of the muscle were evoked during electrolysis.
In a few experiments without isolated muscles in the bath, ROS production in the Krebs-Ringer solution was assayed by fluorescence measurements of dihydrorhodamine 123. During electrolysis, aliquots of 1 mL were taken from the Krebs-Ringer solution between the electrodes, immediately (sampling time, <1 s) placed in cuvettes containing 1 mL of dihydrorhodamine 123 (final concentration, 1 μmol/L), and shaken. Fluorescence was determined with a Shimadzu RF-500 spectrofluorometer at 530-nm emission wavelength, the intensity of the fluorescence being recorded as a millivolt response. Fluorescence was quantified by the xanthine oxidase/xanthine reaction, in which the generation of ROS was measured by the reduction of cytochrome C.
After a stabilization period of at least 2 hours at 29°C, the temperature was increased to 35°C, the temperature at which the actual experiments were performed. Further stabilization of the muscles was continued for ≈1 hour at the muscle length at which the maximal active tension was developed (lmax) until steady state was obtained. After electrolysis, contractile performance, derived from isotonic and isometric twitches, was followed for 1 hour. Contribution of ROS to the electrolysis-induced effects on myocardial performance was verified by performing experiments with muscles incubated (30 minutes before electrolysis) with ROS scavengers, including superoxide dismutase (SOD, 100 U/mL), catalase (CAT, 120 U/mL), and ascorbic acid (1 μmol/L), all obtained from Sigma Chemical Co. After 15 s of electrolysis (current intensity, 30 mA), contractile performance was followed for at least 1 hour.
The role of the EE and EE-derived factors in the ROS-induced response was studied by performing parallel experiments in four groups of muscles. In the first group, the control group, EE was undamaged, and no inhibitors or antagonists were added. In the second group, EE was selectively damaged 30 minutes before electrolysis by a 1-s immersion of muscles in 0.5% Triton X-100 (Sigma) while in their working position, immediately followed by an abundant wash (>250 mL) with control Krebs-Ringer solution at 35°C. As previously described, this procedure resulted in a characteristic alteration of the isometric twitch6 : early onset of relaxation with concomitant decrease in peak twitch but with no significant changes in the maximal calcium-activated force. Selective damage of the EE and functional as well as morphological integrity of the subjacent myocardium after this procedure have also been extensively discussed previously.6 In the third and fourth groups, muscles were incubated 30 minutes before electrolysis with NG-nitro-l-arginine methyl ester (L-NAME, 1 mmol/L, Sigma; an NO synthase blocker) plus indomethacin (1 μmol/L, Sigma; a cyclooxygenase inhibitor) (third group) or with BQ 123 (1 or 3 μmol/L, Sigma; a competitive endothelin-A receptor antagonist) (fourth group). Neither L-NAME+indomethacin nor BQ 123 consistently influenced basal contractile parameters. In addition, in intact muscles cumulative concentration responses were obtained for exogenous endothelin-1 (from 0.1 nmol/L to 0.1 μmol/L, Sigma) in the absence (n=3) and presence (n=3) of BQ 123 (1 μmol/L, added 30 minutes before endothelin-1 administration).
Measurements of Lipid Peroxidation
Lipid peroxidation was determined by using the thiobarbituric acid reaction. This reaction reveals several oxidized substances (thiobarbituric acid reaction substances [TBARS]).
At 5, 10, 15, and 30 minutes after electrolysis (30 mA, 15 s), 1 mL of a 30 mL Krebs-Ringer solution containing the muscle preparation was mixed with 2 mL of 0.375% thiobarbituric acid and 15% trichloroacetic acid in 0.25N HCl. The mixture was left at room temperature for 10 minutes, then heated to 80°C for 15 minutes, cooled to room temperature, and centrifuged at 1000g for 15 minutes. The supernatant was used to measure TBARS absorbance at 532 nm.
Confocal Scanning Laser Microscopy and Viability Tracers
Cellular viability of myocardial and endothelial cells was assayed by using ethidium homodimer, TOPRO, and Bodipy phallacidin (all from Molecular Probes). Both ethidium homodimer and TOPRO are, like propidium iodide and other viability tracers, impermeant to live cells but bind to DNA and RNA in cells with a damaged membrane.21 22 Ethidium homodimer (final concentration, 0.7 μmol/L) was added to the bathing solution after the experiment, before fixation, thus only staining DNA and RNA of cells damaged during the experiment. For quantification of cellular damage, TOPRO staining was performed after fixation, thus staining the nuclei of all cells. In addition, all papillary muscles were stained en block with Bodipy phallacidin (3.3 μmol/L). Bodipy phallacidin binds specifically to filamentous actin (F-actin).23 Strips of stained specimens were mounted, endothelial surface at the top, in a small chamber on a slide. For en face observation, a Polyvar 2 epifluorescence microscope (Reichert) or a model 600 confocal scanning laser microscope (Bio-Rad) was used.
Data are expressed as mean±SEM. In the different muscle groups, the average of the parameter values measured at 20, 30, 40, and 50 minutes after electrolysis was compared with control values (paired Student’s t test). To compare the effect of electrolysis between the different conditions, a nonparametric Kruskal-Wallis test was performed on the percent changes induced by electrolysis, followed by a Dunn-type multiple comparison.
Measurements of ROS
Production of ROS depended on the applied current intensity (Fig 1A⇓). At a constant current intensity (eg, 30 mA), production of ROS was constant (Fig 1B⇓). At this current intensity, electrolysis of the Krebs-Ringer solution produced a 231±4-mV fluorescence response, equivalent to 32 μmol/L ROS. Fig 1C⇓ shows the influence of different ROS scavengers on the fluorescence intensity at 30 mA. SOD (a superoxide anion radical scavenger) depressed fluorescence intensity by 88.8%, although dihydrorhodamine 123 has been shown to be most sensitive to hydrogen peroxide. CAT, a hydrogen peroxide scavenger, almost completely depressed fluorescence, whereas dimethyl sulfoxide (DMSO), a hydroxyl radical scavenger, only slightly reduced fluorescence.
Influence of Electrolysis-Generated ROS on Contractile Performance
Electrolyzing the Krebs-Ringer solution surrounding the muscle for 15 s at 30 mA significantly increased peak twitch total tension by 7.8±0.7% (from 49.5±7.7 to 53.2±8.2 mN/mm2, P<.05) and peak rate of tension development by 9.7±1.5% (from 270±40 to 288±38 mN/mm2 per second, P<.05), whereas peak rate of preloaded isotonic shortening increased by 5.3±1.2% (from 1.03±0.08 to 1.08±0.08 lmax/s, P<.05) (Fig 2⇓). Interestingly, isometric twitch duration (time from stimulus to half relaxation) increased by 2.7±0.6% (from 442±22 to 454±22 ms, P<.05), whereas time from stimulus to peak twitch total tension did not change. The onset of the positive inotropic response was slow (5 to 10 minutes after electrolysis) and remained present until ≥50 minutes thereafter (Fig 2⇓). Replacing the electrolyzed bathing solution by control Krebs-Ringer solution (same pH and temperature) within 30 s after interrupting the electrolysis or after the full appearance of the positive inotropic effect did not influence the above positive inotropic response. These observations almost certainly exclude the possibility that the inotropic response resulted from electrolysis-induced change in pH or other ionic concentrations in the bathing solution. Of note, during the 15-s electrolysis and thereafter, resting properties of the muscle were not affected. In a few muscles (n=4), the mechanical response became highly unstable after electrolysis; because no steady state was attained, these muscles were excluded. Electrolysis-induced mechanical instability was observed only in muscles with an undamaged EE (ie, not Triton treated). Changing the Krebs-Ringer solution did not interrupt this unstable response.
To verify whether ROS were responsible for the contractile changes induced by electrolysis, similar experiments were performed in the presence of ROS scavengers, added at least 15 minutes before electrolysis. A combination of SOD (100 U/mL) and CAT (120 U/mL) was added in eight muscle preparations with an undamaged EE. Ascorbic acid (1 μmol/L) was added in another six preparations with an undamaged EE. In the presence of these scavengers, no inotropic response could be observed after electrolysis (Fig 3⇓). Similarly, CAT alone (n=6, 120 U/mL) completely suppressed the inotropic response. By contrast, SOD alone (n=6, 100 U/mL) did not significantly influence the basic observations (Fig 3⇓).
Atenolol (1 μmol/L), a hydrophilic β-adrenergic–blocking agent, added to the bathing solution 15 minutes before electrolysis did not affect the inotropic response of muscles to electrolysis (data not shown, n=6). A comparable increase in isometric and isotonic inotropic parameters as well as in isometric twitch duration was observed in the presence or absence of atenolol. This observation excluded the possibility that the inotropic response resulted from the release of endogenous catecholamines during electrolysis.
Role of EE and Endothelium-Derived Substances
Electrolysis did not influence contractile performance of isolated cardiac muscle when the EE had been selectively damaged 30 minutes before electrolysis (Fig 4⇓), suggesting that inotropic agents released from the EE were responsible for the positive inotropic effect after electrolysis in muscles with an undamaged EE. Therefore, experiments were repeated with muscles incubated with synthesis inhibitors or antagonists of known EE-derived metabolic substances added to the bathing solution 30 minutes before electrolysis. Preincubation with L-NAME (1 mmol/L) and indomethacin (1 μmol/L) (n=5) did not alter the inotropic response induced by electrolysis (Fig 4⇓), suggesting that neither EE-derived NO nor EE-derived cyclooxygenase products were involved. Preincubation with BQ 123, a competitive endothelin-A receptor antagonist, reduced the positive inotropic response at 1 μmol/L (n=6, Fig 4⇓) and completely suppressed the response at 3 μmol/L (peak twitch total tension, 97.1±1.6% of baseline; maximal rate of tension development, 97.3±0.8% of baseline; n=5).
In intact cardiac muscles (n=6), concentration-response experiments with exogenous endothelin-1 in the absence and presence of BQ 123 (1 μmol/L) revealed that the pattern of endothelin-1–induced positive inotropic effect was similar to the ROS-induced effect. In particular, at a concentration of 0.1 nmol/L, peak twitch total tension increased by 10.0±1.5%, peak rate of tension development increased by 13.9±2.0%, and time from stimulus to half relaxation increased by 2.7±0.2%, whereas time from stimulus to peak isometric tension did not change, suggesting that a similarly effective concentration of endothelin-1 was released by the EE after ROS production. In addition, similar to the ROS-induced effect, the positive inotropic effect appeared only 5 to 7 minutes after endothelin-1 administration, was neither transient nor washable, and was blocked by preincubating the muscles with 1 μmol/L BQ 123 (peak twitch total tension, 101.9±1.1% of baseline; maximal rate of tension development, 103.0±1.8% of baseline).
Effect of Electrolysis-Generated ROS on Cellular Viability
Measurements of Lipid Peroxidation Products
Detection of lipid peroxidation by the thiobarbituric acid reaction at 5, 10, 15, and 30 minutes after electrolysis did not reveal any elevation in the amount of TBARS in the Krebs-Ringer solution (data not shown). This result suggested that the brief electrolysis-induced oxidative stress did not induce major membrane damage or cell death.
Functional Morphology of EE and Myocardium
En face optical sections through the EE of electrolysis-treated muscles and dual-channel observations with the confocal scanning laser microscope revealed two different states of EE cells.
EE cells with nuclei intensively labeled by ethidium homodimer. These cells (Fig 5C⇓) represented nonviable cells, as further suggested by the absence of actin staining and by the absence of cytoplasmic ethidium homodimer staining surrounding the nuclei, which indicated that F-actin was disassembled and that RNA had been washed out.
EE cells without labeling of the nuclei by ethidium homodimer and with a normal F-actin pattern. These cells (Fig 5A⇑ and 5C⇑) represented viable cells with an intact cell membrane. As in the EE of rat hearts,24 actin filaments usually outlined the periphery of cat EE cells and, less frequently, formed centrally located stress fibers. In some cells with a normal F-actin pattern, the nuclei were weakly stained with ethidium homodimer, indicating that some membrane damage had occurred (Fig 5B⇑). The cytoplasmic staining surrounding the nuclei showed, however, that RNA had not been washed out (Fig 5B⇑). These cells were probably still viable but might have been dysfunctional.
The proportion of viable to nonviable EE cells in five electrolysis-treated muscles was quantified and compared with four control muscles. Therefore, the number of nonviable EE cells (ie, the number of EE nuclei intensively stained with ethidium homodimer) was compared with the total number of EE cells (ie, the number of EE nuclei stained with TOPRO). In both muscle groups, some “naked” zones without EE cells were observed. The actual total number of EE cells per 104-μm2 muscle surface was not significantly different in control muscles (19.9±1.2) and in electrolysis-treated muscles (15.5±3.0) (Mann-Whitney, P>.05). Of these latter cells, 42±5% were intensively stained with ethidium homodimer and thus nonviable (versus 14±4% in control muscles; Mann-Whitney, P<.05). Optical sections and dual-channel observations of subendothelial myocardium demonstrated the presence of intact myocytes with a normal striation pattern of F-actin (Fig 5D⇑).
In conclusion, although electrolysis was probably too brief to completely damage the EE or to damage any of the underlying myocytes, significantly more EE cells were damaged in electrolysis-treated muscles than in control muscles.
Electrolysis of a physiological buffer solution has been demonstrated to be a reliable method to produce various ROS, such as superoxide anion radical, hydrogen peroxide, hydroxyl radical, and hypochlorite. Brief exposure of isolated papillary muscles to electrolysis-generated ROS induced a positive inotropic effect, consistently appearing 5 to 10 minutes after electrolysis (Fig 2⇑). Preincubating the muscles with ROS scavengers suppressed the electrolysis-induced inotropic response, confirming that ROS were involved (Fig 3⇑). Because CAT (a specific scavenger of hydrogen peroxide), but not SOD (a specific scavenger of superoxide anion), blocked the inotropic effect (Fig 3⇑), hydrogen peroxide seemed to be at least partly responsible for the inotropic response. Hydrogen peroxide is by far the most stable of the ROS and was shown to influence endothelial function, even at subtoxic concentrations25 (<10 μmol/L; ie, approximately the concentration produced during electrolysis in the present study). In addition, a recent study revealed that subtoxic concentrations of hydrogen peroxide enhanced synthesis of at least 25 proteins in cultured vascular endothelial cells.26 The positive inotropic response was blocked by either damaging the EE before electrolysis or by preincubating the muscle with endothelin-A receptor antagonist. The magnitude, profile, and time course of the electrolysis-induced effect could be mimicked by administration of 0.1 nmol/L exogenous endothelin-1. These observations strongly suggested that ROS released endothelin from EE cells and therefore increased myocardial contractility.
Cardiac endothelium (more specifically, EE) releases at least three chemical messengers that may influence myocardial performance. First, EE cells express NO synthase (constitutive2 and inducible27 form). NO release by EE cells has been shown to elevate myocardial cGMP.10 The influence of EE-derived NO and the NO-cGMP pathway in the modulation of myocardial performance has been the subject of recent experiments in our and other laboratories. Shah and colleagues28 29 have reported that NO and cGMP decreased myocardial contraction in isolated ferret papillary muscles28 and in isolated cardiomyocytes,29 whereas Mohan et al30 have recently shown that the inotropic effect of cGMP in isolated cat papillary muscles is concentration dependent and relies on the state of the EE. Second, Mebazaa and colleagues3 31 demonstrated that cultured EE from sheep in basal conditions released large amounts of prostaglandins, up to 20 times more than vascular endothelium,3 and that this release increased further during hypoxia.31 However, the role of prostaglandins in the control of myocardial performance is not well understood. Because the presently described EE-dependent positive inotropic response was not altered by a combination of an NO synthase inhibitor (L-NAME) and a cyclooxygenase inhibitor (indomethacin) (Fig 4⇑), the response could not be accounted for by EE-derived NO and prostaglandins. Third, EE cells contain endothelin-1 mRNA.1 In the heart, endothelin-1 is highly inotropic1 and vasoconstrictive,32 even at low concentrations, and enhances cardiac cell growth.33 Recently, McClellan et al34 proposed endothelin storage and release from coronary endothelial cells as a cardioregulatory mechanism. Although the present data may add further evidence for endothelin release in vascular endothelium and EE as a cardioregulatory mechanism, a definitive conclusion about a role for endothelin in physiological regulation of cardiac function requires further investigation. The pathophysiological importance of endothelin in the cardiovascular system is suggested by the elevated plasma concentration of endothelin in various disorders, such as hypertension, atherosclerosis, myocardial infarction, and heart failure.35
The present study suggests that ROS may be a trigger for endothelin release from EE cells. To most investigators, ROS are believed to have biological importance only by their toxic effects, eg, by damaging or killing cells in pathophysiological conditions. However, growing experimental evidence suggests that nontoxic properties of ROS may be involved in normal aerobic biology. ROS have been involved in the physiological regulation of in vivo vascular tone by interaction with the endothelium.17 A chronic imbalance between ROS and oxidant defense, however, impairs endothelial function and may be involved in the initiation or progression of various cardiovascular diseases, including hypertension,12 atherosclerosis,13 diabetes,14 and myocardial stunning.15 In our experiments, confocal scanning laser microscopy of muscles only briefly exposed to electrolysis-generated ROS revealed that the myocytes were viable and had a normal actin pattern but that significantly more, but not all, EE cells were damaged when compared with control preparations. The functional state of partly damaged EE after electrolysis seems to resemble dysfunctional vascular endothelium, where an imbalanced release of endothelin compared with vasorelaxant substances (particularly NO) is characteristic.11 Hence, the present endothelin-mediated positive inotropic effect of electrolysis-generated ROS describes, for the first time, in vitro implications of EE dysfunction on myocardial contractility. ROS-induced enhanced endothelin release from dysfunctional EE cells in vivo may influence cardiac performance, coronary tone, and cardiac cell growth and thus be involved in the progression of cardiac diseases.
In conclusion, the present study demonstrates that a brief burst of electrolysis-generated ROS, which partly damaged the endocardial surface, increased contraction of isolated cat papillary muscle by stimulating endothelin release from EE cells. Hence, ROS-induced endothelin release from EE may participate in normal and/or pathological myocardial physiology.
Limitations of the Study
The present results describe in vitro observations of isolated papillary muscles in highly oxygenated aqueous buffer. The function of both EE and myocardium may differ significantly from in vivo conditions. In addition, the present results could be specific to electrolysis-induced oxidative stress. Therefore, extrapolating the present results to oxidative stress in general, and especially to oxidative stress in vivo, where many endogenous antioxidant mechanisms are operative, must be done cautiously. However, electrolysis of a physiological buffer solution has been demonstrated to be a reliable method to produce various ROS, such as superoxide anion radical, hydrogen peroxide, hydroxyl radical, and hypochlorite. We preferred electrolysis rather than pharmacological or enzymatic ROS-producing procedures mainly because it offered the possibility of an on/off oxidative stress that was short enough to avoid myocardial damage. Moreover, because of the gradual appearance of the inotropic response, which was not affected by replacing the bathing solution, it seemed unlikely that electrolysis-induced changes (other than ROS production), such as transient changes in pH or ionic concentrations, participated in the response.
This study was supported by the Belgian Programme on Interuniversity Poles of Attraction initiated by the Belgian State, Prime Minister’s Office, Science Policy Programming. The authors would like to thank C. Bridts and J. Meyers for assistance in free radical measurements.
- Received March 2, 1994.
- Accepted January 19, 1995.
- © 1995 American Heart Association, Inc.
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