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1 Institute for Biomedical Sciences, School of Medical Sciences, University of Sydney F13, NSW 2006, Australia
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Abstract |
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(Received 19 January 2005;
accepted after revision 16 February 2005;
first published online 17 February 2005)
Corresponding author D. G. Allen: Institute for Biomedical Sciences, School of Medical Sciences, University of Sydney F13, NSW 2006, Australia. Email: davida{at}physiol.usyd.edu.au
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Introduction |
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One approach to identifying the mechanisms of muscle fatigue is the use of single muscle fibres loaded with a calcium indicator. In this preparation the decline of isometric force during fatigue can be assigned to three different categories: (i) changes in maximum Ca2+-activated force, (ii) changes in the intracellular calcium during tetani (tetanic [Ca2+]i), and (iii) changes in the Ca2+ sensitivity of the contractile apparatus (Westerblad & Allen, 1991). At room temperature the fatigue caused by intermittent tetani in mouse skeletal muscle fibres has contributions from all three of these categories. It is probable that the well recognized increase in inorganic phosphate concentration ([Pi]i) and acidosis that occur in muscle fatigue are responsible for the reduced maximum Ca2+-activated force and the reduced Ca2+ sensitivity (Fabiato & Fabiato, 1978; Godt & Nosek, 1989). The reduced [Ca2+]i during tetani is thought to result from reduced SR Ca2+ release perhaps caused by precipitation of calcium phosphate in the SR (Fryer et al. 1995; Allen & Westerblad, 2001) or perhaps by the inhibitory effects of raised Mg2+ and reduced ATP on SR Ca2+ release (Dutka & Lamb, 2004).
Many studies of fatigue on isolated mammalian muscle have been performed at room temperature both for reasons of convenience and because isolated skeletal muscles sometimes perform poorly or are unstable at 37°C (Lännergren & Westerblad, 1987). However, in the past decade it has become clear that studies at room temperature may be misleading when extrapolated to 37°C. For instance intracellular acidosis was proposed to reduce force in fatigue by reducing the Ca2+ sensitivity of the contractile proteins (Fabiato & Fabiato, 1978). However the original study was performed at room temperature and when these experiments were repeated nearer 37°C it was found that acidosis under these conditions had very little effect on Ca2+ sensitivity or on muscle fatigue (Pate et al. 1995; Westerblad et al. 1997). Similarly the effects of phosphate are much smaller at 30°C compared to 15°C (Debold et al. 2004). These studies suggest that the effects of metabolites on the contractile proteins may be substantially smaller at 37°C than predicted from studies at lower temperatures.
There is evidence that fatigue develops more rapidly at 37°C than at lower temperatures, raising the possibility that different mechanisms might be involved (de Ruiter & de Haan, 2000). Reactive oxygen species (ROS) have been implicated as these are produced at a higher rate at 37°C compared to room temperature (Arbogast & Reid, 2004) and because ROS scavengers can slow the time course of fatigue at 37°C (Khawli & Reid, 1994). However the mechanism by which ROS might accelerate fatigue is unclear, though effects on SR Ca2+ release have been proposed (Reid, 2001). Studies of the damaging effects of very high temperatures (43–47°C) on muscle function, which are also mediated by ROS, show a pronounced reduction in the maximum Ca2+-activated force (van der Poel & Stephenson, 2002).
In the present study we compared the rate of fatigue at room temperature with that at 37°C in mouse skeletal muscle. We found that muscles fatigued much faster at 37°C than room temperature, but this effect was abolished by the ROS scavenger 4,5-dihydroxy-1,3-benzene-disulphonic acid (Tiron). Using a calcium indicator, indo-1, we show that the accelerated fatigue at 37°C is not caused by changes in tetanic [Ca2+]i or maximum Ca2+-activated force but can be almost entirely explained by a rapid reduction of Ca2+ sensitivity.
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Methods |
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In experiments designed to test the role of reactive oxygen species (ROS) we used the membrane-permeant ROS scavenger 4,5-dihydroxy-1,3-benzene-disulphonic acid (Tiron; Sigma, Australia) (van der Poel & Stephenson, 2002). The study by van der Poel & Stephenson used 20 mM Tiron but we found this concentration rendered the muscles inexcitable. Nevertheless, when 20 mM Tiron was applied during the resting period preceding the fatigue protocol and then washed off, this procedure protected the muscle against the more rapid fatigue observed at 37°C (data not shown). We found that 5 mM Tiron did not affect the contraction of unfatigued muscle but protected against the effects of high temperature when present throughout the fatigue protocol. In a smaller number of experiments we used a second ROS scavenger, 4-hydroxy-2,2,6,6-tetramethylpiperidine-1-oxyl (Tempol; Sigma, Australia).
Experimental protocol
Muscles were stimulated with platinum electrodes using pulses of 0.5 ms at an intensity of about 1.2 x threshold. All tetanic contractions were 0.4 s in duration. We used a fatigue protocol in which the muscles were stimulated with 100 Hz tetani repeated every 4 s for 2 min, and the interval reduced every 2 min thereafter to 3.5 s, 3 s, and so on. The protocol was stopped when force was less than 50% of the initial level. Fatigue protocols were separated by 45 min, which we found was sufficient for complete recovery at 22°C (see also Chin & Allen, 1997). The rate of fatigue was characterized by the time taken for force to fall to 50% of the initial value (T1/1).
Each experiment consisted of at least two fatigue protocols. The initial protocol was always carried out at 22°C (control) and the second protocol was carried out with or without 5 mM Tiron at either 22°C or 37°C. Each muscle was compared with its own control to reduce the intrinsic variability between muscle bundles and single fibres. When Tiron (or Tempol) was used it was applied immediately after the first fatigue protocol and continued throughout the second fatigue protocol. In experiments at 37°C the temperature perfusing the muscle bath was raised over 3–5 min just before the start of the next fatigue protocol.
Use of Indo-1 to measure intracellular [Ca2+]i
The fluorescent Ca2+ indicator, indo-1, was pressure-injected into single fibre preparations to measure [Ca2+]i (for details see Westerblad & Allen, 1991). The injected fibre was illuminated at 360 nm and emitted light measured at two wavelengths (400 and 505 nm) with photomultiplier tubes. The signals were then passed to an analog divide circuit which produced the 400/505 nm ratio signal. Background signal was subtracted from each measured wavelength prior to ratio calculation.
The ratio signal obtained during the experiments was later converted to [Ca2+]i using the following equation:
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Determination of Fmax and Ca50
To determine the calcium sensitivity of the contractile proteins (Ca50) and the maximum Ca2+-activated force (Fmax), we used the protocol initially described by Westerblad & Allen (1991). This protocol consisted of activating single fibre preparations with 0.4 s tetani at a range of frequencies (20, 30, 50, 70, 100 Hz and 100 Hz in the presence of 10 mM caffeine) at 1 min intervals prior to the fatigue protocol. In addition caffeine (10 mM) was rapidly applied after the last tetanus during the fatigue protocol (i.e. when the force had fallen to < 50%) and two or three additional tetani subsequently recorded. For all tetani the tetanic [Ca2+]i and force were estimated by averaging the values over the final 200 ms of the tetanus when the [Ca2+]i and the force appeared to have reached their steady-state relationship. The control (unfatigued) Ca50,CON and Fmax,CON were obtained by plotting the tetanic force versus
[Ca2+]i at each frequency of stimulation. The resulting force–Ca2+ plots were fitted using a Hill equation (see Fig. 5):
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All values are expressed as the mean ± S.E.M. Student's paired t test was used to test for statistical significance between fatigue bouts within each experiment. Significance was accepted at P < 0.05. The Mann-Witney rank sum test was used when a non-parametric test was required (see Table 1).
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Results |
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The initial studies were performed on muscle bundles since these were much more robust, particularly at 37°C. In the first series (n = 6), the bundle was fatigued at 22°C and, after 45 min for recovery, the fatigue was repeated in the presence of 5 mM Tiron (Fig. 1A). The second fatigue run, either in the absence of Tiron (not shown) or in its presence, was not obviously different from the control. The T1/1 of the first fatigue series was 7.7 ± 1.1 min while that of the second fatigue at 22°C + Tiron was similar at 114 ± 12%. There was no change in initial force in the second fatigue run in this series (99 ± 9% of control initial force).
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To test whether the increased rate of fatigue at 37°C was caused by the effects of ROS we performed a third series of experiments (n = 5) in which the bundle was treated with 5 mM Tiron before the fatigue protocol at 37°C. Figure 1C shows that this treatment prevented the deleterious effects at 37°C. The T1/1 at 22°C was 5.5 ± 0.9 min while the T1/1 at 37°C in the presence of Tiron was not significantly different at 103 ± 16%. In addition the reduction in initial force was smaller and not statistically significant in the presence of Tiron (87 ± 7% control). In five experiments we tested the effects of a second ROS scavenger, 1 mM Tempol. The T1/1 at 37°C in the presence of Tempol was 94 ± 6% of the control at 22°C which is significantly longer (P < 0.01) than the T1/1 at 37°C in the absence of ROS scavengers. The initial force was also better maintained at 37°C in Tempol (94 ± 6% of the control at 22°C).
Figure 2 summarizes some of the above data with the T1/1 for each series normalized to 100%. This figure shows that 5 mM Tiron had no significant effect on the duration of fatigue at 22°C. At 37°C muscle bundles fatigue significantly more rapidly and this acceleration was eliminated in the presence of the ROS scavenger Tiron.
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The experiments with muscle bundles showed clearly that at 37°C fatigue was accelerated and that Tiron or Tempol could prevent this effect. The aim of the experiments with single fibres was to determine the mechanism of this accelerated fatigue, specifically to determine the contribution of changes in tetanic [Ca2+]i, myofibrillar Ca2+ sensitivity (Ca50) and maximum Ca2+-activated force (Fmax).
Figure 3 shows the [Ca2+]i and force records of a representative single fibre experiment from a group of seven subjected to the fatigue protocol at 22°C and at 37°C. Note that the fatigue protocol is preceded and followed immediately by application of 10 mM caffeine to determine Fmax. The time course of fatigue is much faster in single fibres than in whole bundles and possible reasons for this are considered in the Discussion. Figure 3 also shows that, as in bundles, force declined more rapidly during fatigue at 37°C compared to 22°C (T1/1 = 77 ± 11 s at 22°C and was reduced to 55 ± 5% at 37°C). Importantly, it was observed that tetanic [Ca2+]i remained more or less constant during fatigue at 37°C whereas at 22°C the previously described decline in tetanic [Ca2+]i was observed. In seven fibres at 22°C the tetanic [Ca2+]i at the end of fatigue was 62 ± 3% of that at the start. In contrast when the same fibres were fatigued at 37°C the decline in tetanic [Ca2+]i was reduced so that the tetanic [Ca2+]i at the end of fatigue was significantly larger at 88 ± 5% of initial value (see Table 1). In addition, at 22°C, the caffeine-activated tetanic [Ca2+]i at the end of fatigue is much reduced compared to the prefatigue value; in contrast at 37°C the caffeine-activated tetanic [Ca2+]i appears unchanged at the end of the accelerated fatigue. These observations all support the idea that decline in tetanic [Ca2+]i has some role in fatigue at 22°C but does not contribute to fatigue at 37°C.
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Figure 4 shows the results one of six experiments in which a control fatigue protocol was performed at 22°C followed by a second fatigue at 37°C + 5 mM Tiron. Note that, as in bundles, the rate of fatigue is now fairly similar at the two temperatures. The T1/1 was 131 ± 37 s at 22°C and was unchanged at 102 ± 3% at 37°C + Tiron. Also, tetanic [Ca2+]i displayed the same declining trend during fatigue at 37°C + Tiron compared to its control at 22°C (for collected data see Table 1).
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Many single fibres did not survive the fatigue protocol at 37°C and it was not usually possible to perform three fatigue protocols as with the muscle bundles. However it was noticeable that the presence of Tiron greatly improved the survival of single fibres at 37°C suggesting that ROS are part of the cause of the failure of survival.
Maximum Ca2+-activated force
Fmax was determined before and immediately after each fatigue protocol by stimulation during a brief application of 10 mM caffeine. At 22°C it can be seen from Fig. 3 that Fmax was reduced by fatigue to about 80% of the prefatigue value. In five experiments Fmax,FAT was reduced to 89 ± 3% of the control value. This was not significantly different from the value of Fmax,FAT at 37°C which was 91 ± 3% of the prefatigue value (see Table 1). It was concluded that decline in Fmax during fatigue was similar in muscles at 37°C and at 22°C and was therefore not the cause of the more rapid fatiguability observed at 37°C.
Figure 4 shows that the change in Fmax was similar in experiments in which Tiron was present to reduce the increased rate of fatigue at 37°C. Fmax,FAT was 89 ± 2% of the control value at 22°C and it was not significantly changed at 94 ± 2% at 37°C in the presence of Tiron.
Myofibrillar Ca2+ sensitivity
Figure 5 illustrates the method of determining Ca50 from fits of Hill curves to prefatigue data at various stimulation frequencies (filled circles). For each tetanus, the force and tetanic [Ca2+]i were measured and plotted on the graph, along with a value of resting [Ca2+]i and the force and [Ca2+]i in the 100 Hz tetanus in the presence of caffeine (filled square). It can be seen that at 22°C (Fig. 5A) the Ca50,CON is about 650 nM. Table 2 collates the mean ± S.E.M. for values of Fmax, Ca50 and Hill coefficients for unfatigued muscles at 22°C, 37°C and 37°C + Tiron. Note that Fmax was significantly reduced to 83 ± 4% at 37°C but this decline was prevented by Tiron. Also the Ca2+ sensitivity increased significantly at 37°C compared to 22°C and this change was not influenced by Tiron. Hill coefficients were not significantly affected by the interventions.
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Ca50) during fatigue was 85 ± 22 nM (see Table 1). This is similar to previous estimates of the reduced myofibrillar Ca2+ sensitivity in fatigue at 22°C (Chin & Allen, 1997). In the same muscle at 37°C the results are strikingly different. Because force declines with no change in tetanic [Ca2+]i and the change in Fmax is similar to that at 22°C, there is a large change in Ca50,FAT (note the large rightward shift in data points for fatigue in Fig. 5B). In this experiment the prefatigue Ca50,CON is about 400 nM while the Ca50,FAT is about 700 nM. In seven experiments the mean Ca50 was 442 ± 146 nM. The sensitivity shift at 22°C and 37°C cannot be rigorously compared on a paired t test because the variance is different in the two groups but a nonparametric test (Mann-Witney rank sum) confirms them to be significantly different (P < 0.02). This substantial loss of myofibrillar Ca2+ sensitivity is clearer when individual tetani at the beginning of the fatigue protocol at 37°C are compared with the end of the fatigue protocol as in Fig. 6. Note that the force declines by a factor of 2 while the tetanic [Ca2+]i is largely unchanged. This figure also shows that there are no important changes in time course of force or [Ca2+]i under these conditions so that the Ca2+ sensitivity can be realistically estimated by this approach.
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Ca50 was 40 ± 6 nM, a value that was not significantly different from the 71 ± 21 nM determined at 22°C in the same fibres (see Table 1). We conclude that at 37°C the accelerated fatigue is largely attributable to a rapid decline in myofibrillar Ca2+ sensitivity and this effect is eliminated by the ROS scavenger Tiron.
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Discussion |
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Isolated skeletal muscles are often studied at subphysiological temperatures because preparations frequently degenerate rapidly at 37°C. In the present study we show that both in isolated muscle bundles and in single fibres, fatigue develops more rapidly at 37°C compared to room temperature and that the acceleration is prevented by the membrane-permeant antioxidant Tiron. Critically Tiron had no significant effect on the development of fatigue at room temperature confirming that other mechanisms dominate under these conditions.
A feature of our results was that fatigue was generally slower in muscle bundles than in single fibres (T1/1 4–8 min vs. 1–2 min) and the deleterious effects of high temperature took longer to develop. Muscles contain motor units and fibres with extremely variable fatigue times (Burke et al. 1973) so it is not surprising that muscle bundles and single fibres on dissection are also very variable. The main reason for the faster fatigue in single fibres is that during dissection there is a preference for large, clear fibres, which are fast-type fibres and therefore fatigue more quickly than the bundles, which are more likely to contain a representative range of fibre types. The greater sensitivity of single fibres to the effects of 37°C may be because the high concentration of O2 used in these studies accelerated the production of ROS. In the single fibres the PO2 would have been 
680 mmHg whereas in the muscle bundles the consumption of O2 by the outer fibres means that the inner fibres would have been subjected to a lower PO2 so that ROS levels may have been lower. Although this seems feasible studies on the intracellular levels of ROS measured with dichlorofluorescein found no changes in ROS over the range of O2 from 25 to 95% (Murrant et al. 1999).
The mechanism of fatigue at 37°C
Although we did not measure ROS in the present study, it is well established that muscles produce superoxide principally from mitochondria but also from xanthine oxidase and NADPH oxidase. Superoxide is broken down by superoxide dismutase and the resulting H2O2 can generate the very reactive hydroxyl radical in the presence of Fe3+. H2O2 is further degraded to H2O and O2 by catalase (for review see Supinski, 1998; Reid, 2001). Of particular relevance to the present study are the demonstrations that ROS production is elevated by higher temperatures (Arbogast & Reid, 2004), that contractile activity accelerates ROS production (Kolbeck et al. 1997) and that extracellular Tiron reduces the intracellular concentration of ROS (Silveira et al. 2003). Tiron is thought to act by scavenging superoxide and hydroxyl radicals; it is also a metal chelator and part of its activity may be by reducing intracellular concentrations of Fe3+ (Krishna et al. 1992). Tempol also scavenges superoxide and may act as a superoxide dismutase mimetic (Krishna et al. 1996). The similarity in the action of these two disparate ROS scavengers supports the idea that ROS have a role in fatigue at 37°C.
Important findings in the present study were that in the accelerated fatigue observed at 37°C there were no significant changes in tetanic [Ca2+]i during the fatigue period and the reduction in maximum Ca2+-activated force was similar to that observed at 22°C. ROS have been shown to increase the opening frequency of isolated SR Ca2+ channels (ryanodine receptors) in artificial lipid bilayers (Favero et al. 1995), but there is dispute as to whether H2O2 influences SR Ca2+ release in more intact preparations. Brotto & Nosek (1996), using a skinned skeletal muscle with intact T-tubule–SR coupling, found that H2O2 reduced depolarization-induced Ca2+ release whereas Posterino et al. (2003) found no such effect. Our results suggest that in the relatively brief exposure to elevated ROS at 37°C which our single fibres would have experienced (5–10 min), there was no apparent effect of ROS on SR Ca2+ release. This result fits well with earlier demonstrations that application of exogenous H2O2 to intact fibres had no effect on SR Ca2+ release (Andrade et al. 1998, 2001). In conclusion we found no obvious evidence that ROS affected tetanic or resting [Ca2+]i. We suspect that the normal sequence of changes in tetanic [Ca2+]i occur at 37°C, as observed in the presence of Tiron, but in the accelerated fatigue that occurs in the absence of Tiron the metabolic changes which cause the modifications to Ca2+ handling have not had time to occur.
Maximum Ca2+-activated force was assessed by application of 10 mM caffeine to a 100 Hz tetanus, which increases tetanic [Ca2+]i by a factor of 2 (Allen & Westerblad, 1995). One concern with this technique is that the elevation of tetanic [Ca2+]i at the end of fatiguing stimulation may not be sufficient to saturate the troponin sites so that the measured force will be less than maximum. For the purposes of the present study the important point here is that errors from this source would have caused an underestimate of the maximum Ca2+-activated force and therefore reinforce our conclusion that there was no substantial reduction in maximum force contributing to the accelerated fatigue at 37°C. This conclusion is perhaps surprising given the findings of van der Poel & Stephenson (2002) in which heating rat skeletal muscle to 43–47°C for short periods produced a reversible loss of force production. The damaged fibres were then skinned and shown to have a reduction in the maximum Ca2+-activated force but no change in Ca2+ sensitivity. As in our study Tiron was able to prevent these effects. Two experimental differences, which might explain the different cellular mechanisms, are the higher temperature in their study and the fact that they did not stimulate the muscle at the elevated temperature.
Reduced myofibrillar Ca2+ sensitivity
Our main finding is that the accelerated fatigue could be explained by a reduction of myofibrillar Ca2+ sensitivity. The reduction of sensitivity appeared to develop sufficiently rapidly so that the fall in force occurred in the absence of changes in tetanic [Ca2+]i and the reduction in maximum force was not different from that observed at 22°C. This result is consistent with the observation that the predominant effect of increasing intracellular ROS by application of extracellular H2O2 to intact single fibres is a reduction in myofibrillar Ca2+ sensitivity (Andrade et al. 1998). In the study by Andrade et al. (1998) this reduction could be reversed by dithiothreitol, which suggested that ROS oxidized free sulphydryl groups to –S–S– linkages. However in a later study Andrade et al. (2001) found that much lower concentrations of H2O2 caused an early increase in Ca2+ sensitivity which subsequently declined. Thus, like many effects of ROS, the effects of H2O2 seem to depend on concentration and duration of exposure.
There are a number of studies in which various free radicals (superoxide, H2O2, hydroxyl) have been applied directly to myofibrils in skinned skeletal preparations. Callahan et al. (2001) and Darnley et al. (2001) both studied skinned respiratory muscle and both found that application of various ROS reduced maximum force but had no effect on Ca2+ sensitivity. In contrast Lamb & Posterino (2003) found that ROS could increase Ca2+ sensitivity particularly when glutathione was present. Thus results from skinned fibres are somewhat varied but none exhibit the fall in Ca2+ sensitivity observed with substantial concentrations of H2O2 (Andrade et al. 1998) or the fall in Ca2+ sensitivity that we observed in muscles fatigued at 37°C. Taken together these results suggest that ROS do not act directly on the myofilaments but act indirectly on a ROS-sensitive pathway which results in loss of myofibrillar Ca2+ sensitivity.
Another situation in which loss of Ca2+ sensitivity has been observed is the postischaemic, reperfused (stunned) myocardium (Gao et al. 1996). These authors showed that activation of calpain by elevated [Ca2+]i resulted in troponin I degradation causing the reduction of Ca2+ sensitivity. Furthermore this loss of Ca2+ sensitivity could be partially prevented if ROS scavengers were present during the period of ischaemia and reperfusion implicating ROS in the development of the loss of Ca2+ sensitivity (Perez et al. 1998). Our findings could be explained if increased levels of ROS were to activate calpain (directly or indirectly) which subsequently caused troponin I degradation. However measurements of both resting and tetanic [Ca2+]i did not show any additional rise in [Ca2+]i at 37°C; nor was the [Ca2+]i higher in the absence of Tiron. Thus the present results provide no clear support for the idea that elevated [Ca2+]i associated with ROS production causes increased calpain activity and contributes to the loss of Ca2+ sensitivity.
Relevance of results to muscular function in intact animals
Our study supports an extensive literature showing that ROS contribute to fatigue in skeletal muscle (for review see Supinski, 1998; Reid, 2001). A feature of our study is that it shows that the contribution of ROS at 37°C is much greater than at 22°C and suggests that future studies of fatigue on isolated muscles may need to be at 37°C if this particular mechanism is to be assessed and studied. It is generally accepted that the consequences of ROS depend on a balance between ROS production and endogenous scavenging pathways and it is very likely that in an isolated single muscle fibre this balance is disturbed. For this reason studies on isolated preparations, while useful for identifying mechanisms, probably do not give a quantitative indication of the magnitude of this mechanism in intact animals. One approach to identifying the role of ROS is the use of ROS scavengers in humans and such studies show that the antioxidant N-acetylcysteine was capable of slowing the onset of some types of fatigue (Reid et al. 1994). The main value of our study is that it suggests that ROS accelerate fatigue principally by reducing the Ca2+ sensitivity of the myofibrils. Further studies will be need to identify the target proteins of ROS and the pathway by which ROS affect myofibrillar Ca2+ sensitivity.
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0.02 on a paired t test. Number of experiments indicated by number inside each column.
