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J Appl Physiol 85: 619-626, 1998;
8750-7587/98 $5.00
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Vol. 85, Issue 2, 619-626, August 1998

Functional aspects of skeletal muscle contractile apparatus and sarcoplasmic reticulum after fatigue

Jay H. Williams, Christopher W. Ward, Espen E. Spangenburg, and Reagan M. Nelson

Muscular Function Laboratory, Department of Human Nutrition, Foods, and Exercise, Virginia Polytechnic Institute and State University, Blacksburg, Virginia 24061-0430

    ABSTRACT
Top
Abstract
Introduction
Methods
Results
Discussion
References

This study examined the effects of fatigue on the functional aspects of the contractile apparatus and sarcoplasmic reticulum (SR). Frog semitendinosus muscles were stimulated to fatigue, and skinned fibers or a homogenate fraction was prepared from both fatigued and rested contralateral muscles. In fatigued fibers, maximal Ca2+-activated force of the contractile apparatus was unaltered, whereas maximal actomyosin-ATPase activity was depressed by 20%. The Ca2+ sensitivity of force was increased, whereas that of actomyosin-ATPase was not altered. Also, the rate constant for tension redevelopment was decreased at submaximal Ca2+ concentration. These latter findings suggest that fatigue slows the dissociation of force-generating myosin cross bridges. Ca2+ uptake and Ca2+-ATPase activity of the SR were depressed by 46 and 21%, respectively, in the fatigued muscles. Fatigue also reduced the rates of SR Ca2+ release evoked by AgNO3 and 4-chloro-m-cresol by 38 and 45%, respectively. During fatigue, the contractile apparatus and SR undergo intrinsic functional alterations. These changes likely result in altered force production and energy consumption by the intact muscle.

calcium; adenosinetriphosphatase activity; muscle energetics; fatigue; skinned fibers; cross-bridge cycling kinetics

    INTRODUCTION
Top
Abstract
Introduction
Methods
Results
Discussion
References

IN RECENT YEARS, the notion that changes in the functional aspects of the contractile apparatus and/or sarcoplasmic reticulum (SR) contribute to the fatigue process has gained increasing support. In intact fibers, the decline in force production during fatiguing stimulation parallels the reduction in the tetanic intracellular Ca2+ concentration ([Ca]i), with the latter thought to occur secondary to depressed rates of SR Ca2+ uptake and release (1, 2, 27). In addition, the development of fatigue is associated with alterations in the force-generating properties of the contractile apparatus (28, 31). A portion of these changes can be attributed to the accumulation of metabolic by-products. Compounds such as H+, ADP, and Pi adversely affect the functional abilities of both the SR and the contractile apparatus (13, 16, 30). However, in many cases, temporal changes in metabolite levels during fatiguing activity do not always parallel changes in force production (20, 25). This has led to the notion that metabolite accumulation may not be the principle cause of fatigue and that other factors may be responsible.

Fatigue is also associated with intrinsic alterations in SR and contractile-apparatus function. That is, fatigue induces changes in function that persist after these structures are removed from the "fatigued" intracellular environment and examined under conditions that mimic those of a rested cell. For example, in SR vesicles isolated from fatigued muscles, the rates of Ca2+ uptake and release and Ca2+-ATPase activity are reduced by as much as 60% (for review, see Ref. 29). These changes occur after both voluntary activity leading to exhaustion as well as electrical stimulation of isolated muscle. It is important to point out that these alterations could not result from the direct action of metabolite accumulation because they are observed in incubation media that are free of elevated H+, ADP, and Pi.

We recently showed that, when frog semitendinosus muscles are stimulated to fatigue, skinned fibers display increased sensitivity to Ca2+ and decreased sensitivity to caffeine (28, 31). In these reports, we proposed that the enhanced Ca2+ sensitivity of the contractile apparatus results from altered cross-bridge cycling kinetics. Also, depressions in caffeine-induced force were thought to reflect depressions in the rates of SR Ca2+ uptake and release. Unfortunately, we were unable to make direct measurements of contractile-apparatus cross-bridge cycling kinetics, nor were we able to directly assess SR Ca2+ handling.

In the present investigation, we extended our previous studies of contractile apparatus and SR function in fatigued frog semitendinosus muscle. Our goal was to more clearly understand the nature of the fatigue-induced changes and to gain insight into the factors underlying these changes. First, we coupled mechanical and energetic measurements to clarify the mechanisms responsible for the increase in contractile apparatus Ca2+ sensitivity. Second, we examined changes in SR function by using a muscle homogenate fraction. This allowed us to directly determine the effects of fatigue on the rate of Ca2+ uptake and energy utilization of the Ca2+-ATPase as well as the rates of Ca2+ release evoked by varied releasing agents.

    METHODS
Top
Abstract
Introduction
Methods
Results
Discussion
References

Whole-muscle preparation. Whole semitendinosus muscles were obtained from small, male grass frogs (Rana pipiens) and placed in a normal Ringer solution that contained the following (in mM): 115 NaCl, 2.5 KCl, 1.8 CaCl2, 0.85 NaH2PO4, and 2.15 Na2HPO4 as well as 0.1 mg/ml d-tubocurarine. The normal Ringer solution was continually aerated with room air, and the pH was adjusted to 7.0. Muscles were mounted vertically between a fixed post and an isometric force transducer (Grass FT-03, 7P122 low-level direct-current amplifier) in a temperature-controlled muscle chamber (20°C). Contractions were electrically evoked by supramaximal square-wave pulses (0.2 ms; Grass S-88 stimulator and SIU-5 stimulus isolation unit) delivered across platinum wire electrodes that were situated at either end of the muscle. Tetanic contractions were evoked by trains of 0.2-ms pulses delivered at 80 Hz for 100 ms. Preliminary data indicate that this protocol elicits maximal force output of frog semitendinosus muscle and that stimulation frequencies >80 Hz cause little increase in tetanic force (Po). Before the fatigue protocol, muscles were equilibrated for 20 min, during which time optimal length was set (i.e., that which resulted in the greatest Po). Low-frequency fatigue was induced by tetanic contractions elicited at 2-Hz intervals for 5 min. Rested muscles were similarly mounted in the muscle chamber but were not subjected to the fatigue protocol. All contractions were displayed in an oscilloscope (Techtronix 2201) and then were digitized (1 kHz, 12-bit analog to digital, Keithley-MetraByte DAS-16) and stored on disk via microcomputer (IBM-PC 386-33 MHz). Each contraction was analyzed for Po.

Skinned-fiber experiments. Two stock solutions, which differed in free Ca2+ concentration ([Ca2+]), contained the following (in mM): 85.0 K+, 85.0 Na+, 1.0 Mg2+, 7.0 EGTA, 5.0 MgATP, and 10 phosphocreatine (PCr). The standard relaxing solution contained no added Ca2+ [-log free [Ca2+] (pCa) 9.0], and the activating solution contained adequate Ca2+ to achieve pCa 4.0. For measurements of actomyosin-ATPase activity (AM-ATPase), PCr was omitted and the following were added (in mM) 0.4 NADH, 5 phosphoenolpyruvate (PEP), 0.2 P1,P5-di(adenosine-5')pentaphosphate) (used to inhibit myokinase activity), 100 U/ml pyruvate kinase (PK), and 140 U/ml lactate dehydrogenase (LDH). Ionic strength of all solutions was adjusted to 0.18 M, and pH was maintained at 7.0 with imidazole. Propionate served as the major anion. The concentrations of the ionic species were determined by solving ionic equilibrium equations by using published binding constants (11) and a computer program kindly provided by Dr. W. Glenn L. Kerrick (University of Miami, Miami, FL).

In both rested and fatigued muscles, fiber bundles were rapidly dissected from the belly of the muscle and placed in a skinning solution for 20 min (relaxing solution containing 1% Triton X-100). With this skinning protocol, force could be elicited by Ca2+ but not by caffeine, suggesting that both the sarcolemma and the SR were disrupted. Also, the use of cyclopiazonic acid did not alter AM-ATPase activity, suggesting that the measured activity was not influenced by the Ca2+-ATPase of the SR. After being skinned, single fibers were mounted between a pair of microtweezers in a Muscle Research System (Scientific Instruments) (19). One pair of tweezers was attached to a photodiode force transducer and the other to a motorized length controller. The fiber was then placed in a quartz cuvette with a cross-sectional area of 1 mm2 and length of 1 cm. Resting length was set by stretching fiber to ~110% of slack length, resulting in a sarcomere length of 2.5 µm (as determined by HeNe laser diffraction). With the use of slack tests, the compliance of this system was found to be 3-4% of fiber length. All skinned fiber experiments were performed at 20°C.

Force and AM-ATPase activity responses to incremental increases in free Ca2+ were determined by stepwise changes in free [Ca2+], which were created with a calibrated gradient device (described below). Solutions were perfused through the cuvette in 28-µl increments, resulting in 60-80 increments of Ca2+ between the range of pCa 9.0-4.0. The solution in the cuvette was exchanged every 15 s. Preliminary experiments show that maximal force and AM-ATPase activity are maintained with repeated exposures to pCa 4.0 and with repeated gradient procedures. In addition, forces and AM-ATPase activities recorded during the gradient procedure were identical to those developed during random exposure to various free [Ca2+] levels. After each individual-fiber experiment, fiber diameter was determined via a video-imaging system (0.28-µm resolution). The smallest and largest diameters were averaged, and cross-sectional area was computed by assuming a cylindrical fiber shape.

The gradient device consisted of an upper chamber and a lower mixing chamber. Before each individual fiber experiment, the cuvette, tubing, and lower mixing chamber were filled with relaxing solution (pCa 9.0) while the upper chamber was filled with activating solution (pCa 4.0). The lower mixing chamber was constantly stirred with a magnetic stirring device. Solutions were perfused through the cuvette via a peristaltic pump. With each pump step, an aliquot (28 µl) of solution was withdrawn from the mixing chamber and replaced with activating solution from the upper chamber, allowing gradual incremental increases in free [Ca2+] within the mixing chamber. Free [Ca2+] delivered to the cuvette with each pump step was computed via an algorithm provided by Dr. Konrad Güth (Scientific Instruments). To verify the values computed by the algorithm, free [Ca2+] was periodically measured by using the fluorescent indicator calcium green-2 (480-nm excitation, 515-nm emission).

AM-ATPase activity was determined simultaneously with force production by an NADH fluorescence-based, coupled-enzyme assay (19, 24). The cuvette was illuminated by a high-pressure xenon lamp with light filtered at 340 nm, and a microscope photometer, housing a 470-nm filter, was used for detection of NADH fluorescence. The decline in fluorescence was determined over the last 10 s of the 15-s incubation interval, during which time the decay in NADH fluorescence was linear. AM-ATPase activities were corrected for basal activity, which was typically <5% of maximal (Amax).

Isometric force and AM-ATPase activities during each step were collected via computer and analyzed offline. The position and the shape of the force and AM-ATPase-free Ca2+ relationships were determined by fitting data obtained from each fiber to the modified Hill equation by using a nonlinear curve-fitting routine (SigmaPlot, Jandel Scientific)
F/F<SUB>max</SUB> = [Ca<SUP>2+</SUP>]<SUP><IT>N</IT></SUP> ⋅ ([Ca<SUP>2+</SUP>]<SUP><IT>N</IT></SUP><SUB>50</SUB> + [Ca<SUP>2+</SUP>]<SUP><IT>N</IT></SUP>)<SUP>−1</SUP>
where N is the slope of the relationship, [Ca2+]50 represents the Ca2+ concentration required to evoke 50% of maximal isometric force (Fmax) or Amax, and F is force output. This nonlinear approach yielded r2 values above 0.98 in all cases. Because of the large number of data points comprising each curve, statistical comparisons were made by using Fmax, Amax, [Ca2+]50, and N values. Preliminary experiments showed that, in six fibers that underwent these procedures three times, coefficients of variation for the above variables ranged from 3.2 to 6.8%. Also, parameters determined by using the gradient procedure were similar to those obtained with random exposures to varied free [Ca2+] solutions.

The rate constant of tension redevelopment (ktr) after a period of unloaded shortening was determined as described by Brenner and Eisenberg (6). The fiber was activated by Ca2+ (pCa 6.0-4.0) until a steady F was reached. It was rapidly shortened by 2% of its initial length and allowed to contract at maximal velocity for 20 ms, and then it was rapidly restretched to its original length and force allowed to redevelop. The fiber was then relaxed with relaxing solution. This procedure was repeated three times at each free [Ca2+]. F was sampled, via computer, at 1 kHz. Tension redevelopment after this procedure was fit by an exponential equation of the form
F = a ⋅ (1 − <IT>e</IT><SUP><IT>tk</IT></SUP>)
where a is the fractional force, k represents ktr, and t is the tension redevelopment duration. This approach yielded r2 values above 0.95 in all cases. Our preliminary experiments showed that the coefficient of variation for the three ktr values recorded during a single Ca2+ exposure was 5.1%. Because ktr was determined at only five levels of free Ca2+, the ktr-free [Ca2+] relationship was not computed as were those of force and AM-ATPase.

SR experiments. The homogenizing buffer contained the following (in mM): 250 sucrose, 20 HEPES (pH 7.5), 0.2 phenylmethylsulfonyl fluoride, and 2% sodium azide (NaN3). Immediately after stimulation, muscles were placed in 8 vol (wt/vol) of ice-cold buffer and minced with scissors. They were homogenized, on ice, with a Pro 200 homogenizer and 5-mm probe by using three 15-s bursts at ~12,000 rpm. The crude homogenates were then centrifuged at 1,600 g for 10 min (2°C) after which the supernatant was removed and stored at -80°C. Total protein concentrations were determined with the Bradford protocol (Bio-Rad).

The Ca2+ uptake/release buffer consisted of the following (in mM): 100 KCl, 20 HEPES, 7.5 pyrophosphate, and 0.5 Mg2+ (pH 7.0, 37°C). In addition, initial free [Ca2+] was 2 µM, and 2 µM fura 2 was added as an extravesicular Ca2+ indicator. Ca2+ uptake and release were measured by adding 200 µg of homogenate fraction protein to 1 ml of buffer. Uptake was initiated by the addition of 1 mM MgATP and was allowed to continue until little or no change in extravesicular free [Ca2+] was observed. Ca2+ release was then initiated by the addition of 25 µM AgNO3 or 5 mM 4-chloro-m-cresol (4-CMC). During this procedure, the buffer solution was continually stirred and was temperature maintained at 20°C.

Extravesicular free [Ca2+] was monitored fluorometrically by using a Jasco CAF-110 Intracellular Ion Analyzer and fura 2 (excitation 340 nm and 380 nm, emission 500 nm), and free [Ca2+] was computed by using the ratiometric method of Grynkiewicz et al. (18). Fluorescence ratios were sampled at 2 Hz (Keithly MetraByte DAS1608, 12-bit analog to digital) and stored on disk for later analysis. The rates of Ca2+ uptake and release were computed from the steepest negative and positive slopes of the extravesicular free [Ca2+] vs. time curve and normalized by the protein concentration. In triplicate assays of the same muscle sample, coefficients of variation for uptake and release were 6.2 and 4.9%, respectively.

Ca2+-ATPase activity was measured in a solution containing the following (in mM): 200 KCl, 20 HEPES, 10 MgCl2, 3 PEP, 0.6 NADH, and 1 EGTA as well as 2 µM ionophore A-23187, 7.5 U/ml PK, and 5 U/ml LDH (pH 7.0, 37°C). Homogenate protein (100 µg) was added to the incubation solution, and the reaction was started with 1 mM Na2ATP. Absorbance changes (340 nm) were monitored for 3 min, and basal activity (Mg2+ stimulated) was determined by using Beer's law and an extinction coefficient for NADH of 6,270 · M-1 · cm-1. Total activity was determined for 3 min after the addition of CaCl2 (2 µM free [Ca2+]). Ca2+-stimulated activity was computed as total minus basal.

Statistical analyses. The effects of condition (rest, fatigue) and stimulation protocol on Ca2+ uptake and release were determined by analyses of variance adjusted for repeated measures made on contralateral muscles. Significance was set at the P < 0.05 level of confidence.

    RESULTS
Top
Abstract
Introduction
Methods
Results
Discussion
References

Fatigue effects on the contractile apparatus. As we have demonstrated previously (28, 31), this protocol of repetitive, tetanic stimulation reduced Po with a rate constant of ~53 s-1 eventually reaching 3-5% of initial within 3 min. In the first set of experiments, the effects of fatigue on force and AM-ATPase of the contractile apparatus were examined by rapidly preparing skinned fibers from both rested and fatigued muscles. The results of these experiments are summarized in Table 1. In fibers taken from fatigued muscles, neither Fmax nor N was significantly different from values of those taken from contralateral rested muscles. However, the [Ca2+]50 of force was significantly lower in the fatigued fibers. This alteration in force production was associated with a 20% reduction in Amax and no alteration in the Ca2+ sensitivity or the slope of ATPase-free [Ca2+] relationship.

                              
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  		Table 1.   Effects of stimulation on contractile-apparatus function

Typical relationships between force and AM-ATPase activity in rested and fatigued fibers are depicted in Fig. 1. In this example, data obtained by using fibers taken from contralateral rested and fatigued muscles are shown. As reported previously (22), there is a clear separation of the force and AM-ATPase curves, with the [Ca2+]50 of force being somewhat greater than that of AM-ATPase. Because of the increase in Ca2+ sensitivity of force in the fatigued fibers and the nonsignificant alteration in that of AM-ATPase, the separation of the two curves is smaller under this condition. Accordingly, the difference in the [Ca2+]50 between force and AM-ATPase was 0.983 ± 0.090 (SE) µM in rested fibers compared with 0.484 ± 0.042 µM in the fatigued fibers (P < 0.05).


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  		Fig. 1.   Typical effects of fatigue on contractile apparatus force and actomyosin (AM)-ATPase activity. Shown are force (open circle , bullet ) and AM-ATPase (, ) in fibers obtained from a rested muscle (open circle , ) and from the contralateral fatigued muscle (bullet , ). pCa, -log free Ca2+ concentration.

In the second set of experiments, changes in ktr as the result of fatiguing stimulation were determined (Fig. 2). At full Ca2+ activation, ktr was 20.21 ± 1.06 and 19.62 ± 1.11 s-1 in rested and fatigued fibers, respectively (P > 0.05). Also, at pCa 5.0 and 4.5, ktr was not significantly different between conditions. However, at lower levels of free Ca2+ (pCa 6.0 and 5.5), ktr was significantly lower in the fatigued fibers than in rested fibers. It should be pointed out that fiber length rather than sarcomere length was controlled during the measurement of ktr. At 20°C, homogeneity of the laser diffraction pattern of frog fibers is good until force reaches ~50% of Fmax (Williams and Ward, unpublished observations). Thereafter, it becomes increasingly diffuse, such that adequate determination and control of sarcomere length are not possible. For this reason, absolute ktr values could be somewhat faster than those presented here (6). However, the lack of sarcomere length control should not markedly influence comparisons made between conditions (see DISCUSSION).


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  		Fig. 2.   Changes in rate constant of tension development (ktr) with Ca2+ activation in rested (open circle ) and fatigued fibers (bullet ). Values were normalized by those recorded under maximal Ca2+ activation (pCa 4.0). Values are means ± SE; n = 6 muscle pairs. * P < 0.05 between conditions.

By using the model of Huxley (21), Brenner (4, 5) proposed that Ca2+ regulation of F by the contractile apparatus occurs via the kinetics of cross-bridge cycling between the strong-binding, force-generating and the weak-binding, non-force-generating states. In this paradigm, fapp represents the rate constant for the transition to strong binding and gapp is the rate constant for transition to weak binding. The fraction of cross bridges in the strong-binding, force-generating state (FS) can be defined as
F<SUB>S</SUB> = <IT>f</IT><SUB>app</SUB>/(<IT>f</IT><SUB>app</SUB> + <IT>g</IT><SUB>app</SUB>) (1)
and isometric F as
F = F<SUB>avg</SUB> ⋅ [M] ⋅ <IT>A</IT> ⋅ <IT>L</IT><SUB>½</SUB> ⋅ <IT>f</IT><SUB>app</SUB>/(<IT>f</IT><SUB>app</SUB> + <IT>g</IT><SUB>app</SUB>) (2)
where [M] is the concentration of myosin per fiber liter, A is the fiber cross-sectional area, L1/2 is the length of one-half sarcomere and Favg is the average force of a single myosin head (4, 5, 22). Assuming that one ATP molecule is hydrolyzed per cross-bridge cycle, AM-ATPase can be defined as
AM-ATPase = [M] ⋅ <IT>A</IT> ⋅ <IT>a</IT> ⋅ <IT>L</IT><SUB>½</SUB> 
⋅ <IT>g</IT><SUB>app</SUB> ⋅ [<IT>f</IT><SUB>app</SUB>/(<IT>f</IT><SUB>app</SUB> + <IT>g</IT><SUB>app</SUB>)] (3)
where a is the number of half-sarcomeres within the fiber. Brenner (4, 5) also states that ktr is equal to the sum of the two rate constants
<IT>k</IT><SUB>tr</SUB> = <IT>f</IT><SUB>app</SUB> + <IT>g</IT><SUB>app</SUB> (4)
With the use of Eqs. 2-4, the ratio of ATPase to F is proportional to gapp and the product of F and ktr is proportional to fapp
AM-ATPase/F = <IT>g</IT><SUB>app</SUB> ⋅ (<IT>a</IT>/F<SUB>avg</SUB>) (5)
F ⋅ <IT>k</IT><SUB>tr</SUB> = <IT>f</IT><SUB>app</SUB> ⋅ F<SUB>avg</SUB> ⋅ [M] ⋅ <IT>A</IT> ⋅ <IT>L</IT><SUB>½</SUB> (6)
By using these equations, both fapp and gapp were estimated from the force, ktr, and AM-ATPase data. For individual fibers, gapp at maximal Ca2+ activation was computed by using Eq. 3, assuming 154 µmol of myosin heads per liter of fiber (14) and FS = 0.95 (22). The value of fapp was calculated by using Eq. 4 and the derived value of gapp. At intermediate free [Ca2+], gapp values were computed using the ratio of fractional AM-ATPase to F as described in Eq. 5. Submaximal values of fapp were computed by using the product of fractional F and ktr as described by Eq. 6.

Figure 3 shows the estimated values of fapp and gapp in rested and fatigued fibers. For simplicity, we calculated values only at the free [Ca2+] where ktr was measured. As shown by Brenner (4, 6), fapp increased with increasing free [Ca2+]. More importantly, fapp was not significantly different between conditions. Conversely, gapp decreased with increasing free [Ca2+] in a manner similar to that shown by Kerrick et al. (22). In addition, gapp was significantly larger in the rested fibers than in the fatigued fibers. The difference was small at higher free [Ca2+] but was considerable at lower free [Ca2+].


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  		Fig. 3.   Changes in fapp (open circle , bullet ) and gapp (, ) as a function of free Ca2+ concentration in rested (open circle , ) and fatigued fibers (bullet , ). See text for definitions of fapp and gapp. Values were computed for individual fibers at free-Ca2+ levels used in ktr experiments. In some cases, symbol hides SE bars. For gapp calculations, n = 8 muscle pairs; for fapp calculations, n = 6 muscle pairs. * P < 0.05 between conditions.

It is important to point out that the values of fapp and gapp determined here agree well with the work of Brenner (4). He reports values obtained under maximal Ca2+ activation at 5 and 15°C. These predict rate constants of 22-24 s-1 for fapp and 2.3-2.8 for gapp at 20°C.1 Our estimate of fapp in rested fibers (17.6 s-1) is somewhat lower than predicted, possibly due to our inability to adequately control sarcomere length. However, our estimate of gapp in rested fibers (2.6 s-1) is within the range predicted from Brenner's data. This suggests that despite potential limitations to our measurements (i.e., sarcomere-length control), our values of fapp and gapp are reasonable.

Fatigue effects on the SR. The use of a skeletal muscle homogenate fraction for assessing SR function was validated by examining the effects of various Ca2+-ATPase and release-channel inhibitors and activators on SR function. Our preliminary work showed that the inclusion of cyclopiazonic acid in the incubation medium completely abolished Ca2+ uptake and Ca2+-stimulated Ca2+-ATPase activity. In addition, Ca2+ release by AgNO3 was attenuated by the reducing agent dithiothrietol, and AgNO3- and 4-CMC-induced releases were completely blocked by tetracaine. Taken together, these findings indicate Ca2+ transport rates measured in this preparation are reflective of SR Ca2+ uptake and release rather than some nonspecific Ca2+ binding and/or release by non-SR organelles or proteins.

Repetitive stimulation substantially depressed the Ca2+ transport capabilities of the SR. The peak rate of Ca2+ uptake was significantly reduced by 46% in the fatigued fibers (Fig. 4). However, the amount of Ca2+ sequestered during loading was not significantly different between conditions (Fig. 4, inset). Associated with the depression in Ca2+ uptake rate, the rates of Ca2+ release evoked by AgNO3 and 4-CMC were significantly reduced by 38 and 45%, respectively, in the fatigued samples. Conversely, the amounts of Ca2+ released by the agents were not different between conditions.


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  		Fig. 4.   Effects of stimulation on rates of SR Ca2+ uptake as well as AgNO3- and 4-chloro-m-cresol (4-CMC)-stimulated release in rested () and fatigued () muscles. n = 8 Muscle pairs. * P < 0.05 between conditions. Inset: effect of stimulation on amount of Ca2+ sequestered and released.

In addition to changes in SR Ca2+ transport rates, Ca2+-ATPase activities in rested and fatigued muscles were altered by repetitive stimulation (Fig. 5). As can be seen, there was no significant difference in basal activity between conditions. However, Ca2+-stimulated activity was significantly reduced by 21% in the fatigued muscles.


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  		Fig. 5.   Effects of stimulation on basal and Ca2+-stimulated Ca2+-ATPase activity in rested () and fatigued () muscles; n = 8 muscle pairs. * P < 0.05 between conditions.

Table 2 shows comparisons of the rates of Ca2+ uptake and release determined in this study, which used the homogenate fraction, and the rates determined by Williams et al. (31), who used skinned fibers. In the study by Williams et al., the rate constant of Ca2+ uptake (kCa) was estimated from caffeine contractures evoked after loading the SR for various time intervals. The relative reduction in kCa after fatigue was statistically greater than the depression of uptake rate determined by the homogenate fraction. However, the difference between techniques was small. In our earlier study, the rate of Ca2+ release was assessed by the rate of caffeine-induced (8 mM) force increase evoked after maximal Ca2+ loading. The depressions in release rate as determined by using the homogenate fraction and skinned fibers were not significantly different. Thus it appears that reductions in SR Ca2+ uptake and release are responsible for the changes in caffeine-evoked force reported earlier (31).

                              
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  		Table 2.   Comparison of the changes in SR function determined by using homogenate fraction and skinned fibers

    DISCUSSION
Top
Abstract
Introduction
Methods
Results
Discussion
References

Our earlier studies show that fatiguing stimulation of frog semitendinosus muscle results in changes in skinned-fiber responses to Ca2+ and caffeine. In fatigued fibers, we found increased Ca2+ sensitivity of force and diminished responses to caffeine challenge. We now provide evidence that the fatigue-induced increase in Ca2+ sensitivity results from alterations in cross-bridge cycling kinetics, specifically a reduction in gapp. Also, the fatigue-induced changes in the force responses to caffeine appear to reflect intrinsic reductions in the rates of SR Ca2+ uptake and release.

Contractile apparatus function. In contrast to the present data, others have demonstrated that acute muscular activity has little effect on myofibrillar-ATPase activity. Turcotte et al. (26) showed that neither maximal activity nor its Ca2+ sensitivity were altered after electrical stimulation of rat plantaris muscle. Also, Fitts et al. (15) showed no effect of 8 h of swimming on basal or maximal activity in the soleus or extensor digitorum longus muscles. However, there are two factors that may account for the discrepant results. First, in the studies of Turcotte et al. (26) and Fitts et al. (15), Po was reduced by only 17 and 26-74%, respectively. In our study, it was reduced by ~95%. It is possible that the smaller reductions in Po, are not associated with changes in AM-ATPase activity, whereas large reductions are. This notion is supported by the finding that reductions in Po of >= 90% are needed to alter the Ca2+ sensitivity of force production by the contractile apparatus (28, 31). Second, the previous studies (6, 15) determined myofibrillar-ATPase activity by using a homogenate fraction that contains isolated myofibrils. In the present study, we used skinned fibers containing structurally intact myofibrils. Perhaps the preparation of isolated myofibrils results in the loss of some structural or regulatory protein that affects myofibrillar-ATPase activity such that the effects of fatigue are masked.

Measurements of force, AM-ATPase, and ktr provide insight into the factors that contribute to the fatigue-induced increase in the Ca2+ sensitivity of force production. By using these measurements, we have determined that gapp is slowed in fatigued fibers, an effect that is more pronounced at intermediate free [Ca2+] than at maximal activating [Ca2+]. As we have made a number of assumptions in our calculations of fapp and gapp, it is possible that we have overestimated the difference in gapp between conditions. First, we assumed that at maximal Ca2+ activation there are 154 µmol of myosin heads per liter (14) and that FS = 0.95 (22). If either of these values is different in the fatigued compared with rested fibers, gapp could be underestimated. To account for the 20% reduction AM-ATPase and gapp (pCa 4.0), one or a combination of these variables would have to have been altered by a similar percentage. However, we do not think that such is the case. If differences in either parameter between rested and fatigued fibers account for the reduction in gapp in the fatigued fibers, then Fmax would be reduced by a similar degree (Eq. 2). As shown in Table 1, Fmax was not different between conditions. Second, the calculation of gapp at intermediate free [Ca2+] from AM-ATPase/F assumes that Favg is similar in rested and fatigued fibers. If Favg is greater in fatigued fibers at intermediate free [Ca2+], then gapp would be underestimated. Again, we think that this is an unlikely scenario. Brenner (4, 5) suggests that Favg does not vary as a function of free [Ca2+]. Thus, if Favg is greater in the fatigued fibers at moderate free [Ca2+], then it would also be greater at maximally activating free [Ca2+]. As a result, Fmax would be increased, again inconsistent with our data. Third, at free [Ca2+] eliciting force responses >= 50% of Fmax, we have observed increasing sarcomere-length heterogeneity. If the extent of heterogeneity is different between rested and fatigued fibers, then our estimates of the difference in gapp between conditions would be inflated. It is important to point out that the greatest difference in gapp between conditions was observed at pCa 6.0 where force is <20% of Fmax and sarcomere heterogeneity is minimal. Furthermore, Brenner and Eisenberg (6) show that increased heterogeneity has a greater effect on ktr than on AM-ATPase. We found that, at maximal Ca2+ activation, ktr was not different between conditions, whereas AM-ATPase was reduced in fatigued fibers. As a result, we do not expect that sarcomere-length heterogeneity in our preparation accounts for the reduced gapp in fatigued fibers. Taking the above arguments into account, we propose that our force, AM-ATPase, and ktr data indicate that gapp is reduced in skinned fibers taken from fatigued muscle, an effect that is greater at intermediate free [Ca2+] than under conditions of maximal Ca2+ activation.

The consequence of reduced gapp is that, during the cross-bridge cycle, the cross bridges remain in the force-generating state for a longer period of time, resulting in increased force. Under conditions of maximal Ca2+ activation, this effect would be minimal because gapp is relatively small compared with fapp. As a result, Fmax would not be markedly altered. However, at intermediate free [Ca2+], gapp is larger and fapp is smaller than at maximal Ca2+ activation (4, 6). As a consequence, gapp exerts greater influence over force. With a reduction in gapp, submaximal force would be increased, resulting in increased Ca2+ sensitivity. The idea that changes in gapp can affect the Ca2+ sensitivity of force was recently demonstrated by Kerrick et al. (22). They showed that reducing MgATP concentration slows gapp, resulting in a marked decrease in [Ca2+]50 of force and little change in Fmax. Thus we propose that the increased Ca2+ sensitivity of contractile apparatus force that accompanies fatigue results from a reduction in gapp.

SR function. Others have used muscle homogenate fractions to show depressions (3, 17) and no change (10) in SR Ca2+ uptake and Ca2+-ATPase activity after exercise leading to fatigue. It is possible that the discrepancies between our study and the above investigations are due to differences in sample preparation. However, we do not think that such is the case (see below). There are a number of other possibilities such as differences in species examined, assay conditions, and exercise protocols. It remains to be seen if any of these later factors alter the potential effect of exercise on SR function.

A unique aspect of this investigation is that we have now demonstrated fatigue-induced reductions in SR Ca2+ handling in frog semitendinosus by using two different methods, skinned fibers (28, 31) and a homogenate fraction (this investigation). In the saponin skinned-fiber preparation (12), the sarcolemma is permeabilized and the SR is left intact. In the homogenate technique, the SR is disrupted and then is reassembled into vesicles. There are unique advantages and disadvantages associated with these preparations. In the skinned-fiber preparation, the SR is examined in a more physiological state (i.e., not disrupted). Unfortunately, when the caffeine contracture method is used, SR Ca2+ handling is inferred from contracture forces and factors other than SR function could influence force including contractile apparatus Ca2+ sensitivity. In the homogenate fraction, a considerable portion of the SR is lost during centrifugation (G. A. Klug, personal communication) and Ca2+ handing must be examined in a nonphysiological state (i.e., vesicles). Despite this, direct measurements of Ca2+ movements can be easily obtained. The fact that similar fatigue-induced reductions in the rates of Ca2+ uptake and release were found with use of the two different preparations suggests that they are not artifacts resulting from sample preparation. We also show that three different compounds, caffeine, AgNO3, and 4-CMC reveal fatigue-induced reductions in SR Ca2+ release. It is interesting to note that similar reductions in the release rate were obtained with all three compounds. This suggests that the effects of fatigue on the Ca2+-release process is complex, probably involving several mechanisms. Taken together, our investigations and previous investigations that used more purified SR preparations strongly suggest that fatigue results in intrinsic alterations SR function.

Summary. We show that the development of fatigue is associated with intrinsic alterations in the functional properties of the contractile apparatus and SR. The increased Ca2+ sensitivity of force production and reduced Amax appear to result from a decrease in cross-bridge cycling kinetics, specifically a reduction in gapp. In addition, the changes in SR function are manifest as depressions in the rates of Ca2+ uptake and release and ATP hydrolysis. Without doubt, these changes have important implications for force production and energy consumption in fatigued muscle (see Ref. 29). It remains to be seen what factors trigger these alterations. The changes reported here probably do not result from the direct effects of metabolite accumulation because both structures were removed from the fatigued, intracellular environment and were studied under conditions that simulate a rested cell. Recently, conditions associated with fatigue such as elevated resting [Ca2+]i (8, 23, 28) and reactive oxygen species (7) have been shown to cause long-term alterations in SR function in muscle. It is also possible that some component that is normally associated with the contractile apparatus or SR that influences function is lost or depleted in fatigued muscle. For example, the loss of muscle glycogen may affect the release and uptake of Ca2+ by the SR (9). Clearly, effort directed toward identifying specific factors that initiate fatigued-induced intrinsic changes in contractile apparatus and SR function is warranted.

    ACKNOWLEDGEMENTS

The authors thank Dr. W. Glenn L. Kerrick for help with simultaneous measurements of force and AM-ATPase activity.

    FOOTNOTES

This project was supported by National Institute of Arthritis and Musculoskeletal Skin Diseases Grant AR-41727.

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. §1734 solely to indicate this fact.

1 At 5°C, fapp = 3.0-3.5 s-1 and gapp = 1.0-1.5 s-1. At 15°C, fapp = 12 s-1 and gapp = 2 s-1 (4). This results in Q10 values of 3.4-4.0 and 2.0-1.3, respectively.

Address for reprint requests: J. H. Williams, Dept. of Human Nutrition, Foods, and Exercise, Virginia Tech, Blacksburg, VA 24061-0430 (E-mail: jhwms{at}vt.edu).

Received 6 February 1998; accepted in final form 8 April 1998.

    REFERENCES
Top
Abstract
Introduction
Methods
Results
Discussion
References

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