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Am J Physiol Regul Integr Comp Physiol 279: R152-R160, 2000;
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Vol. 279, Issue 1, R152-R160, July 2000

Enhanced sarcoplasmic reticulum Ca2+ release following intermittent sprint training

Niels Ørtenblad1, Per K. Lunde2, Klaus Levin3, Jesper L. Andersen4, and Preben K. Pedersen1

1 Institute of Sports Science and Clinical Biomechanics, University of Southern Denmark, Odense University, 5230 Odense M; 3 Institute of Clinical Research, Odense University Hospital, 5230 Odense M; 4 Copenhagen Muscle Research Centre, Rigshospitalet, DK-2200 Copenhagen N, Denmark; and 2 Institute for Experimental Medical Research, Ullevaal Hospital, 0407 Oslo, Norway


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

To evaluate the effect of intermittent sprint training on sarcoplasmic reticulum (SR) function, nine young men performed a 5 wk high-intensity intermittent bicycle training, and six served as controls. SR function was evaluated from resting vastus lateralis muscle biopsies, before and after the training period. Intermittent sprint performance (ten 8-s all-out periods alternating with 32-s recovery) was enhanced 12% (P < 0.01) after training. The 5-wk sprint training induced a significantly higher (P < 0.05) peak rate of AgNO3-stimulated Ca2+ release from 709 (range 560-877; before) to 774 (596-977) arbitrary units Ca2+ · g protein-1 · min-1 (after). The relative SR density of functional ryanodine receptors (RyR) remained unchanged after training; there was, however, a 48% (P < 0.05) increase in total number of RyR. No significant differences in Ca2+ uptake rate and Ca2+-ATPase capacity were observed following the training, despite that the relative density of Ca2+-ATPase isoforms SERCA1 and SERCA2 had increased 41% and 55%, respectively (P < 0.05). These data suggest that high-intensity training induces an enhanced peak SR Ca2+ release, due to an enhanced total volume of SR, whereas SR Ca2+ sequestration function is not altered.

calcium; fatigue; ryanodine receptors; calcium-activated adenosinetriphosphatase; myosin heavy chain distribution


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

REPEATED INTENSE SKELETAL muscle contraction leads to a progressive loss of force generating capacity, i.e., fatigue. The results published to date have consistently implicated low intracellular Ca2+ concentration as a part of the observed fatigue, from both single-fiber preparations (2, 34) and whole body exercise (5, 7). The intracellular free Ca2+ concentration is regulated primarily by the sarcoplasmic reticulum (SR). Skeletal muscle depolarization causes dihydropyridine receptors (DHPR) to trigger the opening of the ryanodine receptors (RyR). The opening of RyR causes release of Ca2+ from SR and subsequent muscle contraction, whereas active reuptake of Ca2+ by the SR Ca2+-ATPase results in muscle relaxation. Given the role of the SR in controlling cytosolic free Ca2+ levels, this organelle is an important factor in the muscle contraction, and impaired SR Ca2+ handling properties potentially could limit the ability to sustain a desired level of force output. The principle of symmorphosis has proposed that the design of all components comprising a system (i.e., fatigue) is matched quantitatively to functional demand, i.e., "enough but not too much" (30). Since impaired SR Ca2+ handling has been linked with fatigue, it might be anticipated that training would reduce the SR impairment following exercise. However, little is known about the training effects on skeletal muscle SR Ca2+ handling properties, i.e., Ca2+ uptake and release and Ca2+-ATPase capacity. In rat fast-twitch muscle, endurance training caused a reduced SR Ca2+-ATPase content, associated with fast-to-slow fiber transition (12). In humans, endurance training does not seem to alter the Ca2+-ATPase concentration or fiber type distribution (20), and the Ca2+-ATPase capacity has been reported to remain unchanged (11). In a cross-sectional study, strength athletes and untrained subjects showed a similar depression of the peak Ca2+ uptake and release rates following repeated maximal contractions, despite that the strength athletes performed more work during the exercise. The present study was undertaken to investigate the role of 5-wk sprint training on intermittent exercise performance, SR Ca2+ sequestration, and release function and SR ryanodine binding.


    METHODS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Subjects

Nine male physical education students, age 24 ± 3 yr (means ± SD), weight 75.5 ± 6.0 kg (and unaffected by training), volunteered for the training study, and six physical education students (age 24 ± 2 yr, weight 73.5 ± 3.8 kg) served as controls. All subjects were fully informed of the potential risks and discomforts associated with the experiment and that they could withdraw from the project at any time, before giving their written informed consent to participate. The study was approved by the local Ethics Committee (file no. 95/4).

Experimental Design

Training was performed on mechanically braked cycle-ergometers. Twenty bouts of 10-s all-out sprints alternating with 50-s rest were repeated three times per week for 5 wk. The load was set to 8% of body weight for the first 3 wk and 8.5% of body wt for the last 2 wk. Every training session was supervised by the investigators, who gave verbal encouragement to the subjects to perform maximally during each sprint. Both the training and the control group were tested at the beginning and end of the training period to determine intermittent sprint performance and maximal oxygen consumption (VO2 max). One week before the first test, the subjects underwent a preliminary test in which they were familiarized with the laboratory equipment and test procedure. Intermittent sprint performance was tested on a modified Monark cycle ergometer. After a 5-min warmup, the subjects performed 10 bouts of 8-s all-out cycling followed by 32-s rest. The subjects were verbally encouraged during the test to mobilize maximal effort. Work load (braking force) was 10% of pretraining body weight; hence, performance depends on revolution frequency, which was kept as high as possible. Power output was calculated every second by a high-resolution computerized system triggered by the passage of each individual tooth on the crank cogwheel through a light diode. With a pedal speed of 120 rpm and a load set to 10% body wt, the power output for an 80-kg person would be 12 W/kg body wt. VO2 max was measured on a separate day using the Douglas bag technique. O2 and CO2 contents were determined with a paramagnetic oxygen analyzer (Servomex model OA 184) and an infrared CO2 analyzer (Beckman model LB2). Expired gas volume was measured with a 130-liter Tissot spirometer. The subjects performed 4 min of light (140 W) and 4 min of heavy (210 W) warmup, followed by a progressive maximal exercise regimen that would lead to exhaustion within 4-8 min, i.e., VO2 max.

Muscle Biopsy Preparation and Analytical Procedures

Muscle biopsy. A resting muscle biopsy was obtained before and 2-4 days after completion of the final training session. After local anesthesia of the skin, subcutaneous tissue, and muscle fascia with 2% lidocaine, a biopsy from the vastus lateralis portion of m. quadriceps femoris was obtained 15 cm above the knee joint, using a Bergström needle with suction (3). This muscle was preferred because it is highly active during cycle exercise (15). The biopsies were taken at random order from the two legs before and after exercise. Each muscle biopsy, yielding about 200 mg wet weight of tissue, was blotted on filter paper, devoided of visible fat and connective tissue and divided into four parts. The first (10-20 mg) was mounted in an embedding medium (OCT compound), rapidly frozen in isopentane precooled by liquid nitrogen, and stored at -80°C for later analysis of myosin heavy chain (MHC) composition. Two specimens were immediately frozen in liquid N2 and stored at -80°C until either freeze dried for analysis of muscle metabolites (20-30 mg) or cut into two parts for the analysis of ryanodine binding and protein immunoblot (~20 and 50 mg, respectively). The rest of the muscle sample (~100 mg) was homogenized in ~10 volumes (wt/vol) ice-cold homogenizing buffer (300 mM sucrose, 1 mM EDTA, 10 mM NaN3, 40 mM Tris-base, 40 mM L-histidine, pH 7.8) (24). Homogenization was done with an Omni model 2000 homogenizer with a 5-mm generator (20,000 rpm), in three 15-s bursts separated by 15-s pause between each burst. The muscle was kept ice cold during the whole procedure. The obtained homogenate was divided into four portions, frozen immediately in liquid N2, and stored at -80°C until analyzed. All the assays were carried out within 5 mo from the last sample collection. Comparisons of SR function measurements on fresh vs. frozen homogenate showed no difference between the two. Protein content in whole homogenates was assessed in triplicate samples using a standard kit (Pierce BCA protein reagent no. 23225).

SR Ca2+-ATPase capacity. SR Ca2+-ATPase capacity was measured by an NADH-linked spectrophotometric technique according to Simonides and van Hardeveld (28), which monitors the rate of ATP hydrolysis in whole homogenates. The method was modified by measuring background ATPase activity in the absence of Ca2+ instead of inhibiting the Ca2+-ATPase by millimolar concentrations of Ca2+. Total ATPase activity was measured in the presence of 25 µM Ca2+, background ATPase was measured in the presence of 1 mM EGTA, and the linear expenditure of NADH was followed for 3 min. SR Ca2+-ATPase capacity was taken to be the difference between the total and the background ATPase activities, since sarcolemma and transverse tubule Ca2+-ATPase capacity only are about 0.2% of SR Ca2+-ATPase capacity. All measurements were carried out with a sample concentration of ~3 mg tissue/ml of buffer, in the presence of 5 µM ionophore A23187 and with high ionic strength (mM) to block the myofibrillar ATPase activity. The assay specificity was tested using the Ca2+-ATPase blocker cyclopiazonic acid (CPA; Sigma catalog number C-1530), which inhibited almost completely the Ca2+-activated ATPase activity. Analyses were performed at 37°C, in a thermostated cuvette holder using a Shimadzu model UV160 spectrophotometer (Teck Science, Mississauga, Ontario, Canada). Activities are expressed as nanomoles per minute per milligram whole homogenate protein.

Ca2+ uptake and release rates. SR Ca2+ uptake and release rates were analyzed using the indo-1 dye as described by Ruell et al. (27). 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 mM oxalate, 11 mM NaN3, 5.5 µM N,N,N',N'-tetrakis(2-pyridylmethyl)ethylenediamine (TPEN), 20 µM CaCl2, and 2 mM MgCl2 (pH 7.0). TPEN, which binds heavy metals without disturbing Ca2+ concentrations, was added to prevent heavy metal ion perturbation of the indo-1 measurements. Oxalate facilitated the SR Ca2+ accumulation, and the addition of NaN3 blocked mitochondrial Ca2+ sequestering activity. ATP and indo-1 were added to a final concentration of 2 mM and 1 µM, respectively, before reaction was initiated by adding ~5 mg tissue to 2 ml of assay buffer (Fig. 1). The subsequent ratiometric data were collected every 0.5 s and converted to free Ca2+ concentrations according to the equation [Ca2+] = Kd · [(R - Rmin)/(Rmax - R)] · (Sf2/Sb2) (13), where R is the ratio value, Rmin is the limiting ratio value when all the indicator is in the Ca2+ free form, Rmax is the limiting ratio value when all the indicator is saturated with Ca2+, and the factor Sf2/Sb2 is the fluorescence intensity measured at 470 nm when all the indicator is free or saturated, respectively. The dissociation constant of indo-1 and Ca2+ (Kd; measured to 142 nM) was measured using a standard Ca2+ calibration buffer kit (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 AgNO3-induced Ca2+ release. The free Ca2+ concentration of the buffer [Ca2+]free was above 1,000 nM before sample addition but decreased to ~700 nM immediately after tissue injection, because of the homogenate EDTA and protein binding of Ca2+ (27). The oxalate-supported Ca2+ uptake was followed for 3 min, which was the point where the Ca2+ uptake plateaued. CPA was added to a final concentration of 40 µM and incubated for 30 s to block the SR Ca2+-ATPase. Addition of AgNO3 (140 µM) induces a rapid spike of Ca2+ release, evoked by thiol oxidation on the SR Ca2+ release channel, followed by a long-term and slow release from the vesicles. Finally Rmin and Rmax were determined in each run, to calibrate the fluorescence signal. Average rates of uptake were determined in 100 nM intervals, between 100 and 700 nM free Ca2+. Peak AgNO3-stimulated SR Ca2+ release rate was determined using the peak first derivative of the calcium concentration ([Ca2+]) vs. time curve (Fig. 1). Since 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+] where the measurements are made. The values obtained for SR Ca2+ uptake and release rates are relative and therefore expressed as arbitrary units of Ca2+ per minute per gram 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 nanomoles free Ca2+ per minute per gram protein. All measurements were performed in duplicate.


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  		Fig. 1.   A representative record of the Ca2+ uptake and release measurements, excluding the calibration procedure. Ratiometric data were used to calculate calcium concentration ([Ca2+]) vs. time. Uptake was initiated by the addition of ~5 mg muscle tissue to the buffer (20 s), and the subsequent Ca2+ uptake was followed for about 3 min until it plateaued. Addition of 40 µM cyclopiazonic acid (CPA) (195 s) blocked the sarcoplasmic reticulum (SR) Ca2+-ATPase, before release was initiated by 140 µM AgNo3 (220 s). Peak Ca2+ release rate was determined using the peak first derivative of the [Ca2+] vs. time curve.

Ryanodine binding. The content of RyR was determined as described by Lunde and Sejersted (19). In short, 40-100 mg biopsy material was homogenized in a HEPES/KCl buffer, pH 7.4, containing protease inhibitors and 1% of the detergent Tergitol. SR membranes were isolated by differential centrifugation. The subsequent SR membrane fraction contains nearly 90% of the [3H]ryanodine binding sites of the tissue, 81% of the muscle Ca2+-ATPases, and only 14% of the whole muscle protein content as previously reported (19). The SR membranes were incubated in a Tris · HCl buffer, pH 8.0, containing 3 mM disodium ATP, 0.1 mM CaCl2, protease inhibitors, and 20 nM [3H]ryanodine for 180 min at 37°C. The Kd value for [3H]ryanodine binding to human gluteus muscle is 8.1 nmol/l, and the Bmax is 1.2 pmol/mg SR protein (19). Unspecific binding of [3H]ryanodine was quantified by incubating the SR vesicles in the same buffer containing 20 µM unlabeled ryanodine. Bound and unbound [3H]ryanodine were separated by filtration through a combination of glass-fiber filter and cellulose nitrate/cellulose acetate filter. The filters were washed with 10% ethanol, dried, and counted in a liquid scintillation counter (model LS5000 CE; Beckman, Palo Alto, CA) after addition of scintillation cocktail.

Protein immunoblot analysis. Muscle membrane proteins were isolated as previously described by Ploug et al. (25). In short, frozen biopsy material (10-20 mg) was homogenized in a buffer containing 210 mM sucrose, 30 mM HEPES (pH 7.4), 2 mM EGTA, 40 mM NaCl, and 2 mM phenylmethylsulfonyl fluoride with a Polytron homogenizer at full speed for two 15-s periods. The homogenate was mixed with a KCl/sodium pyrophosphate buffer (final concentration, 500 mM/25 mM) and incubated on ice for 15 min. All membranes were recovered in a pellet after centrifugation at 200,000 g for 75 min. Membrane protein concentrations were determined by the bicinchoninic acid assay (Pierce 23235) using bovine serum albumin as standard. For quantitation of the different proteins 0.5, 1, and 2 µg of the membrane preparation were loaded onto a polyvinylidene difluoride (PVDF) filter membrane by the use of a filtration manifold (Minifold II; Schleicher & Schuell, Dassel, Germany). PVDF membranes were blocked by incubating for 1 h at room temperature or overnight at 4°C in 10% nonfat dry milk in Tris-buffered saline, pH 7.5, with 0.1% Tween-20 (TBS-T). The PVDF membranes were then incubated for 1 h at room temperature with the primary antibodies [anti-SERCA1 (MA3-912), anti-SERCA2 (MA3-919) and anti-RyR (MA3-925) all from Affinity Bioreagents] and diluted in blocking solution at following dilutions: SERCA1, 1:2,500; SERCA2, 1:1,000; RyR, 1:5,000. After five washes with TBS-T, the PVDF membranes were incubated for 1 h with anti-mouse immunoglobulin G conjugated to horseradish peroxidase (NA-931; Amersham, Oakville, Ontario, Canada) in TBS-T. The PVDF membranes were washed five times with TBS-T, and the immunoreactivity was detected by the enhanced chemiluminescence method (RPN3103H; Amersham). Signal intensity of the slots on the film was quantified with the ImageQuant software (Molecular Dynamics). The results of the blot were calculated by first normalizing the data for each group of the pretreatment value for each protein concentration (i.e., 0.5, 1, and 2 µg) and then calculating the average for the three measurements.

Muscle metabolites. Metabolites were extracted from the freeze-dried muscle samples by perchloric acid treatment and analyzed for the total content of glycogen, creatine phosphate (PCr), and adenosine triphosphate (ATP) by fluorometric assays according to Lowry and Passonneau (18).

MHC composition. MHC analysis was performed using SDS-PAGE (1). Ten serial cross sections (20 µm thick) from each biopsy were placed in 100-200 µl of lysing buffer and heated for 10 min at 60°C (9). One to three microliters of the myosin-containing samples was loaded on a SDS-PAGE gel containing 6% polyacrylamide and 37.5% glycerol. Gels were run overnight at 70 V, followed by 2-4 h of 200 V. Subsequently the gels were silver stained, and the relative proportions of MHC-I, MHC-IIA, and MHC-IIX isoforms were determined densitometrically (Cream 1-D; Kem-En-Tec, Copenhagen, Denmark).

Statistics

Values in the text, Tables 1-3, and Figs. 1-6 are presented as means ± SE or as means and range. Statistical comparisons were made for mean differences using Student's t-test for paired (within group) and unpaired (between groups) observations. Nonparametric statistics (Wilcoxon signed rank test) were used to compare the Ca2+ release rates before and after training, because of the nonparametric distribution of the data (see Fig. 3B). The performance data were analyzed statistically with a two-way repeated measure analysis of variance (ANOVA). Significance was established at the 0.05 level.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Performance

Total amount of work during the 10-repetition intermittent exercise increased by 12% (P < 0.01) after training, from an average of 66.4 ± 2.2 kJ (before) to 74.6 ± 3.2 kJ (after), whereas controls remained unchanged at 66.7 ± 2.0 kJ (before) vs. 67.0 ± 2.4 kJ (after). Corresponding average power output per kilogram body weight in the training group was 11.0 ± 0.2 (before) vs. 12.3 ± 0.2 (after) W/kg body wt (P < 0.01), and controls remained unchanged [11.4 ± 0.3 (before) vs. 11.6 ± 0.4 (after) W/kg body wt]. The average power output, generated before training, decreased 20.5% from bout 1 to bout 10, from 12.8 to 10.2 W/kg (Fig. 2A). However, the experimental group increased significantly (P < 0.001) both the level and the relative maintenance in average power output with training, and controls remained unchanged (Fig. 2A). Splitting up the average power output in seconds, i.e., power through seconds 1-8, as mean for all 10 bouts (Fig. 2B) shows that the performance was significantly higher (P < 0.05) in all seconds after training, but the pattern of power output was equal within each bout. This means that training induced an increase in the rate of power output during the first second of exercise, and the trained subjects developed power output in the same pattern throughout the exercise period before and after training. All in all, the enhanced interval exercise performance following training was achieved through increased power output throughout seconds 1-8, and the subjects fatigued to a lesser extent from bout 1 to bout 10. The mean pedal frequencies during the first and last 8-s sprint, before training, were 128 and 102 rpm, respectively.


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  		Fig. 2.   Average power output before (open circle ) and after () the 5-wk training period, for the training group and controls. A: average power for interval 1-10. B: average power for each second. Values are means ± SE, expressed as watts per kilogram body weight. #Significantly different from before training (time and treatment effect; P < 0.001).

VO2 max values of the subjects were unaffected by training [61.3 ± 1.7 vs. 61.0 ± 1.4 ml · min-1 · kg-1 (mean ± SD) and 64.0 ± 0.5 vs. 64.3 ± 1.6 ml · min-1 · kg-1, for trained subjects and controls, respectively].

SR Ca2+ Release Rate

The 5-wk sprint training induced a 9% (P < 0.05) higher peak rate of AgNO3-stimulated SR Ca2+ release (Fig. 3) from a mean value (ranges are in parentheses) of 709 (560-877) (before) to 774 (596-977) arbitrary units Ca2+ · g protein-1 · min-1 (after). The Ca2+ release rate in the control group remained unchanged [702 (607-780) vs. 701 (590-793) arbitrary units Ca2+ · g protein-1 · min-1]. There were no differences between training vs. control and before vs. after training in the [Ca2+]free levels at which the release rates were performed (averaging 110 ± 12 nmol), and the rates can therefore be compared.


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  		Fig. 3.   A: peak AgNO3-induced rate of SR Ca2+ release before and after 5 wk of high-intensity training. The [Ca2+]free at which the release rates were performed averaged 110 ± 12 nM. The box plots show the median as a line, first and third quartiles as boxes, and minimum and maximum values as extensions. B: the individual changes, posttraining minus pretraining. Values are expressed as arbitrary units Ca2+ · g protein-1 · min-1, in 9 and 6 subjects for trained group and control group, respectively. *P < 0.01.

SR Ca2+-ATPase Capacity and Ca2+ Uptake Rate

The SR Ca2+-ATPase capacities in whole muscle homogenates were not affected by 5-wk intermittent sprint training (95.3 ± 5.4 before vs. 100.5 ± 4.0 nmol · mg protein-1 · min-1, after) (Table 1). Additionally, both total and background ATPase capacities were unaffected by training. The background ATPase was on average 28% of total ATPase capacity. There was no difference in total, background, and SR Ca2+-ATPase capacity between the training group and controls. As for the SR Ca2+-ATPase capacity, there were no differences in SR Ca2+ uptake rates before and after training, when measured at the same [Ca2+]free, in the physiological levels of 100-700 nM [Ca2+]free (Fig. 4). The Ca2+ uptake averaged 3,133 ± 209 and 370 ± 16 arbitrary units Ca2+ · g protein-1 · min-1 between 600-700 and 100-200 nmol [Ca2+]free, respectively.

                              
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  		Table 1.   Homogenate Ca2+-ATPase capacities for trained and control



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  		Fig. 4.   Average rate of Ca2+ uptake before (open bars) and after (solid bars) training for the training group. Measurements were made in 100 nM intervals at free Ca2+ concentrations between 100 and 700 nM. All measurements were performed in duplicate. Activities are expressed as arbitrary units Ca2+ · g protein-1 · min-1, in means ± SE. Note that the arbitrary Ca2+ units are not the same at different values of [Ca2+]free.

Ryanodine Binding

Prior to training, the total binding of [3H]ryanodine to purified SR vesicles was 920 ± 90 pmol/mg SR protein, and the SR ryanodine binding did not change following training (Fig. 5). There were no significant differences between the training and the control group, before and after the training period.


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  		Fig. 5.   [3H]ryanodine binding of purified SR vesicles. Values are means ± SE, expressed as pmol [3H]ryanodine/mg SR protein.

SERCA1, SERCA2, and RyR Immunoblots

The Western blot values are expressed as percentage of value before training, in means of 0.5, 1.0, and 2.0 mg protein on the blots. Western blots of the Ca2+-ATPase isoforms SERCA1 and SERCA2 showed that the relative gel density was increased 41% and 55%, respectively (P < 0.05) (Fig. 6). Similarly, there was a pronounced 48% increase in the membrane fraction RyR following training (P < 0.05) (Fig. 6). There were no significant changes in the amount of protein in the different groups before and after training, neither for SERCA isoforms nor RyR. There were no differences in the total membrane protein concentration per muscle wet weight in the four sample groups, i.e., training and control groups before and after the training period.


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  		Fig. 6.   Relative amount of Ca2+-ATPase isoforms (SERCA1 and SERCA2) and RyR channels, before and after training. The amounts (gel density) are in relative values per milligram membrane protein and are expressed as percentage of pretraining value. The value for each subject is the average immunoblot of 0.5, 1.0, and 2.0 µg protein added. aSignificantly different from value before training.

MHC Composition

The vastus lateralis distribution of MHC isoforms, revealed an almost even distribution of MHC I and MHC II, with no difference between groups (Table 2). For the entire group of subjects before training, the abundance of MHC I varied from 37 to 71%. There were no changes in the distribution of the different isoforms before and after the training period, in neither trained nor control subjects.

                              
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  		Table 2.   Myosin heavy chain isoforms

Muscle Metabolites

The muscle metabolite concentrations before and after training are shown in Table 3. There were no differences in the ATP, PCr, and glycogen levels between the two groups, and no changes in muscle metabolite levels were found following training.

                              
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  		Table 3.   Muscle metabolite concentrations in resting biopsies from vastus lateralis before and after 5 wk sprint training


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Ca2+ Release Results

This study shows a significant increase in the peak rate of AgNO3-stimulated SR Ca2+ release following 5-wk high-intensity intermittent training. Alterations in Ca2+ release in vivo could be caused by a variety of factors, including failure in the conduction of the action potential by the T tubules, impaired coupling between voltage sensors and the SR Ca2+ channels (33), and metabolic or ionic changes (8, 17). However, since the SR function measurements are done in vitro under constant conditions, the measured changes in SR function cannot be due to metabolic changes associated with work. Hypotheses directed at explaining the mechanism(s) of the observed enhanced SR Ca2+ release rate following training could be one or a combination of the following four possibilities: 1) altered fiber type distribution (i.e., type I right-arrow II), 2) increased SR release channel density, 3) increased SR content per se in the existing fiber types, and/or 4) structural changes to the involved proteins (i.e., increased channel opening-time/conductance). To differentiate between these possibilities, we measured the MHC distribution, the relative number of functional RyR receptors before and after training (Fig. 6), as well as the SR RyR density (Fig. 5). First, altered fiber type distribution, measured as the contractile protein distribution, i.e., MHC or ATPase isoform, would have a marked influence on the muscle SR Ca2+ regulation. Although the arrangement and architecture of the T-tubular network and terminal cisternae of the SR are similar in slow-twitch (MHC I) and fast-twitch (MHC IIA and IIB) fibers, important quantitative differences exist. Fast-twitch fibers roughly contain twice the volume and area of terminal cisternae and about four to six times more Ca2+-ATPase content and activity than slow-twitch fibers (26). Furthermore, the Ca2+ release rate is about four to six times higher in fast-twitch compared with slow-twitch fibers. Hence, if high-intensity training induces transformation of slow-twitch right-arrow fast-twitch fibers, then this would lead to an enhanced Ca2+ release rate. However, since the MHC distribution is a product of the number of fibers and the fiber volume, an unaltered MHC distribution means that the relatively amount of the two main fiber types remained unchanged. Thus an enhanced SR content due to an increase in the relative amount of MHC II fibers cannot be the explanation for the observed enhanced Ca2+ release rate. Second, an increased SR release channel density of the SR (i.e., more release channels per surface SR) would lead to an increased Ca2+ release rate per unit SR or muscle, assuming an unaltered SR content. However, measurements of the ryanodine channel density on purified SR vesicles (i.e., pmol [3H]RyR bound per mg SR protein), showed that the density was unaltered by the 5-wk training (Fig. 5). Third, the training could increase the SR content per muscle fiber, without alterations in the contractile apparatus (i.e., ATPase isoforms and/or MHC isoforms). Immunoblot gel density of RyR channels showed a 48% relative increase in the total number of RyRs per unit muscle membrane protein (Fig. 6). The densities are relative units of the total muscle membrane content, i.e., mitochondrion, SR, and sarcolemma membranes. This means that the enhanced immunoblot density could be due to either an increased amount of SR membrane and/or that the number of RyR relative to SR increased. However, these results of an increased total amount of channels, together with unaltered RyR density, lead to the conclusion that the total amount of SR is enhanced. Hence, it seems that the muscle SR content is enhanced, without change in the MHC distribution. The enhanced amount of SR could be the explanation, fully or in part, for the observed training-induced increase in the Ca2+ release rate. Finally, alterations in the RyR channel gating behavior, which could be due to increased channel opening time and/or conductance, could explain an increased SR Ca2+ release rate. The relative importance of this notion can only be speculated. Lactate has been reported to reduce the Ca2+ release channel opening fluctuations on the single-channel level (8); however, little is known about the direct effect on single-channel fluctuations. Favero et al. (7) have shown that diminished AgNO3-stimulated Ca2+ release by SR of fatigued muscle is due to a diminished number of functional channels. An enhanced AgNO3-stimulated SR vesicle Ca2+ release following a training period would likely be expected to increase the number of functional channels. Hence, the relative AgNO3-stimulated Ca2+ release, as we measure, would increase.

The indications of an increase in the muscle SR content, without significant alterations in MHC isoform distribution, following the 5-wk training period, is supported by denervation experiments and studies of continuous low-frequency stimulation. Midrio et al. (22) reported that 2 days of denervation decreased both contraction time and tension of the isometric twitch, whereas an increase was observed in 7-day denervated muscles. The rate of SR Ca2+ uptake and the calculated amount of Ca2+ release showed a close connection with the mechanical data. However, fiber type composition, as well as changes in the Ca2+ sensitivity of the muscle fibers, did not correlate with mechanical changes. It was therefore concluded that the SR plays a prominent role in the early changes of contraction time and tension following denervation. Twenty days of 10-Hz continuous low-frequency stimulation resulted in a steep parallel decline of DHPR, RyR, and triadin (a triad junction marker), as well as Ca2+-ATPase activity, whereas the fast-to-slow transition at the myosin level had not yet occurred at that time (16). Furthermore, significant modifications of SR properties in rat skeletal muscle have been described to occur as early as 1 wk after denervation (21, 22), whereas 6 wk of sprint training did not elicit significant variation in MHC isoform or fiber-type distribution (14). These results, together with the present data, suggest that there is a different time course in the development of the diverse characteristics of the various fiber types, i.e., SR Ca2+ regulation properties and the myosin ATPase and heavy chain isoforms. With this in mind, individual fibers are capable of having SR Ca2+ regulating characteristics of MHC II fibers while also having MHC I properties of the contractile machinery.

Assuming a five times higher RyR content and Ca2+ release rate in fast vs. slow fiber types and an initial 50/50 fiber type distribution, the 5-wk training-induced alteration in fiber type distribution can be calculated. A 9% increase in the Ca2+ release rate indicates a fiber type transformation to 57% fast and 43% slow, regarding SR Ca2+ release rate property. However, a 40% increase in the total SR content, as observed, would denote an 80% fast and 20% slow type, following training. Discrepancies in the calculated distributions could be partly due to different sensitivities in the used analyses and partly due to different time course in the training-induced switching of the SR and RyR content.

Ca2+ Uptake and Ca2+-ATPase Results

Despite the enhanced sprint performance, no differences in SR Ca2+-ATPase capacity and Ca2+ uptake rate at the same [Ca2+]free, between 100 and 700 nM [Ca2+], were observed. Unchanged amounts of Ca2+-ATPases in human vastus lateralis muscle have been reported after intensified endurance training (10, 20); also, 4-12 wk strength training did not alter Ca2+-ATPase capacity (11). The present results confirm these findings and extend them to sprint training. Based on these studies, it appears that both the number of Ca2+-ATPases and the maximal ATP hydrolysis rate are not trainable. Since the SR function measurements are done in vitro under constant conditions, potential changes in SR function may be due to alterations in the ATP binding site or hydrolysis rate and/or alterations in the number of active SR Ca2+-ATPase enzymes. However, we are not able to conclude anything about the SR Ca2+ sequestration function in situ, where substrate availability and the lactate, Pi, ADP, Mg2+, and pH levels have been shown to alter the uptake rate (29, 31, 32).

SERCA1 and SERCA2

Western blots of the Ca2+-ATPase isoforms SERCA1 and SERCA2, showed that the relative gel density, which equals the total amount of SERCA isoforms, was increased 41% and 55%, respectively (P < 0.05). SERCA1 is the major fast-twitch Ca2+-ATPase isoform, constituting up to 90% of the membrane protein, whereas the SERCA2 isoform is found in slow-twitch muscles. A 41% increase in the fast twitch isoform supports very well the finding of a pronounced (48%) increase in the ryanodine binding following training, indicating a slow-to-fast twitch transformation. The finding of an increased amount of Ca2+-ATPases without an increment in Ca2+-ATPase capacity or Ca2+ uptake rate seems enigmatic. However, Hicks et al. (16) demonstrated a diverse time course of the protein level and the activity of the Ca2+-ATPase, verified by a decrease in activity, not the expression, of the ATPase following low-frequency stimulation. This observation might be the explanation for the diversity of protein expression and SR Ca2+ uptake properties observed in the present study.

Intermittent Sprint Performance and Metabolites

Intermittent sprint training enhanced the mean average power output by 12%, as well, and attenuated the development of fatigue from bout 1 to 10, during the intermittent sprint performance test. This demonstrates that the enhanced sprint performance was more distinct in the last part of the 10 sets of 8-s bouts, which reflects the training procedure, consisting of sprint endurance (i.e., 20 times, 10-s all-out), more than maximal power output. Similar to the findings of the present study, Boobis et al. (4) reported an 8% increase in power output, following 8-wk cycle sprint training. Resting ATP concentrations were unchanged, whereas PCr decreased and glycogen increased. In the present training study, the trained subjects did not significantly alter the muscle metabolite concentration; however, there was a tendency to an elevated glycogen level. This is in agreement with Nevill et al. (23), who did not find changes in resting muscle metabolites, after 8-wk sprint run training. Although it is well accepted that the muscle glycogen level is important for the endurance performance, the importance during intermittent high-intensity exercise and SR Ca2+ regulation is less clear. Reduced glycogen availability has been reported to reduce SR Ca2+ release (6). Furthermore, functional coupling of ATP generated by SR-associated glycolytic enzymes may play an important role in cellular Ca2+ homeostasis by driving the SR Ca2+ pump (35).

Conclusions

In summary, a significant new finding in this study is that 5 wk of high-intensity training induced an increase in the peak AgNO3-stimulated SR Ca2+ release rate, together with an enhanced rate of power output and average power output during intermittent cycle sprint. Furthermore, the data indicate that the 5-wk sprint training-induced increase in SR Ca2+ release rate was due to an enhanced SR content without changing the MHC distribution. This shows that there is a different time course in the development of SR and MHC isoform distribution. The present study demonstrates that high-intensity intermittent training does not change SR Ca2+ sequestration function, measured as SR maximal Ca2+-ATPase capacity and SR Ca2+ uptake rate, despite a significant increase in intermittent sprint performance.

Perspectives

Little is known about the training effects on skeletal muscle SR function, and to our knowledge no studies have evaluated the training effect on the SR Ca2+ release. The present study demonstrates that high-intensity training improved muscle performance and enhanced the peak AgNO3-stimulated SR Ca2+ release rate due to an increased SR content, without changing the MHC distribution. Both denervation and low-frequency stimulation studies have shown that adaptations in skeletal muscle SR function correlate with changes in the muscle mechanical properties, e.g., rate of force development (16, 21, 22). Viewed in the context of the present data, these observations suggest that MHC composition is not the sole determinant of skeletal muscle mechanical properties, but that muscle SR content, and with that Ca2+ release rate, may also play a role. Thus the common perception that the contractile properties of a given muscle primarily relate to the isoform of the contractile filaments may have to be broadened with respect to a potential significance of the skeletal muscle SR function. Changes in the SR composition seem to occur rapidly (within days) and precede changes in the MHC or ATPase isoforms of the contractile filaments (14, 16, 22). Hence, the former discussion may not contradict that the contractile properties relate to the isoform of the contractile filament since muscle SR alterations occur prior to the transformation of the contractile proteins.


    ACKNOWLEDGEMENTS

We thank J. T. Rasmussen and H. Bak for skilled assistance during the test and training procedure.


    FOOTNOTES

This study was supported by grants from Team Denmark, the Danish Sports Research Council, and the Danish Medical Research Council.

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.

Address for reprint requests and other correspondence: N. Ørtenblad, Institute 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).

Received 17 November 1999; accepted in final form 19 January 2000.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

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