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1The Dorothy M. Davis Heart and Lung Research Institute, and Departments of 2Internal Medicine (Division of Pulmonary, Critical Care and Sleep Medicine), 3Emergency Medicine, and 4Molecular and Cellular Biochemistry, The Ohio State University, Columbus, Ohio
Submitted 16 September 2004 ; accepted in final form 17 December 2004
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ABSTRACT |
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 TOP ABSTRACT METHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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10% of control. There were no consistent influences of the AOX treatments on high-energy phosphates during hypoxia. The results demonstrate that superoxide scavengers can protect contractile function and reduce contracture in hypoxia through a mechanism that does not involve preservation of high-energy phosphates. skeletal muscle; 4,5-dihydroxy-1,3-benzenedisulfonic acid; N-acetylcysteine; Mn(III)tetrakis(1-methyl-4-pyridyl) porphyrin pentachloride; calcium regulation
In this study, the hypothesis was tested that the mechanism by which antioxidants (AOXs) influence contractile function in hypoxia is through preservation of high-energy phosphates [ATP and creatine phosphate (CrP)] and reduction of Pi. There is considerable rationale for this hypothesis, because glucose transport, glycolysis (5), and oxidative phosphorylation (26) have all been shown to be inhibited by exposure to elevated levels of oxidants. Furthermore, oxidants such as peroxynitrite (ONOO–) appear to inhibit glycolysis in heart muscle during ischemia (10, 11). In hearts treated with the superoxide (O2–·) scavenger Tiron, CrP levels are preserved in rat cardiac muscle at the end of an ischemic exposure (25). The hypothesis was addressed in this study by measuring high-energy phosphates, at the end of a 30-min exposure to severe hypoxia in the presence or absence of various AOXs.
Another purpose of this study was to evaluate the influence of antioxidant treatment on the phenomenon of "contracture," a term used to describe the gradual increase in passive force during exposure to severe hypoxia. Contracture in skeletal muscle is modest compared with cardiac muscle but resembles behavior similar to that seen in ischemic heart, where it is believed to reflect responses to elevated free sarcomeric Ca2+ (22).
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METHODS |
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TOPABSTRACTMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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5 mm in width, with portions of the central tendon and ribs attached. The strips were mounted vertically in four tissue baths with oxygenated Ringer's solution and maintained at 37°C. Optimal length and maximum field stimulation current were determined for each strip by standard methods (34). Strips were thermo-equilibrated for 15 min while being intermittently stimulated with one twitch every 20 s. Then initial, untreated force-frequency relationships were determined by using a twitch stimulation and stimulations at 20, 30, 40, 50, 60, 80, 100, and 150 Hz, each maintained for 400 ms, with a 20-s recovery interval between stimulations. Before hypoxia, preloads (passive tensions) were reset, when necessary, to the predetermined level obtained at baseline optimal length determination (Fig. 1). Once the tissues were exposed to hypoxia, there was no adjustment of passive force, because it was one of the measured variables.
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15 min in well-bubbled buffer at 37°C to avoid excessive oxidation (yellow color), while providing sufficient time for the CO2-HCO3– buffer system to compensate for its mild acidification effect. Antioxidant equilibration and hypoxia challenge. The experimental protocol is illustrated in Fig. 1. Tissues were equilibrated in their respective treatment buffers for 30 min (Fig. 1, AOX Equilibration) while being stimulated with intermittent twitches (0.05 Hz). A new force-frequency maneuver was then performed at the end of the AOX incubation period (Fig. 1, Treatment FF). The strips were rested for 10 min and the preload was readjusted (Fig. 1, arrow). For the AOX experiments, each strip was put into one of four groups: 1) untreated time control in 95% O2 (control), 2) AOX treated in 95% O2 (control + AOX), 3) untreated hypoxic (hypoxia), and 4) AOX treated hypoxic (hypoxia + AOX). To test the response to hypoxia, the gas going to the hypoxia baths was changed to 95% N2-5% CO2 (hypoxia), and the buffers were replaced with buffer preequilibrated with 95% N2-5% CO2 at 37°C. The nonhypoxic baths (with and without AOX treatment) were changed to fresh buffer, preequilibrated with 95% O2-5% CO2 at 37°C (control). By use of a remote Clark PO2 detector (World Precision Instruments, Sarasota, FL), measurement of the PO2 within the hypoxic baths demonstrated values from 5–15 Torr near the center of the chamber after complete equilibration. The tissues remained in hypoxia or 95% O2 for 30 min. For the experimental series in which the extracellular Ca2+ was altered, the muscles were treated similarly, but there were no 95% O2 controls performed; the four muscles were exposed to four different Ca2+ conditions in hypoxia.
For the AOX treatments, two different stimulation paradigms were performed. In one paradigm (stimulated), muscles were intermittently stimulated with twitches (0.05 Hz) during hypoxic exposure to monitor contractile function during the time course of developing hypoxia, followed by a force-frequency test at the end of the hypoxic exposure. In the second paradigm (unstimulated), no twitch stimulations or force-frequency measurements were performed at any time during hypoxia. This latter group was necessary to eliminate the complexities of interpretation due to differing energy costs of contraction between AOX groups, i.e., because AOX-treated tissues generated more force in hypoxia, they presumably consumed more energy. All tissues used in the energetics study were rapidly freeze clamped across the center of the diaphragm muscle mass at the end of the hypoxic exposure period and then stored at –80°C. Our procedures allow us to freeze clamp the muscle within 1 s of dropping the surrounding tissue bath (24).
Extracellular Ca2+ experiments. A separate series of experiments, designed to test the influence of extracellular Ca2+ on passive force during hypoxia, were performed in which [Ca2+] in the baths was manipulated (brackets denote concentration). Four diaphragm strips were mounted, and after the initial force-frequency, the baths were changed to control Ca2+ (2 mM), low Ca2+ (0 mM), 0 mM Ca2+ + 50 µM EGTA, or double Ca2+ (4 mM). To keep osmolarity approximately constant, changes in [NaCl] were used to compensate for changes in CaCl2.
Tissue extraction. Frozen samples were weighed and ground in the presence of 0.6 N perchloric acid under liquid N2 by using liquid N2-cooled mortars and pestles. The N2 was allowed to evaporate, and the resulting powder was quickly placed in a vial containing additional ice-cold perchloric acid. Samples were vortexed and then centrifuged at 16,000 g for 1 min. One milliliter of supernatant was pipetted to a new tube, and the pellet was kept for protein analysis. To adjust the sample pH, 1 M K2CO3 was added to the supernatant and the sample was allowed to sit on ice for >30 min to precipitate KClO4 and then centrifuged. The final supernatant was pipetted into aliquots and stored at –80°C. All added volumes and weights were recorded for later analyses.
Metabolite analysis. Pi was assayed by a modified colorimetric method by Fiske and Subbarow (17a). Briefly, 50 µl of thawed supernatant were added to 1.0 ml water and 1.0 ml assay buffer (0.5% ammonium molybdate, 2.0% ascorbic acid, and 2.8% concentrated sulfuric acid) and mixed, and the absorbance was read after 75 s at 820 nm on a spectrophotometer (Hewlett-Packard 8452A diode array). Concentrations were calculated on the basis of a standard curve. ATP and CrP are acid labile, so samples of known concentration were also measured to determine the portion of the signal attributable to their degradation. Timing of measurements was critical because of this degradation. Creatine, ATP, and CrP were assayed by the enzymatic methods of Bergmeyer (4). This procedure was described in detail in a previous manuscript (24). All assays were performed in duplicate and repeated if measurements varied by >15%. Final values were the means of the two closest values.
Protein assay. Stored protein pellets were thawed and washed with acetone. Dried pellets were resuspended in 500 µl of 1 N NaOH and heated for 10 min at 95–100°C, then placed on ice or frozen at –20°C until assayed (standards were treated similarly). Total dissolved protein was determined by the Lowry method (32). Samples were read with a spectrophotometer at 750 nm and compared with a standard curve.
Statistical analysis. Force measurements were always normalized to initial maximum force. Specific force, or stress, could not be determined in this study because tissues were immediately freeze clamped at the end of the hypoxic period, eliminating the ability to calculate the length and weight of the muscle. The assumption used was that before any treatments all tissues would exhibit equivalent baseline specific forces. Most analyses then used repeated-measures one-way or two-way ANOVA using JMP software (SAS Institute). Comparisons between means were tested, post-ANOVA, by use of contrast procedures. To evaluate differences between the amounts of contracture at the end of the hypoxic period, paired differences between matched controls were compared by using nonparametric matched-pairs statistics. Data points were expressed graphically as the average value ± SE.
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RESULTS |
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85% of matched control values) compared with high frequencies (150-Hz force was reduced only 50–60% of matched control values). Antioxidant treatment in strips not exposed to hypoxia (open circles) resulted in reductions in force at nearly all frequencies, suggesting that, in this setting, high levels of AOXs may have a long-term (>1 h exposure) negative impact on overall contractile function. This did not reach statistical significance for NAC-treated tissues, except at low frequencies.
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High-energy phosphate measurements in hypoxia.
High-energy phosphates and Pi were measured in fresh frozen samples, strips that were stimulated during hypoxia, and strips that were not stimulated during hypoxia. Results are illustrated in Fig. 7. Data for NAC experiments are not shown for simplicity, because there was no protection of contractile function and they could not be used to test the primary hypothesis of the study. However, measurements were performed for NAC-treated muscles (10 mM) contracting during hypoxia. No significant effects of NAC treatment on ATP, CrP, or Pi were observed (not shown). In Fig. 7A, the [ATP] [0.05 µmol/mg protein, corresponds to 3.5 µmol/g wet wt (24)] of fresh frozen diaphragm (open bars) was similar to control diaphragm strips incubated in the tissue baths. In hypoxic muscle strips that were not stimulated during hypoxia there was a small trend for [ATP] to decrease, but this did not reach statistical significance. However, strips that were electrically stimulated to twitch and to generate a force-frequency relationship during hypoxia showed significant (20–30%) reductions in [ATP] compared with matched, stimulated controls. Only Tiron treatment during hypoxia resulted in no significant reduction in [ATP] compared with matched controls in these tissues. When the values for hypoxia and hypoxia + Tiron were tested against each other, however, there were no differences in these groups. In assessing the overall response of all tissue groups, the lack of effect by hypoxia in Tiron-treated strips appears to be similar to the results for other antioxidant treatments.
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0.15 µmol/mg protein, corresponding to 11.6 µmol/g wet wt (24). Hypoxia exposure drastically reduced CrP to 10% of normoxic values. There were no significant effects of any AOX treatment compared with nontreated control tissues, although in unstimulated tissues CrP tended to remain higher at the end of hypoxia compared with the stimulated strips. As shown in Fig. 7C, Pi increased significantly during hypoxia, approaching values corresponding to the reduction in CrP. There were no significant effects of antioxidant treatment on Pi in any of the tissues studied.
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DISCUSSION |
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30% higher maximum tetanic forces compared with untreated tissues, with no apparent additional reserves of high-energy phosphates. Therefore, the primary hypothesis of this study was disproved. Antioxidants also inhibited the development of hypoxia-induced contracture. Because contracture was shown to be dependent on extracellular Ca2+, implying a Ca2+-dependent mechanism, the results suggest that O2–· scavengers may impact hypoxia-induced alterations in Ca2+ regulation. Comparison with previous studies and critique of the approach. The results for the effects of AOXs on contractile function confirm our preceding work with Tiron under similar conditions (34). However, in contrast to our previous work, NAC had little or no influence on tetanic force production in hypoxia. A number of exploratory experiments were performed in an attempt to resolve this discrepancy within our laboratory (data not shown), namely 1) several different formulations and concentrations of NAC were compared, 2) differences in the time of NAC incubation were evaluated (30 min–2 h), and 3) the original fabricated field-stimulating electrodes were tested against currently employed commercial electrodes because of the potential effects of metal chemistry in buffer systems. None of these experiments reconciled the differences in results. We conclude that some unknown variable, e.g., the source of animals, diet, the source of distilled water, or metal-contaminating buffer salts had changed between the sets of experiments. NAC did have the expected influence on twitch force, before hypoxia, as shown in Fig. 5, confirming the previously described pharmacological effects of this agent on skeletal muscle (12, 28, 34).
The present results differ from the recent work of Heunks et al. (23), who found that AOXs, Tiron, and NAC depressed maximum force development during hypoxia compared with hypoxia alone. Differences in the experimental protocols between the two studies suggest that the level of hypoxia was much more severe in the present study. First, bath temperature was 26°C in Heunks et al. vs. 37°C in the present study. Therefore, metabolic rate would be higher and dissolved O2 lower at 37°C, predictably making the tissues in this experiment more hypoxic in the central core of the muscle bundle. Second, the PO2 was 7.5 kPa (56 Torr) for 60 min vs. 1–2 kPa (5.0–15.0 Torr) for 30 min in this study. Third, the average animal size in the Heunks study was 300 g vs. 400 g in the present study, which could have resulted in thicker diaphragm tissues and a greater core hypoxia in the present study. Therefore, it is not surprising that the maximum tetanic force we observed after 30 min of hypoxia was 50% of baseline compared with 90% of baseline in the study of Heunks et al. after 60 min of hypoxia.
As pointed out in a number of recent publications (15, 16), the redox-dependent responses to hypoxia in skeletal muscle vary widely within a relatively narrow range of PO2 exposure. A rough estimate of the PO2 as a function of tissue depth in isolated skeletal muscle is provided from work by Eu et al. (15) in resting mouse extensor digitorum longus muscle, which is 1 mm in thickness, compared with 0.4–0.6 mm in rat diaphragm. When bath PO2 is 1% O2, the PO2 at the core of the extensor digitorum longus was 3–4 Torr; at a 20% O2 core PO2 was 40 Torr; at 95% O2 core PO2 was 387 Torr. These measurements are extremely difficult to make accurately and should be viewed with some caution; nevertheless they suggest that, in a hypoxic tissue bath, PO2 reductions in the muscle core can be substantial. Normal PO2 in perfused skeletal muscle is believed to be 10 Torr at rest and 3–5 Torr during intense exercise (38), not reaching a "critical" cell PO2 where respiration is limited until 1.5 Torr (39). The present study likely resulted in relatively severe hypoxia in the muscle core with more moderate hypoxia in the study by Heunks et al. (23). We speculate that different response mechanisms predominate at different levels of severity of hypoxia and therefore it is difficult but interesting to compare the results from the two laboratories.
Influences of specific AOXs. It is of some interest that the two O2–· scavengers had such potent impacts on contractile function in this setting whereas NAC, a more commonly used antioxidant or reducing agent, had only small effects. O2–· is not considered to be extremely reactive as an oxidant and behaves as a reductant in many microenvironments. However, some targets, such as isoforms of the sarcoplasmic reticulum (SR) Ca2+-ATPase, appear to be specifically sensitive to O2–· oxidation (3) whereas other targets may be uniquely sensitive to peroxynitrite, resulting from reactions that require O2–·.
It is instructive to compare the relative scavenging abilities of Tiron and MnTMPyP to the endogenous enzyme, SOD. Endogenous SOD activity in rat diaphragm has been estimated to be 3.5 U/mg (30). By taking the approach of Gardner et al. (19), assuming a soluble protein concentration of 80 mg/ml in diaphragm (this study), the endogenous SOD concentration can be estimated at 2.5 µM. Because SOD has a reaction rate with O2–· of 109 M–1·s–1 it would scavenge at a rate of 2.5 x 103 s–1. By contrast, Tiron scavenges O2–· with a rate constant of 5 x 108 M–1·s–1 (21), and, assuming a 10 mM intracellular concentration, it would scavenge O2–· at 5 x 106 s–1, a 2,000 times greater rate than endogenous SOD. Similarly, assuming that MnTMPyP is kept in the reduced state [Mn(II)·TMPyP] by cellular reductants, the Mn(II)TMPyP has a rate constant of 4 x 109 M–1·s–1 for oxidation by O2–· (17) and intracellular concentrations of 50 µM would result in a net reaction rate of 2 x 105 s–1, 80-fold higher SOD activity than endogenous SOD. It is highly likely that intracellular concentrations of these applied scavengers are considerably less than their extracellular concentrations and that endogenous SOD is more effective than suggested by these calculations because it is strategically located near O2–·-generating sites such as the mitochondria. Nevertheless, the calculations demonstrate that the scavengers used in this study were extremely potent at the concentrations used. The lack of effect of NAC on the response may simply reflect its comparatively reduced scavenging ability. It is known primarily as a hydroxyl scavenger and precursor for glutathione synthesis. It is not known to have significant scavenging effects on O2–· and only slowly reacts with H2O2 (2).
It is also important to note that neither of the O2–· scavengers is particularly specific. Tiron is an effective metal scavenger (27) and MnTMPyP acts as both a catalase (7) and a peroxynitrite scavenger (18) as well. We infer that it is likely that these AOXs affected a variety of reactive O2 and reactive nitrogen species and therefore the specificity of O2–· in this physiological response to hypoxia should be viewed with some caution.
Potential mechanisms of antioxidant inhibition of contracture. The results showed that contracture could be eliminated by removal of extracellular Ca2+, suggesting that the phenomenon is related to hypoxia-induced alterations in cell Ca2+ homeostasis. Several known mechanisms could account for the influence of AOXs on Ca2+ homeostasis. First, movement of Ca2+ into the cytosol would be expected in response to membrane depolarization, from either the SR or possibly the extracellular fluid. Membrane depolarization is known to occur in rat diaphragm during hypoxia (14). Second, oxidants could influence K+ channel behavior in hypoxia and thus affect Ca2+ influx indirectly. In general, the membrane resists depolarization by outward K+ leakage and recruitment of new K+ channels. One of the molecular mechanisms believed to be responsible for O2 sensing is the influence of local redox state on K+ channel behavior (33), although little is known regarding these influences in skeletal muscle. Third, the SR Ca2+ release channel in skeletal muscle (RyR1) is sensitive to the local redox environment. The open probability of the channel is highly dependent on complex interactions between oxidants, O2, and NO (1, 15, 16, 40). This effect has been described as a form of O2 sensing by RyR1 (15, 16). Fourth, the SR Ca2+-ATPase is responsible for clearance of cytosolic Ca2+ in skeletal muscle, and it is dependent on the presence of an effective creatine kinase system and available CrP (8, 13). Because CrP dropped below 10% of normal during hypoxia, a value shown to inhibit SR Ca2+-ATPase activity in skinned skeletal muscle (13), it is likely that low Ca2+-ATPase pump activity contributed to the accumulation of cytosolic Ca2+. Nearly all isoforms of the Ca2+-ATPase are redox sensitive, with oxidants causing a reduction in function (3, 41). Recent evidence suggests that at least some isoforms of Ca2+-ATPase in skeletal muscle are uniquely inhibited by O2–· (3). Fifth, cell swelling may also contribute to the elevation in passive force during hypoxia. Presumably, as muscle fibers swell, stresses on the elastic elements of the cytoskeleton and membrane could exert passive forces along the long axis of the fiber, contributing to contracture. Swelling occurs in the skeletal muscle in the setting of hypoxia due to the additional osmotic pressure caused by the splitting of CrP to Pi and Cr and the degradation of glycogen to glucose and eventually lactate (29). Interestingly, cell swelling causes an increase in intracellular Ca2+ and subsequent reactive O2 formation via a phospholipase A2-dependent pathway (36).
Mechanisms of tetanic force preservation in hypoxia. Because AOXs did not preserve high-energy phosphates, other mechanisms for preservation of contractile function must be considered. One possible explanation is that AOXs preserved tetanic force in hypoxia by allowing the muscles to contract more economically. Economy is defined as tetanic force developed per rate of ATP hydrolysis. Although the specific rate of ATP hydrolysis was not measured during contraction, the data suggest that there was little or no apparent additional energetic cost to contract at higher forces with AOX treatment because there was no additional decrement in [ATP] and [CrP]. This conclusion is somewhat limited by the inherent inaccuracies involved in the measurement of high-energy phosphates in frozen samples, as discussed previously (24), and because of the fact that the actual rate of hydrolysis was not measured. Furthermore, we cannot rule out the possibility that the rate of ATP turnover was elevated in AOX-treated tissues. It is possible that at specific levels of CrP, ATP, and ADP during hypoxia, the metabolic network is able to maintain better coupling of energy utilization and energy production.
One potential mechanism that could account for improved economy of contraction in hypoxia is a more homogeneous distribution of forces along the fiber length. For example, if certain regions of a muscle, in series along a fiber, are poorly activated during hypoxia compared with other better activated areas, the activated areas will shorten disproportionately and stretch the noncontracting portions of the fibers, thus reducing the net force transmission from tendon to tendon. In hypoxic muscle, it is likely that some regions of a fiber are more vulnerable to energy depletion. Recent data suggest that within sarcomeric regions there is a compartmentalization of energy metabolic systems, at least in cardiac muscle (35), and this could vary with depth inside the tissue and fiber. Alternatively, some other less defined redox-sensitive mechanism that influences the efficiency of cross-bridge interaction could be responsible for the effects of antioxidant treatment. For example, the downward shift in the force-velocity relationship (a characteristic of slower more efficient muscles) was observed with Tiron treatment in more moderate hypoxia (23), suggesting that such a mechanism is possible.
Some of the same potential influences of AOXs on Ca2+ regulation that were discussed in the previous section regarding contracture may also apply to the development of tetanic force in hypoxia. For example, oxidative inhibition of Ca2+-ATPase (3, 41) would presumably lead to reductions in SR Ca2+ stores in hypoxia and therefore reductions in maximum Ca2+ release and maximum force development during contraction. Similarly, oxidation of ion channel activity may limit propagation of the action potential down T tubules, resulting in poor distribution of excitation-contraction coupling in some areas of the muscle. Antioxidants could work in hypoxia by blocking these effects, and the result would be that the muscle would contract in a more homogenous way and presumably do so more economically.
Physiological implications. It is simplistic to conclude that hypoxia represents a form of "oxidative stress" in which AOXs are protective. It is probably more reasonable to consider the low level of oxidants produced in hypoxia as one of many signaling events that are necessary to set up some secondary protective system. For example, is it a good thing to be able to generate more force in a severe hypoxic environment when ATP and CrP are depleted as were observed in the presence of AOXs in this study? Contractions during hypoxia have been shown to cause damage to muscle proteins (9). From the data, it is clear that simply performing a force-frequency curve in hypoxia results in significant decrements in [ATP], whereas [ATP] is reasonably protected in the rested hypoxic muscle. Much like mechanisms of fatigue, muscles are likely to be programmed to protect themselves from overexertion and injury in such environments. For example, in a previous study in heart muscle by Klawitter et al. (25), Tiron was particularly effective in preserving high-energy phosphates in the heart when administered during 20 min of ischemia. However, it resulted in large decrements in energy status in the reperfusion phase, suggesting that oxidants produced during ischemia represent a desired protective mechanism in the heart, possibly providing a type of preconditioning stimulus. Similar mechanisms may be present in skeletal muscle but possibly working through pathways that differ from those in the heart.
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ACKNOWLEDGMENTS |
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Present address of P. Klawitter: Dept. of Emergency Medicine, Upstate Medical Univ., 750 E. Adams St., Syracuse, NY 13210.
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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