Free radicals are produced continuously by skeletal muscle fibers. Extracellular release of reactive oxygen species (ROS) and nitric oxide (NO) derivatives has been demonstrated, but little is known about intracellular oxidant regulation. We used a fluorescent oxidant probe, 2′,7′-dichlorofluorescin (DCFH), to assess net oxidant activity in passive muscle fiber bundles isolated from mouse diaphragm and studied in vitro. We tested the following three hypotheses. 1) Net oxidant activity is decreased by muscle cooling. 2) CO2 exposure depresses intracellular oxidant activity. 3) Muscle-derived ROS and NO both contribute to overall oxidant activity. Our results indicate that DCFH oxidation was diminished by cooling muscle fibers from 37°C to 23°C (P < 0.001). The rate of DCFH oxidation correlated positively with CO2 exposure (0–10%; P < 0.05) and negatively with concurrent changes in pH (7.0–8.5; P < 0.05). Separate exposures to anti-ROS enzymes (superoxide dismutase, 1 kU/ml; catalase, 1 kU/ml), a glutathione peroxidase mimetic (ebselen, 30 μM), NO synthase inhibitors (Nω-nitro-l-arginine methyl ester, 1 mM; Nω-monomethyl-l-arginine, 1 mM), or an NO scavenger (hemoglobin, 1 μM) each inhibited DCFH oxidation (P < 0.05). Oxidation was increased by hydrogen peroxide, 100 μM, an NO donor (NOC-22, 400 μM), or the substrate for NO synthase (l-arginine, 5 mM). We conclude that net oxidant activity in resting muscle fibers is 1) decreased at subphysiological temperatures, 2) increased by CO2 exposure, and 3) influenced by muscle-derived ROS and NO derivatives to similar degrees.
- oxidative stress
- reactive oxygen species
- respiratory muscles
skeletal muscle continually produces free radicals and free radical derivatives, most notably reactive oxygen species (ROS) and nitric oxide (NO) derivatives. Both ROS and NO influence cellular function by modulating excitation-contraction coupling (39, 42, 43, 47), glucose uptake (16), mitochondrial respiration (20), and gene expression (27). Accordingly, the regulation of intracellular oxidant cascades and the factors that influence oxidant activity are of considerable physiological importance.
Oxidant activity in muscle has primarily been studied using indirect indexes. These have included biochemical markers of ROS- or NO-mediated reactions, e.g., glutathione oxidation, malondialdehyde, carbonyl or nitrotyrosine adducts (3, 13, 14, 25, 48, 49, 54), and measurements of extracellular ROS and NO release (2, 19, 45). Only a few studies have measured intracellular oxidants in viable muscle fibers. Most of these have focused on the increases in oxidant production caused by biological stressors: fatiguing exercise (4, 21, 41, 50), inflammatory mediators (44), and heat stress (10, 55, 56). Little is known about oxidant regulation under less dramatic conditions, e.g., unfatiguing contractions and muscular inactivity. Such conditions are more common for muscles in vivo and provide a more physiological state for studying redox homeostasis. This manuscript describes experiments that evaluated metabolic factors that might influence cytosolic oxidant activity in resting muscle fibers. We tested the following three hypotheses.
Hypothesis 1: Oxidant Activity in Resting Muscle Fibers is Decreased by Cooling
The temperature of peripheral muscles may fall below core temperature in individuals subjected to extreme environments (30), and experimental muscle preparations are often studied at temperatures below 37°C to enhance stability (47). Cooling is expected to diminish production of ROS and NO derivatives since mitochondrial oxygen consumption is decreased (7) as are the enzymatic activities of oxidoreductases, e.g., NADPH oxidase (31) and NO synthase (51). The activities of antioxidant enzymes are also depressed by cooling, decreasing the capacity of muscle to buffer endogenous oxidants. We postulated that cooling would shift the net balance among these processes, depressing the overall activity of muscle-derived oxidants. This was tested by comparing cytoplasmic oxidant activities at 23°C vs. 37°C.
Hypothesis 2: CO2 Exposure Inhibits Oxidant Activity
Skeletal muscle is exposed to elevated CO2 levels under a variety of circumstances, including exercise (40), ischemia (22), and chronic obstructive pulmonary disease (1). Published reports suggest CO2 influences redox homeostasis. CO2 and bicarbonate react directly with NO derivatives (3, 52) and undergo electron exchange reactions to form carbonate radicals (28). Dissolved CO2 also promotes acidosis in biological systems, stimulating cellular production of ROS and NO (8). Studies in nonmuscle cell types are conflicted about the net effect of CO2-mediated reactions that appear to either promote (23, 37) or protect against (9, 12, 52) oxidative and nitrosative stress. In skeletal muscle, elevated CO2 levels inhibit ROS release by skeletal muscle and CO2 is proposed to limit the biological activity of NO (47). Based on these reports, we tested a range of CO2 levels (0–10%) for inhibitory effects on intracellular oxidant activity.
Hypothesis 3: Myogenic ROS and NO Both Contribute to Oxidant Activity in Resting Muscle Fibers
Prior experiments have shown that resting muscle fibers release either ROS (45) or NO (2) into the extracellular space. Interventions that interrupt either ROS or NO signaling have been shown to alter contractile function (32, 41). Surprisingly, however, detectable levels of NO derivatives have never been demonstrated within muscle fibers, nor have intracellular activities of the ROS and NO cascades been compared directly. We did both in this experiment.
2′,7′-Dichlorodihydrofluorescin diacetate (DCFH-DA; Molecular Probes, Eugene, OR) was dissolved in 100% ethyl alcohol, diluted in Krebs-Ringer solution, and stored at −80°C for later use. Reagents to interrupt free radical signaling included Nω-nitro-l-arginine methyl ester (l-NAME; Alexis, San Diego, CA), Nω-nitro-d-arginine methyl ester (d-NAME; Alexis), hemoglobin (Hgb; Oxis, Portland, OR), Cu/Zn superoxide dismutase (SOD; Oxis), catalase (Oxis), and N-(2-aminoethyl)-N-(2-hydroxy-2-nitrosohydrazino)-1,2-ethylenediamine (spermine NONOate or NOC-22; Calbiochem, San Diego, CA). All other chemicals were obtained from Sigma (St. Louis, MO). Reagents were dissolved in Krebs-Ringer solution before the experiment.
All procedures were conducted in accordance with the National Institutes of Health Guide for the Care and Use of Animals and were approved by the Institutional Review Board of Baylor College of Medicine. Adult male ICR mice [31 ± 0.37 (SE) g] were anesthetized and killed by rapid exsanguination. Muscle fiber bundles (133 ± 8 mg) were surgically isolated from the costal diaphragm and placed in Krebs-Ringer solution for subsequent study.
Measurement of Cytosolic Oxidant Activity
As described previously (41), oxidant activity was measured by use of a diffusible fluorochrome probe, DCFH-DA. The acetate group is cleaved by cytosolic esterases to yield DCFH, a polar nonfluorescent molecule retained by the cell. Cytosolic oxidants convert DCFH to its fluorescent derivative, 2′,7′-dichlorofluorescein (DCF; 480-nm excitation, 520-nm emissions). DCFH conversion to DCF is a nonspecific oxidation reaction that can be mediated by ROS (16), NO derivatives (15), oxidized thiols, metal centers, and other intracellular oxidants. DCFH competes for electrons with other redox-active molecules, notably intracellular antioxidants. The rate of DCFH conversion to DCF therefore reflects the dynamic balance between oxidant production and buffering, i.e., net oxidant activity, in the cytosolic compartment.
Fiber bundles were loaded with DCFH by in vitro incubation with DCFH-DA, 50 μM, for 45–60 min, time periods that enable probe equilibration (Fig. 1A). Accumulation of oxidized DCF was measured from representative areas of the fiber bundle surface (0.27 mm2) by use of an epifluorescence microscope (Labophot-2; Nikon Instruments, Melville, NY) with xenon lamp, 480-nm low-pass excitation filter, and 520-nm high-pass emissions filter. Emissions were recorded using a charge-coupled device camera (series 72; Dage-MTI, Michigan City, IN). A mechanical shutter in the excitation light pathway was controlled by computer using commercial data acquisition and analysis software (Optimas 4.02; Bioscan, Edmonds, WA) to standardize excitation time (33 ms). Images were acquired in real time and stored in a desktop computer for later analysis of mean emission intensity.
Photooxidation artifact was controlled by conducting experiments in a darkened laboratory and by standardizing excitation parameters to minimize the cumulative delivered energy (32). The protocol of Murrant et al. (32) was adapted for use in the current study based on observations that photooxidation is directly proportional to the number of excitation exposures at a given site (Fig. 1B, top). Photooxidation was quantified using excitation doublets delivered at 10-s intervals (Fig. 1B, bottom). A regression was fitted to the measured emission intensities; the y-intercept provided an estimate of DCF emissions 10 s before the initial excitation, in the absence of photooxidation artifact. Value of the y-intercept was calculated for each condition and used as a photooxidation-corrected signal in subsequent analyses of biological responses. Background subtraction is not required for this assay; in the absence of DCFH, neither muscle fibers nor Krebs-Ringer solution emits detectable fluorescence at 520 nm.
In paired comparisons, two muscle fiber bundles were isolated from homologous regions of the same muscle. Each was placed in a separate dish containing Krebs-Ringer solution aerated with 95% O2-5% CO2 (pH 7.3), loaded with DCFH, and incubated under passive conditions at either 23°C or 37°C. DCF emissions were measured using a standardized protocol. Autoxidation of DCFH-DA was measured under identical conditions but without fiber bundles.
For each CO2 condition tested, paired fiber bundles were isolated, placed in Krebs-Ringer solution aerated with 90% O2-5% CO2-5% N2, and loaded with DCFH. Basal emission intensity was measured from a selected site on each fiber bundle. Each fiber bundle then was incubated in buffer aerated with one of three gas mixtures: 90% O2-5% CO2-5% N2 (control), 90% O2-0% CO2-10% N2 (low CO2), or 90% O2-10% CO2-0% N2 (high CO2). After 45 min, a second measurement was made on each fiber bundle using a separate site. Changes in emission intensity during the second incubation period were expressed as a percentage of basal emissions and analyzed for CO2 effects. The effect of CO2 on pH of the Krebs-Ringer solution was measured using a Corning 240 pH meter.
ROS and NO effects.
Paired fiber bundles were mounted in separate chambers containing Krebs-Ringer solution aerated with 95% O2-5% CO2 at 37°C. Fiber bundles were pretreated for 20 min with probes to interrupt ROS or NO signaling and were loaded with DCFH. After 60 min passive incubation, fluorescence emissions were measured to assess oxidant activity. In separate experiments, we screened for chemical interactions between DCFH (generated in vitro by esterase conditioning of DCFH-DA; Ref. 32) and each ROS- or NO-selective probe in a muscle-free system. As expected (32), H2O2 and NOC-22 directly oxidized DCFH. Other probes did not.
Fluorescence images were analyzed post hoc for mean emission intensity using commercial software (Optimas) and were corrected for photooxidation (Fig. 1B). Separate software (Sigma Stat; SPSS, Chicago, IL) was used for statistical analyses. Data are reported as means ± SE. Student's paired t-test (53) was used to test the effect of individual interventions imposed for a single time period. Changes produced by a range of graded interventions were tested by either linear or second-order regression analyses. Statistical significance was accepted at P < 0.05.
Temperature Effects on Cytosolic Oxidants
Fluorescence emissions from fiber bundles studied at 23°C and 37°C are shown in Fig. 2. As in all current experiments, a paired design was used to compare DCFH oxidation rates in two fiber bundles from each animal. In six of six comparisons, DCF fluorescence emissions from the bundles exposed to room temperature (23°C) were less than emissions from bundles incubated at 37°C [mean 39.7 ± 7.8 (SE) vs. 87.3 ± 12.8 arbitrary units (AU); P < 0.001]. Autoxidation of DCFH over 60 min was greater at 37°C but remained negligible, only 5.7% of the signal detected from fiber bundles.
Data from studies performed using different CO2 concentrations are shown in Fig. 3. In paired comparisons (Fig. 3A), emission intensities measured in bundles exposed to 0% CO2 were less than intensities measured at 5% CO2 (26.7 ± 5.3 vs. 49.2 ± 5.0; P < 0.05). Intensities at 10% CO2 tended to be higher (31.5 ± 5.3 vs. 27.9 ± 4.5), but this difference was not significant. In a combined analysis across all CO2 levels (Fig. 3B), emission intensity was directly proportional to CO2 exposure (P < 0.05) with the greatest emission changes occurring at CO2 levels <5%. Figure 3B also depicts the inverse relationship between CO2 supply and pH of the bathing solution (P < 0.05). In a post hoc analysis (not shown), intracellular oxidant activity also was negatively correlated with pH [AU = 3.765 − (0.377 × pH); n = 20; P < 0.001]. Control studies in muscle-free systems showed that DCF fluorescence was diminished by CO2 exposure (P > 0.001) and by acidification using hydrochloric acid (P < 0.04). This represents a modest artifact that opposed the biological signal in our experiments but did not prevent resolution of CO2 effects.
Contributions of Myogenic ROS and NO
Data in Fig. 4 show that exposure to ROS-specific antioxidant enzymes depresses the rate of DCFH oxidation in muscle fibers. Incubation with either Cu,Zn SOD, 1 kU/ml, or catalase, 1 kU/ml, reduced DCF emissions significantly (P < 0.01). Treatment with a glutathione peroxidase mimic, ebselen, 30 μM, also decreased emissions (P < 0.05). In contrast, positive control studies using H2O2, 100 μM, caused a marked increase in DCF fluorescence (P < 0.05). These data confirm the expected contribution of muscle-derived ROS to this signal (41).
NO contributions to DCFH oxidation were assessed using a panel of complementary interventions. As shown in Fig. 5A, inhibition of NO synthase activity by use of l-NAME, 1 mM, decreased emissions by 38.5% (P < 0.05). d-NAME, the enzymatically inactive enantiomer, had no effect. In results not shown, a second NO synthase inhibitor, l-NMMA at 1 mM, had a similar effect, inhibiting DCFH oxidation by 30% (P < 0.05); d-NMMA had no effect (data not shown). The action of NO synthase inhibitors was mimicked by a nitric oxide scavenger, hemoglobin, 1 μM, which also depressed DCFH oxidation (P < 0.05). In positive control studies, exposure to the NO donor NOC-22, 400 μM, dramatically increased DCFH oxidation (P < 0.05). Figure 5B shows dose-dependant effects of l-arginine, the enzymatic substrate for NO synthesis. Exposure to either l-arginine or its inactive enantiomer d-arginine had no effect at low concentrations (1 mM). At a higher concentration (5 mM), l-arginine doubled the rate of DCFH oxidation (P < 0.05). d-Arginine exerted a nonspecific effect, increasing DCF emissions, but this was significantly less than the action of l-arginine (P < 0.05). Taken together, these results indicate that muscle-derived NO derivatives contribute substantially to the basal rate of DCFH oxidation in resting muscle fibers.
This study addressed factors thought to influence oxidant activity in the cytoplasm of intact muscle fibers. Experiments evaluated oxidant regulation under basal conditions, in the absence of muscle contraction or underlying pathology, using an intracellular fluorochrome probe to assess net oxidant activity. Our results established three points. 1) Oxidant activity is sensitive to temperature, diminishing significantly when muscle fibers are cooled to room temperature. 2) CO2 promotes oxidant activity under basal conditions. 3) Muscle-derived free radicals, both myogenic ROS and myogenic NO derivatives, are major determinants of overall oxidant activity.
DCFH Oxidation Assay
As described previously (41), we assessed cytosolic oxidant activity by use of DCFH, a redox-sensitive fluorochrome probe. DCFH oxidation to DCF is a nonspecific reaction mediated by biological oxidants that include ROS (16), NO derivatives (15), oxidized thiols, and metal centers. This reaction may involve multiple mediators; for example, DCFH oxidation by hydrogen peroxide requires peroxidase activity. Also, the transition from DCFH to DCF may also involve intermediate states, complicating the oxidation reaction mediated by NO derivatives. It is important to recognize that intracellular conversion of DCFH to DCF occurs in competition with other oxidation reactions. Among these, the traditional antioxidant mechanisms are included: glutathione oxidation, Cu,Zn- and Mn-SOD activities, catalase activity, and ascorbate oxidation. The overall rate of DCFH conversion to DCF therefore reflects a complex, highly dynamic balance between simultaneous rates of oxidant production and oxidant buffering by multiple pathways. We refer to this integrated signal as net oxidant activity under a given condition.
Factors that may distort this signal include DCFH availability and DCF retention. To avoid limitation by DCFH availability, we preload muscle fibers with excess DCFH as assessed by maximal fluorescence induced by photooxidation (32, 41). Experimental interventions did not alter DCFH loading in the current study. Cooling to 23°C did not inhibit DCFH loading relative to 37°C (data not shown); in other studies, muscle preparations were preloaded with DCFH under identical control conditions before interventions were introduced. DCF retention is a separate factor that may also affect assay performance. Transport of oxidized DCF out of muscle fibers is detectable at 37°C but not at 23°C (32). This difference tends to increase DCF fluorescence and apparent oxidant activity at cooler temperatures. This artifact may have caused us to underestimate the inhibitory effects of cooling in our current experiments. Fluorescein derivatives such as DCF also undergo protonation in acidic environments, which opposes transport across the cell membrane. We have no means of assessing DCF protonation in our current system. To the extent it occurred, protonation would favor DCF retention and enhance the apparent increase in oxidant activity observed under conditions of high CO2/low pH. Water loss from tissue has the opposite effect, concentrating DCF and increasing apparent oxidant activity. In the current experiments, SOD, catalase, and hemoglobin probably did not cross the sarcolemma and therefore tended to alter osmotic pressure, favoring water loss from the fibers. Any such changes would have been small, tending to lessen the observed decreases in DCF fluorescence.
Effects of Temperature
Temperature is expected to influence oxidant production by intracellular sources. Mitochondrial oxygen consumption (7) and the enzymatic activities of oxidoreductases, e.g., NADPH oxidase (31) and NO synthase (51), are strongly affected by changes in temperature. Antioxidant enzyme activities are also temperature sensitive, affecting the pathways that buffer muscle-derived oxidants. Increases or decreases in total oxidant activity reflect changes in net balance among these variables.
Only two other studies have directly addressed the thermal sensitivity of oxidant regulation in skeletal muscle fibers. Zuo and colleagues (55) focused on the cellular response to heat stress. Their work demonstrates that heating muscle from 37°C to 43°C causes intracellular superoxide anion levels to increase. This change was mirrored by increased release of superoxide anions into the extracellular space. A followup study by the same group (56) determined that this response does not depend directly on mitochondrial complex activity or sarcoplasmic anion channels. Our current study tested the opposite intervention, muscle cooling, and observed the opposite response. Oxidant activity dropped by half when muscle fibers were cooled from 37°C to 23°C. Thus, between 23°C and 43°C, oxidant activity within skeletal muscle fibers appears directly related to temperature.
Our findings are relevant to physiology in cold environments, where muscles may function at subphysiological temperatures in vivo. For example, our data are consistent with the report by Pendergast and coworkers (38) that nitric oxide production by exercising humans is diminished by lowering core temperature. The current data also relate to basic research conducted using isolated muscle preparations. Such studies are commonly conducted at 23°C to promote stability (47). Our data indicate that this strategy blunts endogenous oxidant activity, an “antioxidant effect” that biases the experimental system against oxidant-mediated processes. For example, Diaz et al. (10) showed that antioxidant inhibition of muscle fatigue, a common finding at 37°C (10), is abolished at room temperature. The current data suggest that by decreasing oxidant activity, muscle cooling lessens the role of muscle-derived oxidants in the fatigue process and renders antioxidants ineffective.
CO2 and Oxidant Activity
CO2 levels in skeletal muscle are altered under a variety of conditions, most notably exercise (40) and chronic obstructive lung disease (1). Changes in dissolved CO2 alter intracellular oxidant activity via a complex chemistry that is not well studied in muscle. In general, CO2 and bicarbonate react with peroxynitrite and other NO derivatives to influence the kinetics and end-products of NO metabolism. Via these and related pathways, CO2 can undergo electron exchange reactions to form carbonate radicals (28). Dissolved CO2 also influences the pH of biological systems, as observed in this study, and extracellular acidosis increases cellular production of both ROS and NO (8). Studies in nonmuscle cell types are conflicted about the net effect of CO2-mediated reactions, which are reported both to exaggerate NO-mediated injury (23, 37) and protect against nitrosative and oxidative stress (9, 12, 52).
The only prior study of CO2 effects on redox homeostasis in skeletal muscle was conducted by Stofan and associates (47). They measured reduction of cytochrome c in the arterial perfusate of a rat diaphragm preparation. Their data show that repetitive isometric contraction accelerates cytochrome c reduction. This signal was inhibited by SOD, identifying the reductant as superoxide anions. The signal was also inhibited by increasing CO2 levels in the superfusate. They concluded that elevated CO2 levels inhibit ROS release by exercising diaphragm.
We saw the opposite response. CO2 promoted oxidant activity in the cytosol, an effect most apparent at levels below 5% CO2. This apparent conflict can be resolved based on compartmentalization of the two assays used in these studies. CO2 reacts with NO and other oxidants to form redox-active intermediates, e.g., peroxynitrite and carbonate radicals, that are more unstable, have shorter half-lives, and diffuse shorter distances than the parent molecules. This tends to localize nitrosative and oxidative reactions near the site(s) of production, which are likely within the cell. Thus reactions with intracellular targets become more likely, enhancing our signal, and diffusion out of cells becomes less likely, lessening the signal measured by Stofan et al. (47).
Contributions of Myogenic ROS and NO
Healthy skeletal muscle continually produces ROS and NO at low levels under resting conditions (4, 5, 32, 45) and at higher levels during contractile activity (4, 45). Myogenic ROS and NO modulate intracellular processes, including excitation-contraction coupling (39, 42, 43, 47), acute fatigue (4, 21, 41, 50), glucose uptake (16), and gene expression (27). Yet virtually all of the data in this field derive from measurements made in the extracellular space (1a, 6, 21, 29, 45, 55, 56) or microvasculature (19, 47). This seriously limits our understanding of redox homeostasis in muscle because free radical kinetics in cellular systems are compartmentalized and difficult to model (33).
Our current data confirm prior reports of myogenic ROS activity in resting muscle fibers (4, 5, 11, 17, 18, 32, 34, 35, 41). The ROS signal represented approximately half of the total oxidant activity that we measured. Inhibitory effects of SOD and catalase demonstrate the contributions of superoxide anions and hydrogen peroxide, respectively. We do not expect SOD (molecular mass 125 kDa) or catalase (65 kDa) to have entered muscle fibers in significant amounts. Rather, we postulate that enzyme activity created a perisarcolemmal “sink” for superoxide anions and hydrogen peroxide, lowering ROS concentrations in the near-membrane extracellular compartment and promoting outward flux from the cytosol. Ebselen, a glutathione peroxidase mimic that enters the cell (24, 36), had a similar inhibitory effect. Hydrogen peroxide also crosses the sarcolemma and had the opposite effect, increasing oxidant activity. In combination, these data suggest that DCF fluorescence primarily reflected the activity of cytoplasmic hydrogen peroxide or its derivatives.
NO activity has not been detected in muscle fibers previously. In this study, we used complementary interventions to identify the NO signal. These included NO synthase inhibitors, an NO scavenger, and the substrate for NO synthesis.
Like SOD and catalase, the NO scavenger hemoglobin (molecular mass 64 kDa) is expected to have remained outside the cell, creating an extracellular sink for NO derivatives and promoting outward diffusion. The other reagents are known to cross membranes freely. Each NO-selective reagent altered cytosolic oxidant activity as predicted. Specificity of the observed changes is supported by negative control studies using d-enantiomers and by positive effects of an NO donor. Overall, our results indicate the contribution of NO derivatives to total oxidant activity is ∼40%, similar in magnitude to the contribution of muscle-derived ROS.
The methods and experimental interventions used in this study are established techniques with which we have prior experience. Under the current conditions, fiber bundles isolated from rat diaphragm are stable for at least 60 min, as reflected by decrements in maximal force of <10%/h (32). Fiber bundles also remain stable after DCFH loading (32).
Photooxidation is a major artifact of DCFH use (41) that we control by conducting experiments in a darkened laboratory, by acquiring each data point from a unique site on the fiber bundle surface, and by standardizing excitation stimuli (intensity, duration, timing, and total number). We used photooxidation as a tool to determine that DCFH loading of muscle fibers is complete within 40–45 min and that intracellular DCFH availability does not limit the biological signal.
In the absence of muscle fibers, DCFH spontaneously autoxidizes to DCF at a rate 1–2 orders of magnitude less than the biological signal. Our current data indicate autoxidation is more prominent at 23°C than 37°C, is accelerated by exposure to hydrogen peroxide or an NO donor, and is insensitive to other drugs that we used to test free radical activity. DCF fluorescence shows pH sensitivity, decreasing with acidosis. In the current protocols, condition-matched controls were used to correct for autoxidation artifact in the final data expression.
In conclusion, oxidant activity in skeletal muscle fibers is strongly influenced by common physiological variables. Basal activity is diminished by cooling or CO2 depletion, environmental factors that are regulated in experimental systems. Studies of isolated muscle preparations at room temperature or in CO2-deficient buffer systems are likely to underestimate the magnitude of oxidant-mediated effects in muscle. Myogenic ROS and myogenic NO appear to influence cytosolic oxidant activity to a similar extent. This is the first explicit demonstration that ROS and NO signaling cascades are simultaneously active in the cytosol of muscle fibers. It reinforces the biological importance of ROS and NO interactions and the potential interplay of these two cascades in modulating intracellular events.
This work was supported by the National Space Biomedical Research Institute, the Muscular Dystrophy Association, and National Heart, Lung, and Blood Institute Grant HL-45721.
We thank Dr. J. Moylan for assistance in editing this manuscript.
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