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Effects of progressive exercise and hypoxia on human muscle sarcoplasmic reticulum function

Department of Kinesiology, University of Waterloo, Waterloo, Ontario, Canada N2L 3G1

Submitted 5 September 2003 ; accepted in final form 9 January 2004

	ABSTRACT

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This study examined the effects of progressive exercise to fatigue in normoxia (N) on muscle sarcoplasmic reticulum (SR) Ca²⁺ cycling and whether alterations in SR Ca²⁺ cycling are related to the blunted peak mechanical power output (PO_peak) and peak oxygen consumption (O_{2 peak}) observed during progressive exercise in hypoxia (H). Nine untrained men (20.7 ± 0.42 yr) performed progressive cycle exercise to fatigue on two occasions, namely during N (inspired oxygen fraction = 0.21) and during H (inspired oxygen fraction = 0.14). Tissue extracted from the vastus lateralis before exercise and at power output corresponding to 50 and 70% of O_{2 peak} (as determined during N) and at fatigue was used to investigate changes in homogenate SR Ca²⁺-cycling properties. Exercise in H compared with N resulted in a 19 and 21% lower (P < 0.05) PO_peak and O_{2 peak}, respectively. During progressive exercise in N, Ca²⁺-ATPase kinetics, as determined by maximal activity, the Hill coefficient, and the Ca²⁺ concentration at one-half maximal activity were not altered. However, reductions with exercise in N were noted in Ca²⁺ uptake (before exercise = 357 ± 29 µmol·min^–1·g protein^–1; at fatigue = 306 ± 26 µmol·min^–1·g protein^–1; P < 0.05) when measured at free Ca²⁺ concentration of 2 µM and in phase 2 Ca²⁺ release (before exercise = 716 ± 33 µmol·min^–1·g protein^–1; at fatigue = 500 ± 53 µmol·min^–1·g protein^–1; P < 0.05) when measured in vitro in whole muscle homogenates. No differences were noted between N and H conditions at comparable power output or at fatigue. It is concluded that, although structural changes in SR Ca²⁺-cycling proteins may explain fatigue during progressive exercise in N, they cannot explain the lower PO_peak and O_{2 peak} observed during H.

calcium cycling; muscle; normoxia; fatigue

The SR, an intracellular membranous network that envelopes the myofibrils, regulates [Ca²⁺]_f levels by selective control of the rate of Ca²⁺ release from Ca²⁺ stored inside the SR and by the rate of Ca²⁺ uptake (4). Ca²⁺ release depends on the open state of the Ca²⁺-release channels (CRC) or ryanodine receptors, an 450-kDa protein embedded in the SR membrane, most of which is localized to the terminal cisternae, which is in close apposition to the T tubules (12). It is now generally acknowledged that, in skeletal muscle, the action potentials that are conducted along the sarcolemma and into the T tubule result in a physical-chemical coupling between the T tubules and the CRC and opening of the CRC, enabling the rapid release of Ca²⁺ (4). The sequestration of [Ca²⁺]_f back into the SR is controlled by the Ca²⁺-ATPase, an 110-kDa protein located primarily in the longitudinal SR, which uses the energy from the hydrolysis of ATP to pump Ca²⁺ against its concentration gradient for storage in the SR (27). Increases in [Ca²⁺]_f are a potent stimulus for increasing the activity of Ca²⁺-ATPase and, consequently, the rate of Ca²⁺ uptake into the SR (32).

The possibility that disturbances in SR Ca²⁺ cycling may be involved in fatigue during submaximal exercise is also supported by several studies that have demonstrated reductions in both Ca²⁺-ATPase activity (6, 7, 14), Ca²⁺ uptake (6, 7), and Ca²⁺ release (10) in both rat and human locomotor muscles. Information is emerging to indicate that the disturbances in Ca²⁺ uptake and Ca²⁺ release are mediated by structural adaptations in the Ca²⁺-ATPase enzyme (26) and the CRC (9) secondary to oxidation and/or nitrosylation mediated by reactive O₂ species (ROS) (23, 24).

During intense dynamic exercise, increases in the rate of Ca²⁺ cycling in the muscle cell and consequently in the rate of Ca²⁺ release and Ca²⁺ uptake are necessary to realize increases in PO. A disturbance in either Ca²⁺ release and/or Ca²⁺ uptake would be expected to alter the [Ca²⁺]_f integral and disrupt force profiles (1).

Intuitively, one might expect that supramaximal fatiguing exercise would produce even more dramatic reductions in SR Ca²⁺ cycling. There is evidence to support this idea since pronounced reductions in Ca²⁺-ATPase activity and Ca²⁺ uptake have been reported with exhaustive running in horses (8) and rats (52) and with intense knee extensions in humans (13, 25). Pronounced depressions in SR Ca²⁺ release have also been reported (25, 43). However, not all studies are consistent, because reports have been published indicating no change in Ca²⁺-ATPase activity (19, 31) and in Ca²⁺ uptake (18, 31) with intense, repetitive contractions.

Progressive dynamic exercise to fatigue is commonly employed to assess peak aerobic power [peak O₂ consumption (O_{2 peak})], one of the most important properties in applied physiology. However, it is unclear whether the peak mechanical power output (PO_peak) attained is limited by a failure in SR Ca²⁺ cycling.

Hypoxia is known to result in a reduction in PO_peak and O_{2 peak} during progressive exercise to fatigue (15, 33). It is generally believed that the reduction in O_{2 peak} observed in hypoxia occurs because of limitations in O₂ availability to the mitochondria, secondary to reductions in arterial O₂ content and O₂ delivery to working muscles (51). However, this interpretation could be simplistic because it assumes that the viability of the E-C processes in muscle are protected, allowing power output (PO) to increase and available O₂ to be fully utilized for ATP regeneration to contribute the increasing demands of the cellular ATPases involved in E-C. It is possible that the reverse may be true, namely that E-C failure may occur prematurely, resulting in fatigue and an inability to utilize available O₂ for oxidative phosphorylation (OxPhos).

The purpose of this study was to investigate the effects of progressive cycle exercise in normoxia and hypoxia on muscle SR function. We have hypothesized that progressive reductions in Ca²⁺-ATPase activity, Ca²⁺ uptake, and Ca²⁺ release would occur with increases in PO. We have also postulated that, at similar PO, the depression in the Ca²⁺-cycling properties would be more pronounced in hypoxia compared with normoxia. However, at fatigue in normoxia vs. hypoxia, the disturbances would be comparable, consistent with the lower PO_peak and O_{2 peak} typically observed with hypoxia.

	METHODS

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Participants.   Participants were nine healthy male university students who were not engaged in vigorous physical activity on a regular basis. Age, height, and body mass were 20.0 ± 0.37 yr, 175 ± 2.3 cm, and 80.0 ± 3.5 kg (means ± SE), respectively. All procedures were approved by the Office of Research Ethics, and all experimental protocols and risks were carefully explained to each volunteer before written consent was obtained.

Experimental design.   To investigate the effects of progressive cycle exercise during normoxia and hypoxia, a randomized cross-over design was employed. In this design, all participants completed each condition within a minimum of 3 wk between test sessions for each individual. At least 1 wk before the beginning of the experiment, volunteers reported to the laboratory and completed a preliminary incremental exercise test to fatigue during normoxia. This test was used to ensure a normal response to exercise and to verify that a maximal or near maximal physiological response could be achieved. For all tests, fatigue was defined as an inability to maintain pedal frequency at 60 revolutions/min, even with verbal encouragement.

During the actual test days, the participants reported to the laboratory 60 min before the beginning of the exercise test. During this time, the individual was prepared for muscle biopsies. For tissue sampling, 0.5 ml of a local anaesthetic (2% xylocaine) was administered to selected sites over the vastus lateralis muscles and then prepared for needle biopsies (5). For the test in normoxia, four separate sites were prepared balanced between the two legs. For the test in hypoxia, three separate sites were prepared, two in one leg and one in the other. After the preliminary preparations, the volunteers reported to the exercise testing area, mounted an electrically braked cycle ergometer (Siemens Elma 380 B), and were prepared for heart rate and respiratory gas-exchange measurements (21). These measurements were obtained continuously before and throughout exercise.

For the test conducted in normoxia, the participant breathed room air for 15 min before the beginning of exercise (Fig. 1). The exercise protocol consisted of 4 min of cycling at 25 W followed by step increases in work rate of 15 W/min until fatigue (Fig. 1). Before the start of cycling, resting tissue samples were obtained. In addition, tissue samples were obtained at mechanical POs equivalent to 50 and 70% O_{2 peak} as determined in normoxia and at fatigue. To obtain the exercise biopsies, the cycling was briefly interrupted and the tissue quickly extracted from previously prepared sites. The order of biopsies was alternated between legs.

View larger version (17K): [in this window] [in a new window]   Fig. 1. Characteristics of the progressive exercise protocol and tissue sampling schedule used in normoxia and hypoxia. The bold stairs indicate the step increases in power output (PO) from 25 to 300 W. Arrows indicate the time of tissue sampling; 50% O2 peak (N), PO at which 50% peak aerobic power (O2 peak) for each volunteer, determined in normoxia, occurred; 70% O2 peak (N), PO at which 70% O2 peak for each volunteer, determined in normoxia, occurred; B(N), biopsy for normoxic condition; B(H), biopsy for hypoxic condition; respiratory gas, period of respiratory gas collection. During exercise, respiratory gas collection was continuous throughout the exercise. After the first work load (25 W), which was 4 min, work loads were increased by 15 W each minute.

On a given experimental day, the participant was instructed to consume a light snack consisting of juices. Approximately 3 h before reporting to the laboratory, a meal-replacement beverage was provided (Ensure; 250 kcal; 61, 24, and 15% Kcals from carbohydrates, lipids, and proteins, respectively). This procedure was utilized to standardize the nutritional intake before the two test conditions. Before testing, water intake was not controlled. All volunteers were instructed not to consume alcohol and caffeine nor to engage in heavy exercise for at least 24 h before testing. For each subject, the two exercise tests were performed at approximately the same time of day. For respiratory gas collection, an open-circuit system was employed as previously described (21). The electronic cycle was calibrated on a daily basis, and care was taken to ensure that the seat height was standardized for each volunteer between tests. Average environmental conditions in the testing room were not different between conditions and ranged between 50 and 60% relative humidity and 21 and 22°C in temperature.

Measurement of SR function.   Immediately after extraction of the tissue, homogenates were prepared and stored at –80°C until analyses of Ca²⁺-ATPase activity, Ca²⁺ uptake, and Ca²⁺ release. Homogenates, prepared under ice-cold conditions, consisted of 11:1 (vol/wt) dilution of buffer containing (in mM) 250 sucrose, 5 HEPES, 10 NaN₃, and 0.2 PMSF (pH 7.5). Tissue (30–40 mg) was homogenized by using a handheld glass homogenizer (Kontes, Duall 20) (29). Once prepared, the homogenate was separated into a number of aliquots and quick frozen in liquid N₂ before storage. On a given analytical day and just before analysis, aliquots were thawed and processed for specific measurements. For a given property, duplicate measurements were performed. Care was taken to ensure that a complete set of measurements were made on all tissues for a given participant during a specific analytical session.

Ca²⁺-ATPase activity.   Ca²⁺-ATPase activity was assessed spectrophotmetrically using the basic procedures developed by Simonides and van Hardeveld (40), as modified in our laboratory (45). The reaction buffer contained (in mM) 200 KC1, 20 HEPES, 15 Mg Cl₂, 1 EGTA, 10 NaN₃, 5 ATP, and 10 phosphoenolpyruvate. The pH of the buffer was adjusted to 7.0 at 37°C. Just before the reaction was started, 18 U/ml lactate dehydrogenase, 18 U/ml pyruvate kinase, 0.3 mM NADH, and 25 µl of homogenate were added to a cuvette containing 1 ml of reaction buffer. One micromolar Ca²⁺ ionophone A-23187 (Sigma C-7522) was also added to prevent intraluminal Ca²⁺ accumulation in SR and inhibition of Ca²⁺-ATPase activity (40). Assays were performed in duplicate at 37°C and 340 nm (Shimadzu UV 160) using 1 mg wet wt of tissue per assay. After the baseline absorbance of NADH was recorded for 1 min, the reaction was initiated by adding 1 µl of 100 mM CaCl₂ and monitored for 2 min.

Maximal Ca²⁺-ATPase activity and Ca²⁺ dependency of Ca²⁺-ATPase activity were measured using a 0.5-µl addition of 100 mM CaCl₂. The additions were continued until a plateau and subsequent decline in Ca²⁺-ATPase activity were observed. Basal or Mg²⁺-ATPase activity was determined in the presence of 40 µM of the specific inhibitor of the Ca²⁺-ATPase cyclopiazonic acid (38). Ca²⁺-ATPase activity is based on the difference between the total and basal ATPase activities. The validity of SR Ca²⁺-ATPase activity assay has been previously established in whole muscle homogenates (34, 40). It should be emphasized that a potential limitation of the regenerating assay employed is the possible loss of ADP to AMP via the myokinase reaction. However, we have shown using a myokinase inhibitor (AP5A) that Ca²⁺-ATPase activity is unaffected (unpublished observations).

Measurement of [Ca²⁺]_f, which is used to assess Ca²⁺ dependency of the Ca²⁺-ATPase reaction, was assessed with dual-wave spectrofluorometry and the Ca²⁺ fluorescent dye indo 1 according to procedures previously detailed (46, 48). In summary, the measurement of [Ca²⁺]_f is based on the difference in maximal emission wavelengths between the Ca²⁺-bound-indo 1 complex and the Ca²⁺-free indo 1 complex. An excitation wavelength of 355 nm and the emission maxima were recorded at 405 and 485 nm for Ca²⁺-bound- and Ca²⁺-free indo 1. Felix software (Photon Technology International) was used to calculate the ionized Ca²⁺ concentration according of Eq 2.5 of Grynkiewicz et al. (17). The dissociation constant that was used for the interaction between Ca²⁺ and indo 1 was 250 nM (17). It should be emphasized that in homogenates the actual dissociation constant can be significantly affected by the protein composition. As such, given the uncertainty as to the true value of the dissociation constant, the pCa values used throughout the text are only apparent.

The kinetic properties of the Ca²⁺-ATPase activity that were measured included the maximal activity (V_max), the [Ca²⁺]_f needed to obtain half-V_max (Ca₅₀), and the Hill coefficient (n_H). To obtain these properties, Ca²⁺-ATPase activity was plotted against the negative logarithm of [Ca²⁺]_f (pCa). V_max represented the peak value, Ca₅₀ represented the [Ca²⁺]_f obtained from a sigmoid fit of the data that yields 50% of V_max, and n_H was obtained through nonlinear regression with computer software (GraphPad Software) by using a portion of the curve that corresponded to between 20 and 80% of V_max. These procedures have been described in recent papers from our laboratory (37, 49).

Ca²⁺ uptake and Ca²⁺ release.   Both Ca²⁺ uptake and Ca²⁺ release were measured during the same assay. In general, the procedure employed involved using the Ca²⁺ fluorescent dye indo 1 according to the methods of O'Brien et al. (30) as modified by our group (46). The reaction buffer for these assays contained (in mM) 200 KC1, 20 HEPES, 15 MgCl₂, 10 NaN₃, 0.005 N,N,N¹,N¹-tetrakis(2-pyridlymethyl)-ethylenediamine, 5 oxalate, and 10 phosphoenolpyruvate. Before each assay, 1.5 µM indo 1, 18 U/ml lactate dehydrogenase, and 18 U/ml pyruvate kinase were added to 2 ml of reaction buffer. In addition, 2.5 µl of CaCl₂ (10 mM) were added to the cuvette to produce a consistent starting [Ca²⁺]_f of 3.5 µM. After a constant [Ca²⁺]_f was achieved, 5 mM ATP was added to the cuvette to initiate Ca²⁺ uptake. Ca²⁺-uptake rates were assessed over a range of [Ca²⁺]_f by using a single assay. [Ca²⁺]_f was measured as described earlier. The maximal rate of Ca²⁺ uptake for each [Ca²⁺]_f was determined by differentiating the linear fit curve using four concentrations of cytosolic free Ca²⁺, namely 500, 1,000, 1,500, and 2,000 µM. We have found that with this procedure there is a rapid initial drop in [Ca²⁺]_f. A large component of the initial drop in [Ca²⁺]_f has been credited to the binding of Ca²⁺ with proteins in the homogenates (34). As a consequence, the initial changes in [Ca²⁺]_f were not used in the calculation of Ca²⁺ uptake (34).

Ca²⁺ release was measured immediately after Ca²⁺ uptake according to the general procedures of Ruell et al. (34). However, unlike Ruell et al., who employed AgNO₃, we have employed 4-chloro-m-cresol (4-CMC). We have recently shown that effects of 4-CMC are specific to the CRC and do not cause Ca²⁺-ATPase pump reversal, as was noted for AgNO₃ (47). To assess Ca²⁺-release rates, 20 µM of 4-CMC was added to the cuvette after [Ca²⁺]_f declined to a plateau after the measurement of Ca²⁺ uptake. We have found that a biphasic Ca²⁺ release can be distinguished, an early release, which we have labeled phase 1, and a slower, more delayed release, which we have labeled phase 2 (47). Maximal release rates for each phase were calculated using the same method as for Ca²⁺ uptake and differentiating using a linear-fit curve.

It should be noted that, because Ca²⁺ uptake and Ca²⁺ release were measured in the same assay, the reaction buffers were the same. It is now possible that optimal assay conditions may vary with the property measured. As an example, 15 mM MgCl₂ was used for both Ca²⁺ uptake and Ca²⁺ release. In the case of Ca²⁺ release, optimal MgCl₂ levels appear to be between 20 and 30 mM (unpublished observations).

For all SR properties, protein was determined by the method of Lowry as modified by Schacterle and Pollock (36). This involved separate protein assessments in homogenate used for Ca²⁺-ATPase activity and in homogenates used for Ca²⁺-uptake and Ca²⁺-release assays. There were no significant differences in protein levels between normoxic and hypoxic conditions. The coefficients of variation for duplicate measurements for Ca²⁺ uptake and Ca²⁺-ATPase activity were 7.6 ± 0.9 and 7.9 ± 2.2%, respectively. Only single measurements were used for Ca²⁺-release determinations.

Statistics.   Group data are expressed throughout the text as means ± SE. To examine the effects of progressive exercise and O₂ condition on SR function, linear regression techniques were employed, and the slope of the regression line for each condition was established. Student's t-tests were used to examine differences between slopes and, hence, conditions. Conventional two-way ANOVA procedures were not possible because of the unbalanced nature of the tissue-sampling schedule (see Fig. 1). Where only one variable was of interest (such as during exercise in normoxia or hypoxia), a one-way ANOVA with repeated measures was employed. Two-way ANOVA procedures were used to examine the effects of condition and exercise on the gas-exchange properties. We have also performed a secondary analysis, using two-way ANOVA procedures, to determine the effects of exercise and O₂ condition on SR properties using matched samples. Where significance was found, Newman-Keuls techniques were applied to locate differences between specific means. The significance level was set at P < 0.05 for all comparisons.

	RESULTS

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Peak mechanical and aerobic power.   PO_peak generated during progressive cycle exercise to fatigue in normoxia was 300 ± 8.7 W. During hypoxia, PO_peak was 250 ± 7.9 W, 19% lower than the value obtained during normoxia. As expected, O_{2 peak} was also reduced with hypoxia compared with normoxia (4.07 ± 0.15 vs. 3.21 ± 0.11 l/min, respectively).

Measurements of exchange during normoxia and hypoxia were obtained at matching POs of 137 ± 51, 214 ± 6.9, and 250 ± 7 W (Table 1). These data indicated that no differences existed in O₂ (l/min) between normoxia and hypoxia at corresponding POs. As expected, CO₂ production, expiratory minute volume (BTPS), and respiratory exchange ratio were all higher during exercise in hypoxia compared with normoxia. In the case of CO₂ production and expiratory minute volume, the differences were observed both at rest and during each PO.

View larger version (12K): [in this window] [in a new window]   Fig. 2. Representative curves for a single participant illustrating the Ca2+-dependent changes in Ca2+-ATPase activity in normoxia and hypoxia. These measurements were made at rest. The kinetic properties of the Ca2+-ATPase were obtained from this curve.

View larger version (22K): [in this window] [in a new window]   Fig. 3. Ca2+-dependent changes in Ca2+ uptake during progressive exercise in normoxia and hypoxia. Ca2+ uptake was assessed at 4 different systolic free Ca2+ concentrations ([Ca2+]f; 500, 1,000, 1,500, and 2,000 nM). O2, O2 consumption assessed during the progressive exercise tests. Main effects (P < 0.05) for [Ca2+]f and exercise were observed. For Ca2+, 2,000 > 1,500 > 1,000 > 500 nM. For exercise, before exercise was greater than fatigue. Values are means ± SE (n = 9).

View larger version (11K): [in this window] [in a new window]   Fig. 4. Coupling ratios obtained during progressive exercise in normoxia and hypoxia. Coupling ratio is defined as the ratio of Ca2+ uptake (2,000 nM) to Ca2+-ATPase activity. Values are means ± SE (n = 10) determined at rest and during exercise to fatigue.

View larger version (15K): [in this window] [in a new window]   Fig. 5. Representative trace for the measurement of Ca2+ uptake and Ca2+ release. Ca2+ uptake was initiated with the addition of ATP to the homogenate. After Ca2+ levels had plateaued (baseline), 4-chloro-m-cresol (4-CMC) was added to the cuvette to initiate Ca2+ release. Uptake, tracing for Ca2+ uptake by sarcoplasmic reticulum. Two phases of Ca2+ release were identified: a fast phase (phase 1) and a slow phase (phase 2).

View larger version (15K): [in this window] [in a new window]   Fig. 6. Changes in Ca2+ release during progressive exercise in normoxia and hypoxia during phase 1 (A) and phase 2 (B). Ca2+ release has been plotted against O2 before exercise and during progressive exercise. Reductions (P < 0.05) in phase 2 of Ca2+ release were observed during exercise for each condition. *Significantly different from before exercise (P < 0.05). Significantly different from 70% O2 peak in normoxia (P < 0.05). Significantly different from 50% O2 peak in normoxia (P < 0.05).

	DISCUSSION

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Although reductions in Ca²⁺ cycling, Ca²⁺ uptake, and Ca²⁺ release with various exercise protocols have previously been shown to occur in humans (6, 25, 45), our study appears to be the first to demonstrate disturbances in SR function during progressive exercise. The reduction in Ca²⁺ uptake that we have observed in vitro appears to occur in the absence of changes in the kinetic properties of the Ca²⁺-ATPase because we could find no effect of exercise on V_max, n_H, or Ca₅₀ either during normoxia or hypoxia. This was unexpected because previous studies in both animals (7) and humans (6) have reported parallel reductions in V_max and Ca²⁺ uptake with prolonged exercise. However, there are reports where changes in Ca²⁺ uptake are dissociated from changes in Ca²⁺-ATPase activity (3, 11). Our results are similar to a previous study (19) in humans investigating the effects of intense dynamic knee extensor exercise in which reductions in Ca²⁺ uptake occurred in the absence of changes in Ca²⁺-ATPase activity.

At present, it is not clear what the underlying mechanism is to explain the dissociation between these two properties that has been observed. Several possibilities exist. First, it should be emphasized that the assays for Ca²⁺ uptake and Ca²⁺-ATPase activity are performed under different conditions. In the case of Ca²⁺ uptake, oxalate is employed to bind Ca²⁺ in the lumen of the SR (28). Oxalate has been previously shown to increase Ca²⁺-uptake rates in homogenates, conceivably by preventing the inhibitory effects of increases in luminal [Ca²⁺]_f (22). For the measurement of Ca²⁺-ATPase activity, the Ca²⁺ ionophone A23187 [GenBank] was employed, which prevents the accumulation of Ca²⁺ in the lumen of the SR and causes back inhibition of Ca²⁺-ATPase activity by allowing diffusion of Ca²⁺ out of the SR (40). The reduction in Ca²⁺ uptake that we have observed in exercise in the absence of changes in Ca²⁺-ATPase activity could occur as a result of slippage of the Ca²⁺-ATPase during the catalytic cycle of Ca²⁺ transport. This has been observed to occur as a result of sarcolipin, a small peptide located in association with the Ca²⁺-ATPase (42). How a similar effect would be induced by progressive exercise is unclear. It should be emphasized that the reductions in Ca²⁺ uptake that we have observed at fatigue were relatively modest and insufficient to significantly alter the coupling ratio between Ca²⁺-ATPase activity and Ca²⁺ uptake. Because coupling ratios were not altered with exercise either during normoxia or hypoxia, it would appear that the energy costs per unit of Ca²⁺ transport into the SR were unchanged (4, 27).

The most pronounced effect of progressive exercise on the SR function was noted for Ca²⁺ release. Reductions in Ca²⁺ release assessed in vitro have been commonly observed with a variety of exercise protocols (6, 9, 19, 25, 31). Reductions in Ca²⁺ release appear to be associated with a reduction in [³H]ryanodine binding (9), which has been used as evidence to support the hypothesis that structural damage to the CRC occurs (9). [³H]ryanodine is known to bind specifically to a 245-amino acid-long polypetide, which is located in a region encompassing most of the myoplasmic loop-2, which is closely associated with the transmembrane segments M₅ of the CRC (12). This has been identified as the putative functional domain of the CRC, and as a consequence [³H]ryanodine binding reflects the open state of the channels (10).

In this study, we have used 4-CMC as the CRC-releasing agent. The selection of 4-CMC, instead of AgNO₃, which has previously been commonly employed as a releasing agent (9), was based on a recent publication by our group (47). In that study, our laboratory found that AgNO₃, in contrast to 4-CMC, induces Ca²⁺ release by acting on both the CRC and Ca²⁺-ATPase. We have also been able to identify two phases of Ca²⁺ release, phase 1 and phase 2. We have observed that only reductions in phase 2 were observed with exercise. At present, the physiological significance of this observation is not clear because, in our in vitro system, the Ca²⁺-release rates occur at a much slower rate than observed in single-fiber and isolated-channel preparations (9). Our homogenate system is based on a preparation from whole muscle and consequently contains all of the proteins regulating SR behavior. This factor would appear important in the regulation of Ca²⁺ release and explain, at least in part, differences that exist between the homogenate system and other preparations, such as isolated vesicles and single channels. It must be acknowledged, however, that, regardless of the preparation, some functional alterations could be expected during the homogenization process.

The reduction in Ca²⁺ release that we have observed with our progressive exercise protocol probably results from a decrease in the number of functional CRC (9). Although definitive evidence is lacking, it is strongly suspected that oxidative and/or nitrosylative processes mediated by ROS are involved in the structural modifications to the CRC that occur (9, 44). It would be expected that ROS would accumulate during progressive exercise given the increase in OxPhos and the increase in inosine 5'-monophosphate (IMP) that occurs (16). It has been estimated that 2–4% of OxPhos results in ROS generation by the election transport system (39, 41). In addition, the conversion of IMP to hypoxanthine and xanthine by xanthine oxidase also increases ROS formation (39, 41). Conceivably, the increase in ROS generation by xanthine oxidase during exercise would occur given the larger IMP accumulation (35). It is possible that the increase in ROS was insufficient to result in a greater reduction in Ca²⁺ release with hypoxia compared with normoxia. At present, it is not clear why reductions in Ca²⁺-ATPase activities during progressive exercise did not occur in either normoxia or hypoxia, given the apparent sensitivity of this enzyme to ROS (23).

A potential factor that could influence our measurements of the SR properties has to do with potential oxidation during the homogenate preparation. In our study, as well as most similar type studies, the DTT, a sulfhydryl-reducing agent, was not added. Consequently, it would be expected that some oxidation would occur, possibly masking the actual changes that occurred with exercise. DTT was not added because of the possibility that the effects of exercise, if due to oxidation, would be reversed.

In this study, we have confirmed what has been reported on many previous occasions, namely that, with hypoxia (inspired O₂ fraction = 0.14), O_{2 peak} is reduced compared with normoxia (15, 33). The 23% reduction that was observed in O_{2 peak} is consistent with what has been reported earlier in untrained subjects under similar conditions (20). We have also been able to demonstrate, again similar to previous studies (20), that, over a range of submaximal POs, compensatory adjustments allow OxPhos to remain similar or nearly similar to normoxia despite a lower arterial O₂ saturation and arterial O₂ content. Even at the highest PO, attained at fatigue in hypoxia, we could find no differences from normoxia when matched to the same PO. Some insight into these adjustments can be obtained from the minute ventilation and respiratory exchange ratio measurements, both of which were increased in hypoxia compared with normoxia at similar POs. Increased ventilation in conjunction with increased blood flow to working muscles appears important in minimizing the reductions in arterial O₂ delivery in hypoxia (50, 51). Our results indicate that a substantial blunting of PO and O_{2 peak} can occur even though OxPhos is relatively well protected throughout submaximal exercise. This observation could suggest that fatigue may occur somewhat independently of a failure in ATP supply by OxPhos. Conceivably, fatigue could also prevent further increases in OxPhos (and O_{2 peak}) even though delivery, extraction, and utilization of O₂ by mitochondria have not been fully exploited. Failure in E-C coupling as a cause of fatigue remains an inviting possibility in this regard.

To examine differences between normoxic and hypoxic conditions in Ca²⁺-cycling properties, it was not possible to use conventional two-way ANOVA procedures for repeated measures as a result of the unbalanced nature of our tissue-sampling schedule. Although the first exercise samples occurred at the same O₂ (and PO), this was not possible at the higher exercise intensities due to the randomization of conditions and ethical limitations in the number of biopsies that could be performed. To examine for differences between conditions, we have used regression analyses and slopes from the lines of best fit for the relationship between O₂ (or PO) and the property of interest. These plots clearly demonstrate a trend (P > 0.05) toward a greater effect of hypoxia on Ca²⁺ uptake and Ca²⁺ release. It should be emphasized that this approach using regression analyses depends on the assumption of linearity between the measured parameters and PO.

Although not measured in this study, reductions in the [Ca²⁺]_f time integral remain a viable possibility to explain the inability to continue the progressive exercise task in both normoxia and hypoxia. Decreases in [Ca²⁺]_f time integral could occur specifically as a consequence of decreases in Ca²⁺ release. Reductions in the Ca²⁺ time integral, mediated by decreases in Ca²⁺ release, have been previously shown to associate with fatigue in repetitively stimulated mouse muscle fibers (2). The fact that the depression in Ca²⁺ release was comparable to normoxia in hypoxia at fatigue, even though PO was considerably lower in hypoxia, is also suggestive that impaired Ca²⁺ cycling may be involved in the early fatigue observed during progressive exercise in this condition.

It must be emphasized that other, more central processes may be involved in the inability to continue progressive exercise. Because our measurements were conducted on homogenates under supposedly optimal in vitro conditions, the depression in Ca²⁺ release can be attributed directly to disturbances in the CRC. However, in vivo, other processes such as an inability of the sarcolemma and T tubule to conduct a repetitive action potential and/or a deficit in the mechanical signaling between the T tubule and the CRC could lead to reductions in Ca²⁺release (2). It should also be emphasized that with progressive exercise profound changes occur in the intracellular environment, which may impair Ca²⁺ cycling as well as a variety of other excitation and contraction processes (53). Consequently, isolating the direct site and process involved in fatigue during voluntary activity is not possible.

In summary, our results demonstrate that, during progressive exercise to fatigue in normoxia, disturbances in SR Ca²⁺ cycling occur as indicated by the in vitro measurements of Ca²⁺ uptake and Ca²⁺ release. With hypoxia, similar disturbances in Ca²⁺ cycling occur but at a reduced PO_peak and O_{2 peak}. These findings raise the possibility that fatigue, both in normoxia and hypoxia with progressive exercise, is related to an inability to regulate the [Ca²⁺]_f integral.

	GRANTS

TOP ABSTRACT METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
Financial assistance for the study was provided by grants to H. Green from the National Sciences and Engineering Research Council of Canada.

	FOOTNOTES

 

Address for reprint requests and other correspondence: H. J. Green, Dept. of Kinesiology, Univ. of Waterloo, Waterloo, Ontario, Canada N2L 3G1 (E-mail: green{at}healthy.uwaterloo.ca).

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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