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Institute of Sports Science and Clinical Biomechanics, University of Southern Denmark, Odense University, Denmark
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ABSTRACT |
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 TOP
 ABSTRACT
 INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
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The purpose of
the study was to characterize the sarcoplasmic reticulum (SR) function
and contractile properties before and during recovery from fatigue in
the rat extensor digitorum longus muscle. Fatiguing contractions (60 Hz, 150 ms/s for 4 min) induced a reduction of the SR Ca2+
release rate to 66% that persisted for 1 h, followed by a gradual recovery to 87% of prefatigue release rate at 3 h recovery.
Tetanic force and rate of force development (+dF/dt) and
relaxation (
dF/dt) were depressed by ~80% after
stimulation. Recovery occurred in two phases: an initial phase, in
which during the first 0.5-1 h the metabolic state recovered to
resting levels, and a slow phase from 1-3 h characterized by a
rather slow recovery of the mechanical properties. The recovery of SR
Ca2+ release rate was closely correlated to
+dF/dt during the slow phase of recovery
(r2 = 0.51; P < 0.05).
Despite a slowing of the relaxation rate, we did not find any
significant alterations in the SR Ca2+ uptake function.
These data demonstrate that the Ca2+ release mechanism of
SR is sensitive to repetitive in vitro muscle contraction. Moreover,
the results indicate that +dF/dt to some extent depends on
the rate of Ca2+ release during the slow phase of recovery.
extensor digitorum longus; muscle fatigue; Ca2+-ATPase; rate of force development; recovery
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INTRODUCTION |
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
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REPEATED MUSCLE CONTRACTIONS are often accompanied by a progressive loss of force-producing capacity. The precise mechanisms that mediate fatigue have yet to be identified. However, a growing amount of evidence implicates alteration in intracellular Ca2+ handling as a major contributor in fatigue (3, 30). Nevertheless, knowledge about the direct effect of exercise on sarcoplasmic reticulum (SR) Ca2+ handling is limited. Repetitive stimulation of single fibers to fatigue is accompanied by a reduction in maximum Ca2+-activated force, decreased Ca2+ sensitivity, and a declined tetanic cytosol free Ca2+ concentration ([Ca2+]) (3, 30). It has for a long time been proposed that the observed decline in tetanic free [Ca2+] may be due to a decrease in Ca2+ release from SR (30). Recently, Ward et al. (29) and Favero et al. (13) have shown that the SR Ca2+ release rate is diminished with fatigue. These measurements are done in vitro under optimal conditions, indicating that, after the SR is removed from the intracellular environment, changes in function persist. Hence, the depressed Ca2+ release rate is due not only to changes in the intracellular environment, e.g., decreased pH, metabolic disturbances, and altered ion homeostasis. However, an exercise-induced altered intracellular environment impairs both in vivo cross-bridge cycling (23) and SR function (15). Most studies of SR function in relation to exercise have examined changes in direct relation to fatigue; however, data comparing the Ca2+ regulation at various states of recovery after fatigue are scarce. Because the contraction-induced altered metabolic environment is recovered within the initial (0.5-1 h) recovery, the metabolic parameters can be excluded as modifiers of the contractile apparatus after the initial recovery phase. The inclusion of recovery may offer an opportunity to further establish the relationship between contractile properties in vivo and SR function. Thus the deleterious effects of the intracellular metabolic alterations on the contractile function per se can be separated from the changes in SR function.
We have hypothesized that if changes in force during fatiguing contractions are due at least in part to diminished SR Ca2+ release rates, then alterations in these parameters would parallel each other. The question was addressed by measuring the SR Ca2+ release and uptake rates as well as Ca2+-ATPase activity in relation to contractile function at fatigue and in the recovery phase.
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METHODS |
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
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Animals
Male Sprague-Dawley rats (n = 28) purchased from the Institute of Biomedicine, Odense University Hospital, were used in the study. The rats were ~4 wk old and weighed 87 ± 2 g. Animals were housed in box cages, four to five animals per cage, and maintained in a temperature-controlled room (22 ± 1°C) with a 12:12-h light-dark cycle (light 0800-2000). The rats were provided unrestricted access to food and water.Experimental Procedure
In vitro muscle preparation. Rats were weighed and killed by cervical dislocation, and both extensor digitorum longus (EDL) muscles [98% fast-twitch fibers (2), weight ~40 mg] were quickly excised and placed in Krebs-Ringer bicarbonate buffer containing (in mM) 120.1 NaCl, 25 NaCHO3, 4.7 KCl, 1.2 KH2PO4, 1.2 MgSO4, 1.3 CaCl2, and 5 D-glucose (pH 7.3). A small loop of surgical silk was tied to each tendon of the muscle, and the muscles were mounted vertically between a fixed hook and an isometric force transducer (K30 type 351, Hugo Sachs Elektronick, March-Hugstetten, Germany) in a temperature-controlled chamber at 30°C (Schuler organbad, Hugo Sachs Elektronick). The muscle chamber contained Krebs-Ringer bicarbonate buffer (pH 7.3) with 5 mM D-glucose and was continuously gassed with a mixture of 95% O2-5% CO2. Contractions were electrically evoked through the bathing solution by two platinum electrodes located close to either side of the muscle. The muscles were gently adjusted to the length at which maximum force with a single twitch stimulus was recorded, after which a control tetanus was evoked (60 Hz, 1.5-s duration) to ensure that the muscle was properly mounted, i.e., 20% deviation from the expected force output compared with rat size. Subsequently, the muscles were allowed to equilibrate for 30 min without stimulation.
Test tetanus. To evaluate the mechanical properties of the muscle, a maximum tetanic contraction was induced by trains of 0.1-ms pulses delivered at 100 Hz for 1.5 s (test tetanus). All stimulations were delivered with 10 V. Preliminary experiments showed that this stimulation elicits a maximum force output, which can be repeated at 2-min intervals for at least 10 min without affecting the force characteristics.
Fatigue protocol. Before the start of the experiment, muscles were exposed to the test tetanus (prefatigue) and randomly either used as controls or subjected to a 4-min fatigue protocol in which contractions were evoked by 60 Hz on a duty cycle of 150 ms every second. After the fatigue protocol, the muscles were randomly assigned to one of five groups (n = 8 in each group) in which the test tetani were elicited either immediately after fatiguing stimulation (0 h recovery) or after 0.5 h, 1 h, 2 h, or 3 h of recovery, respectively. Control muscles (n = 8) were evoked with test contractions at the same time points but were not subjected to the fatigue protocol.
Analytical Procedures
Mechanical properties.
The mechanical properties monitored during the test tetanus included
peak tension (Fmax), half peak tension time
(PT1/2), half relaxation time (RT1/2), maximum
rate of force development (+dF/dt) and relaxation
(dF/dt), and latent time. Latent time was defined as the
time from stimulation until the start of contraction, defined as a
signal that was 30% higher than the noise level. The muscle
stimulation and subsequent on-line collection of mechanical data were
obtained through a computerized data acquisition system (LabVIEW 5.0, National Instruments). The setup enabled the stimulation, measurements,
and analysis of the mechanical properties during each contraction.
Muscle preparation.
The muscles were allowed to rest for 2 min after the test tetanus to
ensure minimal effect of the contraction and then were quickly removed
and divided in two specimens of ~20 mg. One-half of the muscle was
immediately frozen in liquid nitrogen and stored at 80°C until
freeze dried and analyzed for ATP, phosphocreatine (PCr), glycogen, and
lactate content. The other half of the muscle was put in ice-cold
homogenizing buffer containing (in mM) 300 sucrose, 1 EDTA, 10 NaN3, 40 Tris-base, and 40 L-histidine (pH 7.8) (24). The muscle was homogenized in 30 wt/vol (~20
mg muscle in 600 µl buffer), minced with scissors, and homogenized
with a Omni 2000 homogenizer (20.000 rpm) in three 15-s bursts
separated by 15-s pauses between each burst. The muscles were kept on
ice during the whole procedure. Aliquots of 200 µl were frozen in liquid N2 and stored at 80°C until analysis. Whole
homogenate protein content was assessed by using a standard kit (Pierce
bicinchoninic acid protein reagent no. 23225).
Analysis of SR function. The evaluation of the SR function involved measurements of the SR Ca2+-ATPase capacity and the SR Ca2+ uptake and release rates. The Ca2+-ATPase activity was measured under constant, nearly optimal conditions and hence was defined as SR Ca2+-ATPase capacity. All measurements were carried out on whole homogenates and expressed relative to the whole homogenate protein content.
SR Ca2+-ATPase capacity was measured according to the method of Simonides and van Hardeveld (27). In brief, the NADH expenditure was followed spectrophotometrically (Beckmann DU 600, Fullerton, CA) at 37°C for 2 min. The method was modified because SR Ca2+-ATPase capacity was taken to be the difference between the total and the background ATPase capacities, measured in the presence of 25 µM Ca2+ and 1 mM EGTA, respectively. All measurements were carried out with a sample concentration of ~3 mg tissue/ml buffer in the presence of 5 µM of the Ca2+ ionophore A-23187. Activities are expressed as micromoles per minute per gram of whole homogenate protein. SR Ca2+ uptake and release rates were analyzed by use of the fluorescent Ca2+ indicator indo 1 as described by Ruell et al. (26). Analysis was performed on a fluorometer (Ratiomaster RCM, Photon Technology International, Brunswick, NJ) with thermostated cuvette holder at 37°C and continuous stirring by a magnetic bar. The excitation wavelength was 355 nm, and the emission was continuously measured at 400 and 470 nm. The assay buffer consisted of 165 mM KCl, 22 mM HEPES, 7.5 µM oxalate, 11 µM NaN3, 5.5 µM TPEN, 20 µM CaCl2, and 1 mM MgCl2 (pH 7.0). ATP and indo 1 were added to a final concentration of 2 mM and 1 µM, respectively, before the assay was initiated by adding ~4 mg tissue to 2 ml of assay buffer. The subsequent Ca2+ uptake was followed until it plateaued. Thapsigargin (Sigma T-9033) was added to a final concentration of 1 µM and was incubated for 30 s to block the SR Ca2+-ATPase, before 1 mM 4-chloro-m-cresol (4-CMC) was used to induce a rapid spike of Ca2+ release from the vesicles. Ratiometric data were sampled at 2 Hz and converted to free Ca2+ concentration of the buffer [(Ca2+]free) according to the method of Grynkiewicz et al. (19). A dissociation constant of 142 nM for the binding of indo 1 and Ca2+ was measured by using a standard Ca2+ calibration buffer kit with Mg2+ (Molecular Probes, Eugene, OR). The resulting curve was smoothed over 15 points (Savitsky-Golay algorithm) and differentiated (point-to-point slope) to determine the rate of Ca2+ release. Average rates of uptake were determined in 100 nM intervals between 100 and 600 nM free Ca2+. The 4-CMC-stimulated Ca2+ release rate was determined as the peak of the first derivative of the [Ca2+]-vs.-time curve (Fig. 1). Because the assay buffer employed with this method contains various Ca2+ chelators (i.e., oxalate, homogenate protein, and EDTA), the Ca2+ buffering capacity is strongly dependent on the [Ca2+] at which the measurements are made. The values obtained for SR Ca2+ uptake and release rates are relative and are therefore expressed as arbitrary units of Ca2+ per minute per gram of protein. As a result, SR Ca2+ uptake and release rates cannot be compared unless measured at similar [Ca2+]free and buffering conditions. For the measurement conditions employed in the present study, the values reported had the dimension micromoles per gram protein per minute. All measurements were performed in duplicate.
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Validation of SR Ca2+ handling measurements. To validate the method for determining SR function, the dose dependency of 4-CMC-induced Ca2+ release was established, and the reproducibility of methods to determine SR Ca2+ uptake and release rates and SR Ca2+-ATPase activity was assessed.
A total of 14 EDL muscles taken at random from right and left legs of rats (~150 g body wt) were each analyzed by using three to five different concentrations of 4-CMC ranging from 0.5 to 20 mM, and eight muscles were analyzed by using 141 µM AgNO3 as release agent. The Ca2+ release rates (means ± SE in arbitrary units Ca2+ · min
1 · g1 protein) when 4-CMC was used were 0.80 ± 0.15 at
0.5 mM (n = 6), 1.77 ± 0.17 at 1 mM
(n = 9), 2.39 ± 0.34 at 2 mM (n = 7), 3.60 ± 0.10 at 5 mM (n = 14), 3.55 ± 0.2 at 10 mM (n = 7), and 3.59 ± 0.17 at 20 mM
(n = 5). When AgNO3 was used, the
Ca2+ release rate was 1.76 ± 0.80 arbitrary units
Ca2+ · min1 · g1
protein. The data show that a maximum SR Ca2+ release rate
was attained at concentrations of 5 mM 4-CMC and above. The release
rate at 1 mM 4-CMC was similar to the release rate when using 141 µM
AgNO3, both of which elicited about 50% of maximum release
rate. Measurements of 4-CMC-evoked SR Ca2+ release using
the submaximal concentration of 1 mM 4-CMC to evoke SR Ca2+
release, as in the present study, reflect a combination of the number
of functional release channels and the sensitivity of the channels to
4-CMC. Hence, changes in each of these variables may cause a change in
measured release rate.
Coefficients of variation (CV) for the rates of Ca2+ uptake
and 5 mM 4-CMC-evoked Ca2+ release were assessed as the CV
of 13 pairs of resting rat soleus and EDL muscles from left and right
leg of the same rat. The CV of Ca2+ uptake measurements was
7.6% (EDL) and 11.7% (soleus) for uptakes between 100 and 200 nM
[Ca2+]free and 4.2% (EDL) and 7.1% (soleus)
for uptakes between 500 and 600 nM
[Ca2+]free. The CV values for the peak
4-CMC-induced SR Ca2+ release rate when 5 mM were used were
determined to be 9.6% (EDL) and 15.1% (soleus). Given the sensible
assumption that SR function is equal in left and right muscles from the
same rat, the CV includes both analytical and biological variation. The
coefficient of variation for duplicate determinations (CVd)
of 1 mM 4-CMC-evoked SR Ca2+ release rate, pipetted from
the same homogenate, was 8.6% (EDL) and 12.3% (soleus). CV
for the Ca2+-ATPase activity, determined from nine pairs of
soleus muscles, was 9.9%. CVd for Ca2+-ATPase,
determined from the 55 pairs of EDL muscles used in the present study,
was 8.2%.
Measurements of metabolites. Muscle metabolites were extracted from the freeze-dried muscle samples by perchloric acid treatment and were analyzed for the total content of ATP, PCr, glycogen, and lactate by fluorometric assays according to the procedure of Lowry and Passonneau (22).
Statistics
Values are presented as means ± SE, and significance was tested at the 0.05 level. Statistical comparisons were performed with one-way ANOVA. The reproducibility of the methods used was calculated as
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RESULTS |
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TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
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Muscle mechanical properties, fatigue protocol. Force output declined significantly during the 4-min stimulation protocol (60 Hz, 1-ms pulses, 150 ms/s). The mean prefatigue force of 26.8 ± 1.8 g had declined to 72 ± 3% and 49 ± 3% after 1 and 2 min of stimulation, respectively, and decreased further to 14 ± 2% at 4 min. The +dF/dt increased significantly to 128 ± 7% after 20 s of stimulation; this increase was followed by a pronounced decline to 38 ± 6% and 20 ± 3% after 2 and 4 min of stimulation, respectively. Rate of relaxation followed a similar pattern, averaging 114 ± 4%, 17 ± 3%, and 12 ± 1%, respectively. Altogether, these mechanical properties provide good evidence for the development of fatigue.
Muscle mechanical properties, test tetanus.
The mechanical properties obtained during the test tetanus (100 Hz,
1.5-s duration) at prefatigue, at 0 h recovery, and until 3 h
recovery appear in Fig. 2. All values are
expressed as percents of prefatigue values. The absolute values at
prefatigue averaged 42.4 g (Fmax), 3.6 g/ms
(+dF/dt), 4.4 g/ms (dF/dt), 12.4 ms (PT1/2), 9.7 ms (RT1/2), and 1.69 ms (latent
period), respectively. Fmax was depressed to 20 ± 1%
at 0 h recovery but recovered to 64 ± 1% of prefatigue
force during the first 0.5 h of recovery and to 73 ± 1%
after 1 h recovery. Subsequently, Fmax recovered more
slowly to 85 ± 1% force after 3 h recovery; this was termed the slow phase of recovery (Fig. 2). The +dF/dt and
dF/dt showed a pattern similar to Fmax, and,
when ±dF/dt was expressed relative to Fmax, no
significant changes occurred in these variables. An initial recovery of
+dF/dt and dF/dt from 21 ± 1% and
18 ± 1%, respectively, immediately after stimulation to 56 ± 1% and 65 ± 1% after 0.5 h recovery was followed by a
slower phase from 1 h until nearly full recovery after 3 h
(84 ± 1% and 86 ± 1% prefatigue +dF/dt and
dF/dt, respectively). The PT1/2 was not significantly changed by the fatigue protocol, the only exception being
a 5 ± 1% increase (P < 0.05) after 0.5 h
recovery. RT1/2 and latent time both increased immediately
after the fatiguing stimulation (116 ± 3% and 132 ± 3%,
respectively; P < 0.05), but both properties were back
to prefatigue value after 0.5 h recovery.
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SR Ca2+ release rate.
The fatigue protocol induced a decrease in the SR Ca2+
release rate to 66 ± 2% that persisted for 1 h and was
followed by a gradual recovery to 87 ± 5% of prefatigue release
rate 3 h after the fatiguing contractions (Fig.
3). There were no significant differences
after 3 h recovery between controls and stimulated muscles or
between prefatigue release rate for the stimulated muscles and for the
3-h incubated controls. There was a significant correlation between the
SR Ca2+ release rate and +dF/dt
(r2 = 0.51; P < 0.05) at
prefatigue and after 1 h, 2 h, and 3 h recovery, i.e.,
the slow phase of recovery (Fig. 4).
Values from 0 h and 0.5 h recovery were excluded because
muscle metabolites were not fully recovered back to prefatigue levels.
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SR Ca2+ uptake function.
The effects of fatiguing protocol on SR Ca2+ uptake
function, measured as Ca2+ uptake rate and
Ca2+-ATPase capacity, are shown in Fig.
5. The Ca2+ uptake rates,
measured in 100 nM intervals between 100 to 600 nM intracellular
[Ca2+]free, were unaffected by
the stimulation protocol at [Ca2+] higher
than 300 nM. However, the average Ca2+ uptake rate was
depressed after 0.5 h and 1 h recovery at 100-200 nM
intracellular [Ca2+]free, and after
1 h at 200-300 nM. We did not find any significant alterations in the Ca2+-ATPase capacity after fatigue or in
the total and basal ATPase capacities for any of the muscle groups.
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Muscle metabolites.
The muscle metabolite concentrations were all significantly
(P < 0.05) altered with fatigue (Table
1). The muscle ATP, PCr, and glycogen
levels were depressed immediately after the fatiguing stimulations to
42%, 43%, and 65%, respectively, of prefatigue levels. The muscle
PCr content was recovered after 0.5 h recovery; however, ATP and
glycogen content was still depressed to 84% and 74% of
prefatigue levels and required 1 h to recover fully. PCr content showed a subsequent overshoot to ~125% of prefatigue content at 2 and 3 h, as was the case in the 3-h incubated control
muscles. Muscle lactate concentration increased to 143 mmol/kg dry wt, i.e., 6.5-fold increase, immediately after the 4-min stimulation, but
returned to the resting value after 1 h recovery. All measured metabolites were back to prefatigue level at 1 h of recovery, corresponding to the slow phase of recovery of Fmax.
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DISCUSSION |
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TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
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The present study demonstrates that the performance of repeated
isometric contractions for 4 min (60 Hz, 150-ms/s duty cycle) leads to
a significant reduction in Fmax and decrements in
+dF/dt and dF/dt, confirming the general
changes seen with muscle fatigue development (17,
28). The recovery of Fmax after fatigue
occurred in two phases: an initial phase within the first 0.5 to 1 h of recovery followed by a slow phase of recovery to prefatigue
tension (28). The length of each phase depends on the
degree of fatigue. The etiology of the slow recovery phase is unknown,
but it is apparently not related to sarcolemma inactivation, because
resting and action potentials are unaltered during the slow phase of
recovery (21). Furthermore, the slow recovery phase after
muscle contraction cannot be explained by metabolic alterations in the
cellular environment, because the alterations in contractile properties
are observed well after metabolic homeostasis is reestablished
(11). In the present study, Fmax and rates of
force development and relaxation were depressed for >3 h after the
4-min exercise. However, muscle metabolite environments were similar to
prefatigue values after 1 h, further suggesting that metabolic
factors were not a primary mechanism of the slow phase of recovery.
A major finding in this investigation is that 4 min of high-intensity exercise results in a significant and sustained depression in the SR Ca2+ release rate, indicating that the release mechanism of skeletal muscle SR is sensitive to repetitive in vitro muscle contraction. Recently, Ward et al. (29) demonstrated that the SR Ca2+ release rate was depressed by ~40% in frog sartorius muscles at fatigue. Furthermore, it was shown that the depression in release rate depended on the severity of fatigue more than on the stimulation protocol.
Favero et al. (13) reported a reduced rate of AgNO3-stimulated Ca2+ release in isolated rat SR vesicles after prolonged treadmill run. Moreover, Ca2+-activated ryanodine binding was depressed by ~20%, suggesting that the number of functional Ca2+ release channels is diminished by repetitive contractions. However, in the present study, an even larger decrease of 34% in Ca2+ release rate was observed, which may be accounted for at least in part by a reduced number of functional channels. Additionally, a reduced sensitivity of the channels to 4-CMC may contribute. Because the SR Ca2+ release in the above-mentioned studies was measured in vitro under optimal conditions, changes after fatigue hypothetically could be due to alterations in the structure of the SR membrane, release proteins, and/or irreversible binding of metabolic end products (i.e., Pi, lactate, H+). In the present study, the SR Ca2+ release was still depressed after 1 h recovery, whereas the muscles were fully recovered metabolically (ATP, PCr, lactate, and glycogen). This finding substantiates that metabolic factors are not the primary mechanism for the observed long-term depression of Ca2+ release. Depression of SR Ca2+ release by high lactate is alleviated once the SR is diluted into a lactate-free medium, demonstrating a reversible inhibition of the release channel (16). Taken together, these observations demonstrate that, once the SR is removed from the fatigued intracellular environment, depression in the Ca2+ release rate persists, indicating that the depression is due to structural changes in the release mechanism.
The cause of the reduced SR Ca2+ release associated with fatigue is not known at present, but various factors have been hypothesized, including reactive oxygen species (ROS) (14), elevated cytosolic Ca2+ (7, 32), and reduced muscle glycogen concentration (7). Several recent reports have stated that ROS are produced in the active muscle (25) and that the produced ROS are operative in the muscle dysfunction and altered Ca2+ regulation after exercise (12, 14, 25). Small amounts of ROS tend to stimulate the SR Ca2+ release (14); however, high concentrations of ROS target reactive sulfhydryl groups, which are abundant in SR Ca2+-ATPases and release channels (12). The relative importance of ROS tend to on the SR function is at the moment unknown. Exposure of skinned fibers to 0.5 µM free Ca2+ for 5 min depressed caffeine sensitivity of Ca2+ release, suggesting that at least a portion of the change in SR Ca2+ release seems to be due to an elevated cytosolic [Ca2+] (33). Chin et al. (7) have shown that a long series of tetani, elevating the free cytosolic Ca2+ level, resulted in a prolonged (1 h) reduction in Ca2+ release. When fibers recovered without glucose, Ca2+ release was reduced to a greater extent. This indicates that the decrease in SR Ca2+ release associated with fatigue has at least two components: a metabolic component, which, in the presence of glucose, recovers within 1 h, and a component dependent on the elevated Ca2+, which recovers more slowly. In another study, Chin and Allen (6) demonstrated that the reductions in force and Ca2+ release observed during fatigue were closely associated with reduced muscle glycogen concentration.
Another interesting finding is that the present data indicate that there is a close correlation (r2 = 0.51; P < 0.001) between the +dF/dt and the SR Ca2+ release rate in the slow phase of recovery. The corresponding data of Ca2+ release and +dF/dt were obtained by using all data points except for the values at 0 h and 0.5 h of recovery. The exclusion of these values was justified to distinguish in situ metabolic alterations on the SR Ca2+ release (6, 15) and contractile proteins (8, 23) from the nonmetabolic related alterations observed. Peak dF/dt is thought to be related to the expression of the myosin heavy and light chain isoforms in the muscle (for review, see Ref. 18). However, Kugelberg and Thornell (20) measured isometric contraction time and histochemical fiber type, as well as volume of the SR terminal cisternae, which relates to SR Ca2+ release rate. They found that the volume density of SR terminal cisternae was inversely related to isometric contraction time, irrespective of type of muscle. The present finding of a positive relationship between the relative SR Ca2+ release rate vs. +dF/dt (Fig. 4) during the slow phase of recovery supports the belief that the SR Ca2+ release characteristics may influence the +dF/dt during fatiguing conditions.
Despite a slowing of the relaxation rate after fatiguing stimulation, we did not find any significant alterations in the SR Ca2+ uptake function, measured as homogenate SR Ca2+ uptake and Ca2+-ATPase capacity. Single-fiber studies have shown that fatiguing stimulation produced slower rates of both force relaxation and Ca2+ removal from the myoplasm (1). This suggests a causal relationship between the two, although Westerblad et al. (31) have shown that slowing of relaxation in fatigued mouse fibers is caused by altered cross-bridge kinetics and not by impaired Ca2+ handling. It is the general opinion that severe exercise impairs both the Ca2+-ATPase capacity and the SR Ca2+ uptake rate (5, 33). However, considerable inconsistencies have been found in the effect of exercise on SR Ca2+ uptake function (5, 9, 10, 11, 29, 32). These observations do not support that fatiguing exercise is inevitably associated with alterations in SR Ca2+ uptake or, with the present result in mind, that slowing of relaxation is coupled to diminished SR Ca2+ uptake. Of note is that these measurements of Ca2+ uptake were executed in vitro under optimal conditions, and hence they would not reflect any in vivo alterations in homogenates, which could influence the SR Ca2+ uptake.
In conclusion, the present data demonstrate that 4 min of high-intensity exercise results in a depression of the SR Ca2+ release rate, indicating that the release mechanism of skeletal muscle SR is sensitive to repetitive in vitro muscle contraction. The decreased SR Ca2+ release rate persisted over several hours, despite a full recovery of muscle metabolites (ATP, PCr, lactate, and glycogen) after 30-60 min. The results showed that, after the SR is removed from the fatigued intracellular environment, altered SR function persists. Furthermore, the depression of SR Ca2+ release rate during the slow phase of recovery (1-3 h) showed a close correlation with the +dF/dt, indicating some degree of causality in the fatigued muscle. Despite a slowing of the relaxation rate, we did not find any significant alterations in the SR Ca2+ uptake function, measured as homogenate SR Ca2+ uptake rate and Ca2+-ATPase capacity.
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ACKNOWLEDGEMENTS |
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We thank Preben K. Pedersen for contributing criticism of the manuscript.
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FOOTNOTES |
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This study was supported by grants from Team Denmark, the Danish Sports Research Council, and the Danish Medical Research Council.
Address for reprint requests and other correspondence: N. Ørtenblad, Inst. of Sports Science and Clinical Biomechanics, Univ. of Southern Denmark, Odense Univ., Campusvej 55, 5230 Odense M, Denmark (E-mail: niels.ortenblad{at}agrsci.dk).
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.
Received 9 February 1999; accepted in final form 28 February 2000.
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REFERENCES |
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TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
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) and
stimulated (
) muscles, before and in the recovery phase after
fatiguing stimulations. Maximum (max) tetanic force (Fmax),
rate of force development (+dF/dt), rate of relaxation
(
) and 0.5 h (
) of
recovery are not included in the regression (