HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS		QUICK SEARCH:   [advanced] Author: Keyword(s): Year:  Vol:  Page:

This Article Abstract Full Text (PDF) Free All Versions of this Article: 103/6/1986    most recent 01440.2006v1 Submit a response Alert me when this article is cited Alert me when eLetters are posted Alert me if a correction is posted Citation Map Services Email this article to a friend Similar articles in this journal Similar articles in Web of Science Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Citing Articles Citing Articles via HighWire Citing Articles via Web of Science (3) Citing Articles via Google Scholar Google Scholar Articles by Duhamel, T. A. Articles by Ouyang, J. Search for Related Content PubMed PubMed Citation Articles by Duhamel, T. A. Articles by Ouyang, J.

Muscle metabolic, SR Ca²⁺-cycling responses to prolonged cycling, with and without glucose supplementation

Department of Kinesiology, University of Waterloo, Waterloo, Ontario, Canada

Submitted 19 December 2006 ; accepted in final form 28 September 2007

	ABSTRACT

TOP ABSTRACT METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
This study investigated the effects of prolonged exercise, with and without glucose supplementation, on metabolism and sarcoplasmic reticulum (SR) Ca²⁺-handling properties in working vastus lateralis muscle. Fifteen untrained volunteers [peak O₂ consumption (O_2peak) = 3.45 ± 0.17 l/min; mean ± SE] cycled at 60% O_2peak on two occasions, during which they were provided with either an artificially sweetened placebo beverage (NG) or a 6% glucose (G) beverage (1.00 g carbohydrate/kg body mass). Beverage supplementation started at 30 min of exercise and continued every 15 min thereafter. SR Ca²⁺ handling, metabolic, and substrate responses were assessed in tissue extracted from the vastus lateralis at rest, after 30 min and 90 min of exercise, and at fatigue in both conditions. Plasma glucose during G was 15–23% higher (P < 0.05) than those observed during NG following 60 min of exercise until fatigue. Cycle time to fatigue was increased (P < 0.05) by 19% during G (137 ± 7 min) compared with NG (115 ± 6 min). Prolonged exercise reduced (P < 0.05) maximal Ca²⁺-ATPase activity (–18.4%), SR Ca²⁺ uptake (–27%), and both Phase 1 (–22.2%) and Phase 2 (–34.2%) Ca²⁺-release rates during NG. The exercise-induced reductions in SR Ca²⁺-cycling properties were not altered during G. The metabolic responses to exercise were all unaltered by glucose supplementation, since no differences in respiratory exchange ratios, carbohydrate and lipid oxidation rates, and muscle metabolite and glycogen contents were observed between NG and G. These results indicate that the maintenance of blood glucose homeostasis by glucose supplementation is without effect in modifying the muscle metabolic, endogenous glycogen, or SR Ca²⁺-handling responses.

Ca²⁺ regulation; glucose supplementation; human skeletal muscle; metabolism

The SR in skeletal muscle is uniquely structured to perform its primary role, namely the ability to rapidly respond to the transmission of neural command signal by rapid and precise changes in the [Ca²⁺]_f transient, necessary to activate the myofibrillar apparatus for the production of a desired force response (51). To regulate [Ca²⁺]_f, Ca²⁺ cycling increases dramatically during contractile activity by increasing both Ca²⁺ release and Ca²⁺ uptake in a coordinated manner (51). As might be expected, both the CRC and the Ca²⁺-ATPase are under complex intrinsic regulatory control (49).

Disturbances in the [Ca²⁺]_f transient, as measured by Ca²⁺-sensitive dyes, are a well-documented effect of repetitive contractile activity (1). The disturbances in the [Ca²⁺]_f transient have been shown to be accompanied by reductions in both Ca²⁺ release (1, 5) and Ca²⁺ uptake (1, 5) in single fibers during induced contractions of the mouse, ostensibly as a result of changes in the intracellular environment mediated by the contractile activity (40). Reductions in Ca²⁺ release (20, 37), Ca²⁺ uptake (5, 37), and maximal Ca²⁺-ATPase activity (V_max) (6, 37, 47) have also been shown to occur in rat muscle with repetitive contractions when measured in homogenates "in vitro" under supposedly optimal conditions. However, not all studies employing rats show disturbances in SR Ca²⁺ cycling in muscle with prolonged, submaximal exercise (21, 66). Of the few studies that have examined the SR responses to a similar exercise challenge in humans, all have found disturbances in SR Ca²⁺ handling, namely V_max, Ca²⁺ uptake, and Ca²⁺ release (4, 14, 15). Based on the limited evidence available, it appears that no changes occur in coupling efficiency, defined as the ratio between Ca²⁺ uptake and V_max, or in the sensitivity of the enzyme to Ca²⁺ (15, 66). However, one study (13) has reported that, late in prolonged exercise, changes in membrane permeability to Ca²⁺ occurs, which conceivably should alter coupling efficiency.

One factor that appears to affect the SR Ca²⁺ cycling function during repetitive contractions is the carbohydrate (CHO) status of the muscle cell. Several studies have linked the depletion of muscle glycogen with reduced [Ca²⁺]_f transients in rat (43), mouse (7), and toad (70), while others (16) have reported that exercise-induced reductions in Ca²⁺-handling properties in human muscle measured in vitro occur earlier during low-glycogen compared with high-glycogen states. In this latter study, no changes were observed in the Ca²⁺ sensitivity of the enzyme, the coupling ratio, or membrane permeability to Ca²⁺ (16).

The most inviting hypothesis to account for the relationship between muscle SR Ca²⁺ cycling and CHO status is based on the substrate and/or energy status of the cell. According to this rationale, contracting muscle with depleted glycogen reserves experience a decreased ability to regenerate ATP. As a consequence, ATP levels decline and metabolic by-products accumulate, both of which may affect Ca²⁺ release (18) and Ca²⁺ uptake (41). Evidence also exists to support the existence of a SR-glycogenolytic complex containing glycogen phosphorylase, glycogen debranching enzyme, many of the enzymes involved in the glycolytic pathway, and Cr phosphokinase (86), the dissociation of which could alter Ca²⁺-cycling behavior (44, 85). The depletion of cellular glycogen stores could effectively dissociate this complex and result in a disturbance in energy homeostasis in regions close to the Ca²⁺-ATPase (12) and CRC (30), or result in structural alterations in the SR, both of which could impact on SR Ca²⁺ cycling (43).

A limitation in our laboratory's previous work designed to investigate the relationship between muscle glycogen and SR Ca²⁺-handling properties during prolonged whole body exercise (16) was the inability to control blood glucose levels and, consequently, the regulatory hormones, insulin (Ins) and the catecholamines, norepinephrine (NE) and epinephrine (Epi). In our laboratory's previous work, alterations in the dietary intake of CHO resulted in differences during exercise in blood glucose concentrations that were also accompanied by differences between conditions in plasma NE and Epi (16) and most probably Ins as well (23). Differences in blood glucose availability could alter SR Ca²⁺-cycling function via improved energy homeostasis and/or protection of muscle glycogen reserves (44, 85), while differences in the hormonal responses could affect intrinsic behavior through second-messenger regulation (49, 84). As an example, Epi could alter the phosphorylation status of the regulatory protein phospholamban (PLN) via cAMP signaling, resulting in alterations in the catalytic activity of the enzyme (49). Interestingly, as with experiments designed to manipulate muscle glycogen levels by exercise and diet (3), oral glucose supplementation during exercise also has an ergogenic effect (11, 31).

Maintenance of blood glucose homeostasis during prolonged exercise could affect SR Ca²⁺-cycling responses, by altering one or more of the factors involved in its regulation. In this regard, the literature is contradictory with regard to the effects of glucose supplementation on muscle metabolism (8, 69, 72), glycogen depletion patterns (11, 55, 72), and blood hormonal responses (10, 24, 31).

The main purpose of this study was to investigate the effects of prolonged exercise and prolonged exercise plus oral glucose supplementation on muscle SR Ca²⁺-cycling function. We have hypothesized that prolonged exercise would result in a progressive reduction in the maximal activity of the Ca²⁺-ATPase, Ca²⁺ uptake, and Ca²⁺ release in active muscle and that the reduction in these properties with exercise would occur in the absence of changes in Ca²⁺ sensitivity, Ca²⁺-transport efficiency, and SR membrane permeability to Ca²⁺. Oral glucose supplementation is proposed to attenuate the exercise-induced changes in SR Ca²⁺-cycling properties by protecting muscle energy homeostasis and glycogen content during exercise.

	METHODS

TOP ABSTRACT METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
Participants.   Fifteen volunteers (14 men and 1 woman) were recruited from the general student population at the University of Waterloo to participate in the study. Volunteers were healthy and recreationally active (assessed by a questionnaire), but not involved in exercise utilizing large-muscle groups on a regular basis (i.e., not more than once per week). The physical characteristics of the participants included the following: age, 19.3 ± 0.4 yr; height, 179 ± 4 cm; and body mass, 78.5 ± 3.7 kg. Peak aerobic power (O_2peak) was 3.45 ± 0.17 l/min. All volunteers were healthy, as determined by questionnaire. Testing was completed during the midfollicular phase of the menstrual cycle for the female participant in this study. This study received approval from the Office of Research Ethics at the University of Waterloo.

Experimental design.   To investigate the effects of oral glucose supplementation on muscle SR Ca²⁺-transport properties, two sessions of prolonged, moderate-intensity exercise were employed. One session served as the control, or no glucose (NG), condition and was used to investigate the effects of exercise in isolation without oral glucose supplementation. The other session served as the glucose supplementation (G) condition and was identical to NG, with the only difference being the ingestion of oral glucose supplements, designed to maintain blood glucose levels during the late stages (i.e., >30 min) of exercise. The order of the two experimental conditions was randomized for 9 of 15 participants. The remaining six participants completed the NG condition before the G condition. The latter sequence allowed for muscle tissue to be sampled at a matched time point in both conditions within this subgroup. The matched time point was selected to correspond to fatigue in the NG condition and was unique to each individual. The time to fatigue in NG was 115 ± 6 min, whereas average time to fatigue in G was 137 ± 7 min. The increase in time to fatigue in G was 19.1% (P < 0.05).

Before the commencement of each test session, participants were asked to complete a 7-day diet journal to allow for the assessment of average daily nutritional intake. Based on the analysis of these 7-day diet journals, no differences were observed in total caloric intake or macronutrient intake during the lead-in period between experimental conditions.

The prolonged cycle task was performed in a neutral environment (20°C; 50% relative humidity) at an intensity that was 60% O_2peak. Exercise was continued until volitional fatigue or when the participant could not maintain a cadence of at least 50 revolutions/min, with verbal encouragement.

Glucose supplements.   The glucose supplement was a 6% solution of glucose, without the addition of any electrolytes. Participants were provided with a drink (volume dependent on body mass; >1.00 g CHO/kg body mass in a 6% solution), starting after 30 min of exercise and continuing every 15 min thereafter. The average total volume ingested at each time point ranged between 100 and 300 ml. A placebo (Sugar Twin; Alberto-Culver Canada, Toronto, ON, Canada) consisting of a 7.5% Sugar Twin [water, sodium cyclamate (10%), benzoic acid, methyl paraben] solution was provided according to the same schedule using similar individual volumes, as received during G. Beverages were served at room temperature (20°C). For participants who completed the NG condition before G, beverage volumes were matched to the volume that was to be consumed during the G condition. Test sessions were separated by at least 4 wk and were conducted in the morning, following an overnight fast.

The total amount of CHO ingested during G was 1.23 ± 0.11 g CHO/kg body mass. The average volume ingested at each time point was 243 ± 17 ml, which amounted to a total volume of 1,564 ± 142 ml for G over the course of the exercise. The volume of fluid ingested at each time point for NG was not different from G.

O_2peak determination and respiratory gas collection.   A progressive exercise test was performed on an electrically braked cycle (Quinton 870), as previously described (35), to measure O_2peak using an open-circuit gas collection system with continuous measurements (36). The gas collection system was calibrated daily, 30 min before all test sessions, using standardized gas samples of known concentrations. All exercise sessions were performed on the same cycle ergometer and using the same respiratory gas collection system. Respiratory gas properties measured during the exercise tests include oxygen uptake (O₂), carbon dioxide production (CO₂), and expiratory ventilation (E). From these data, respiratory exchange ratios (RER = CO₂/O₂) have been calculated. Stoichiometric equations and appropriate caloric equivalents (22, 82) were used to calculate CHO and lipid oxidation rates during the exercise.

Blood sampling.   Blood samples were collected at rest, during exercise (15, 30, 45, 60, and 90 min), and at fatigue (115 ± 6 min) from a catheter inserted in the prewarmed dorsal region of the hand during the NG condition. During G, blood samples were collected at rest, during exercise (15, 30, 45, 60, and 90 min), at a time corresponding to fatigue during the NG condition (115 ± 6 min) and at fatigue (137 ± 7 min).

The remaining aliquots of whole blood, serum, and plasma samples were processed and frozen at –20°C until analyses. Blood glucose and lactate (Lac) were determined fluorometrically, in triplicate, from the plasma aliquots that were deproteinized using perchloric acid and centrifuged (26). On a given day, all samples for a given individual for a given property were analyzed in triplicate.

Tissue sampling.   Four tissue samples for each condition were collected from the vastus lateralis under local anesthesia (2% xylocaine). For NG, tissue samples were collected at rest, after 30 and 90 min of exercise, and at fatigue. During the G condition, tissue samples were collected at rest, after 30 and 90 min of exercise, and at fatigue in 9 of 15 participants. In the remaining six participants, the resting tissue sample in G was omitted and replaced with a tissue sample taken at a time corresponding to fatigue during the NG condition (i.e., matched NG fatigue). The decision to replace the resting tissue sample with a matched placebo fatigue tissue sample was based on observations made in this study (data not shown), in addition to numerous experiments from our laboratory (14–16) that have shown no differences in resting SR Ca²⁺-handling or metabolic properties between conditions when the experimental treatment was preceded by a standardized diet. This approach allowed for direct comparison of tissue samples collected at a time corresponding to the placebo fatigue time point in NG and G. Statistical analyses of all data were adjusted to account for this subgroup of participants, as described in Statistical analyses below.

Two tissue samples were taken from each site at each sampling time. The first sample was immediately placed into liquid N₂ and stored at –80°C until metabolite analyses were performed. The second tissue sample was used for determination of SR Ca²⁺-handling properties.

Assessment of muscle metabolites.   Muscle metabolite analyses were performed as previously reported (27), using fluorometric procedures (46) and HPLC techniques (38). The specific metabolites analyzed include ATP, PCr, ADP, AMP, IMP, inorganic phosphate (P_i), creatine (Cr), and a range of glycolytic intermediates, including Lac. The concentrations of free ADP (ADP_f) and free AMP (AMP_f) were calculated, as previously described (61).

Quantification of total glycogen, proglycogen, and macroglycogen was accomplished using fluorometric techniques (50). All measurements, with the exception of muscle glycogen content, were performed using an extract from the same piece of freeze-dried tissue and were corrected to the average total Cr content (TCr = PCr + Cr) of all tissue samples collected for each individual. Neither exercise nor glucose supplementation altered TCr. During a given analytic session, all tissue samples from three individuals were analyzed in duplicate for a given metabolite.

Assessment of SR Ca²⁺-handling properties.   To assess changes in Ca²⁺ transport across the SR membrane, a variety of functional properties of SERCA, the CRC, and the SR membrane were performed using crude muscle homogenates. Muscle samples (40–60 µg) were diluted 1:11 (wt/vol) in ice-cold homogenizing buffer (pH 7.5) containing (in mM) 250 sucrose, 5 HEPES, 0.2 PMSF, and 0.2% sodium azide (NaN₃). Dithiothreitol was not used in the preparation of crude muscle homogenates, since dithiothreitol could potentially reverse exercise-induced sulfhydryl oxidation during sample preparation (64). The muscles were hand homogenized with a Duall glass on glass hand homogenizer (Kontes Glass). Tissue samples were stored on ice from the time of extraction until homogenized. The total time between tissue extraction and homogenization was typically <10 min. Muscle homogenate aliquots (115 µl per aliquot; approximately four aliquots) were rapidly frozen in liquid N₂ and stored at –80°C for future analysis of SR function. Assessment of the kinetics parameters of SERCA included V_max, Hill coefficient (n_H) (defined as the relationship between SERCA activity and [Ca²⁺]_f for 10–90% V_max), which is a measure of the cooperative binding behavior of SERCA for Ca²⁺, and the Ca₅₀ (defined as the [Ca²⁺]_f at 50% V_max), which is also a measure of cooperative binding behavior. Additionally, we determined the relative amounts of passive Ca²⁺ leak across the SR membrane by comparing the ratio between V_max in the presence and absence [V_max(–)] of 1 µM Ca²⁺ ionophore A23187 [GenBank] (ionophore ratio).

SERCA activity.   Measurement of SERCA kinetic properties was performed using a spectrophotometric assay (68), as modified in our laboratory (75). Assay protocols were further adapted for use on a plate reader (SPECTRAmax Plus; Molecular Devices). With this protocol, we analyzed three different muscle samples simultaneously on a single plate. Each sample was prepared by adding 40 µl of crude muscle homogenate to 5 ml cocktail buffer, and then each sample was aliquoted (300 µl) into 16 Eppendorf tubes and mixed with Ca²⁺ to generate 14 different Ca²⁺ concentrations, ranging between 7.6 and 4.7 pCa units. The difference in assay conditions was limited to one Eppendorf tube, in which Ca²⁺ ionophore A23187 [GenBank] was not included in the reaction cocktail. In all other Eppendorf tubes and for all 14 Ca²⁺ concentrations used to generate the substrate-SERCA activity curve, the Ca²⁺ ionophore A23187 [GenBank] (1 µM Sigma C-7522) was included. The Ca²⁺ ionophore A23187 [GenBank] was used to prevent the formation of a large Ca²⁺ gradient across the SR membrane. By measuring maximal SERCA activity in the presence (V_max) and in the absence [V_max(–)] of Ca²⁺ ionophore A23187 [GenBank] , passive Ca²⁺ leak through the SR membrane was assessed. Assessment of V_max, V_max(–), and calculation of ionophore ratios [V_max/V_max(–)] were completed at a pCa of 4.9 for all samples. On a given analytic day, complete sets of samples from six individuals were analyzed for SERCA kinetics in duplicate.

In the measurement of V_max, phosphatase inhibitors were not employed in the extraction buffer. The possibility exists that exercise and/or glucose effects on phosphorylation on the CRC or SERCA directly or via PLN could be obviated via activation of phosphatases during homogenization. To examine this possibility, we have measured V_max in four to six individuals at rest and during exercise, with and without phosphatase inhibitors (Phosphatase Inhibitor Cocktail I, Sigma P2850) in the extraction buffer. For the control or NG condition, the difference between means for the two conditions ranged between 2.0 and 6.3%. For the experimental or G condition, the differences at similar exercise times ranged between 0.6 and 6.1%. These data suggest that V_max is not affected by the addition of phosphatase inhibitors.

SR Ca²⁺ uptake.   Oxalate-supported Ca²⁺-uptake rates were measured using the Ca²⁺ fluorescent dye indo 1, according to published methods (56, 57), as modified by Ruell et al. (60) and our laboratory (76). Fluorescence measurements were made on a spectrofluorometer (Ratiomaster system, Photon Technology International) equipped with dual-emission monochromators. The measurement of [Ca²⁺]_f using this procedure is based on the difference in maximal emission wavelengths between the Ca²⁺-bound indo 1 complex and the Ca²⁺-free indo 1 complex, as described previously in this paper and earlier publications (56, 57).

The reaction buffer (pH 7.0) for muscle homogenates contained 200 mM KCl, 20 mM HEPES, 15 mM MgCl₂, 10 mM NaN₃, 10 mM phosphoenolpyruvate, 5 mM oxalate, and 5 µM N,N,N',N'-tetrakis(2-pyridlymethyl)ethylanediamine. Before each assay, 1.5 µM indo 1, 18 U/ml Lac dehydrogenase, and 18 U/ml pyruvate kinase were added to the 2 ml of reaction buffer. Immediately before collection of emission spectra, a volume of muscle homogenate was added to the cuvette containing the reaction buffer. Following initiation of data collection, 2.5 µl of 10 mM CaCl₂ were added to the cuvette, which produces a consistent starting [Ca²⁺]_f of 3.5 µM. Shortly after the attainment of a constant [Ca²⁺]_f, 5 mM ATP were added to the cuvette to initiate Ca²⁺ uptake. The curve generated from [Ca²⁺]_f vs. time was then smoothed over 21 points using the Savitsky-Golay algorithm. Linear regression was performed on values ranging ±100 nM, at [Ca²⁺]_f of 500, 1,000, 1,500, and 2,000 nM. Differentiating the linear fit curve allowed determination of Ca²⁺-uptake rates. On a given analytic day, complete sets of samples from three individuals were analyzed for Ca²⁺-uptake kinetics in duplicate.

We have also assessed the apparent coupling ratio, which provides an indication of the efficiency of SERCA to transport Ca²⁺ from the cytosol into the lumen of the SR per ATP hydrolyzed, by calculating the ratio between Ca²⁺ uptake at 2,000 nM and V_max. It should be emphasized that the ratio was determined under different assay conditions that were optimized for measurement of each property. Maximal SERCA activity was assessed in the presence of the Ca²⁺ ionophore A23187 [GenBank] and at [Ca²⁺]_f 12.5 µM, whereas, Ca²⁺-uptake rates were determined in the absence of Ca²⁺ ionophore A23187. [GenBank] Additionally, our measurements of Ca²⁺ uptake are submaximal, since the Ca²⁺ sensitivity of indo 1 is limited at high [Ca²⁺]_f. We have used the term "apparent coupling ratio" to indicate that the parameters used to calculate this ratio were measured under different assay conditions.

SR Ca²⁺ release.   SR Ca²⁺-release rates were measured according to the methods of Ruell et al. (60), as modified by our group (76). SR Ca²⁺-release assays were conducted similar to the Ca²⁺-uptake assay procedures, where a dual-emission spectrofluorometer (Ratiomaster system, Photon Technology International) records simultaneous photon counts per second for indo 1 emission wavelengths previously defined. To assess Ca²⁺-release kinetics, homogenate samples were actively loaded with Ca²⁺ until a characteristic plateau in [Ca²⁺]_f was achieved. At this point, 20 mM 4-chloro-m-cresol were added to the assay mixture to chemically stimulate Ca²⁺ release in vitro. Two different properties of Ca²⁺ release have been reported in this paper. Phase 1 Ca²⁺ release has been characterized as the peak rate of the initial, fast phase of Ca²⁺ release that lasts 1–3 s in duration, while Phase 2 Ca²⁺ release has been characterized as the more prolonged, slower rate of Ca²⁺ release lasting from 4–10 s. Both phases of Ca²⁺ release have been calculated using the same methods as described for Ca²⁺ uptake, where the ionized Ca²⁺ concentration is calculated as described previously (29). Subsequently, differentiating the linear fit curves allows determination of Ca²⁺-release rates. On a given analytic day, Ca²⁺-release kinetics were assessed from complete sets of samples from three individuals using the same assay sample as that used to assess Ca²⁺ uptake.

Protein determination of homogenates was made by the method of Lowry and Passonneau (46), as modified by Schacterle and Pollock (63). Samples were analyzed in triplicate to determine protein concentration.

Western blot analysis.   To assess PLN phosphorylation site-specific polyclonal antibodies for anti-serine 16 (Ser16) PLN (sc-12963) and anti-threonine 17 (Thr17) PLN (sc-17024), phosphorylated forms of PLN were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). Samples (5–50 µg protein with quantity dependent on specific antibody) were loaded on 12.5% polyacrylamide gels. Proteins were separated using standard SDS-PAGE protocols and transferred to nitrocellulose membranes. After blocking with a 5% skim milk suspension, the membranes were treated with site-specific polyclonal antibodies raised against the Ser16 and Thr17 phosphorylated forms of PLN, washed in Tris·HCl, pH 7.5, 150 mM NaCl, 0.1% Tween 20 (Tris-buffered saline/0.1% Tween), and treated with horseradish peroxidase-conjugated secondary antibody (Santa Cruz Biotechnology). Membranes were washed in Tris-buffered saline, 0.1% Tween, and the signals were detected with an enhanced chemiluminescence kit (Amersham Biosciences) using a bio-imaging system and the GeneSnap software (Syngene) obtained from Fisher. Relative phosphorylation levels were determined by quantifying the optical density of bands at 10 and 25 kDa, as specified on the material data sheet for the PLN antibodies. For each antibody, the linearity of progressive increases in protein content was established before experiments were conducted (data not shown). Relative protein levels were determined by scanning densitometry, and values were expressed as a percentage of standard. When direct comparisons were made between sampling times, values were normalized to tissue samples collected at rest (0 min) and expressed as percentage of rest. All samples were analyzed in duplicate and on different gels.

Statistical analyses.   Data are presented as means ± SE. A one-way ANOVA for repeated measures was utilized to compare differences between the exercise times within each condition. A two-way ANOVA (2 repeated measures) was utilized to discriminate between differences resulting from experimental condition and exercise time for matched samples. The data collected in the subgroup of participants (i.e., 6 of 15 volunteers) who had a tissue sample collected at a matched time corresponding to fatigue in NG (i.e., 115 ± 6 min) during both conditions were analyzed using a different two-way ANOVA to discriminate between differences resulting from experimental condition and sampling time for matched samples. Where significant differences were found, Neuman-Keuls post hoc procedures were used to compare specific means. Significance was accepted at P < 0.05. Where a difference between means is indicated in the text, significance is implied.

	RESULTS

TOP ABSTRACT METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
Respiratory gas and substrate oxidation.   The relative exercise intensities based on measurements performed at 15 min were 57.2 ± 1.9 and 57.8 ± 1.2% O_2peak for the NG and G conditions, respectively. No differences in O₂, CO₂, or RER were observed between conditions at rest or during exercise (Table 1). However, main effects of exercise were found for O₂, CO₂, and RER. For O₂, rest < 15 min, and 30 min < 60 min, 90 min, 115 ± 6 min, and 137 ± 7 min. For CO₂, rest < 15 min, 30 min, 60 min, 90 min, 115 ± 6 min, and 137 ± 7 min. For RER, rest < 15 min, and 30 min > 60 min, 90 min, 115 ± 6 min, and 137 ± 7 min.

View larger version (13K): [in this window] [in a new window]   Fig. 1. Blood glucose (A) and lactate (B) concentrations during prolonged exercise, with (G) and without oral glucose (NG) supplementation. Values are means ± SE (n = 15). Exercise end points represent the time to fatigue for NG and G. For NG and G, the times to fatigue were 115 ± 6 and 137 ± 7 min, respectively. Significantly different from *0 min, 15 min, 30 min, and °60 min (P < 0.05). Significantly different from #G and $NG fatigue time point (P < 0.05).

Muscle metabolites and substrates.   Muscle ATP concentrations were not altered by exercise or by glucose supplementation (Table 3). The total content of both ADP and AMP was also unaffected by our exercise or glucose supplementation (data not presented). Additionally, PCr, P_i, and Cr concentrations measured at rest and exercise were not different between NG and G. However, main effects of exercise were found for PCr, P_i, and Cr concentrations. For PCr, rest > 30 min, 90 min, 115 ± 6 min, and 137 ± 7 min. For P_i and Cr, rest < 30 min, 90 min, 115 ± 6 min, and 137 ± 7 min. A main effect of exercise (rest < 30 min, 90 min, 115 ± 6 min, and 137 ± 7 min) was found for IMP. Glucose supplementation did not alter this exercise effect.

Muscle glucose increased with exercise such that rest < 30 min and 90 min, and 30 min > 90 min, 115 ± 6 min, and 137 ± 7 min (Table 4). For muscle Lac, rest < 30 min, 90 min, 115 ± 6 min, and 137 ± 7 min, and 30 min > 90 min, 115 ± 6 min, and 137 ± 7 min.

Preexercise and exercise total glycogen, proglycogen, and macroglycogen contents were not different between NG and G conditions (Fig. 2). During NG, total muscle glycogen content was reduced by 42, 66, and 76% after 30 and 90 min of exercise and at fatigue, respectively. During G, total muscle glycogen content was reduced by 40, 62, 68, and 72% after 30, 90, 115 ± 6, and 137 ± 7 min of exercise, respectively. Similar percent changes were also observed during exercise for proglycogen and macroglycogen.

View larger version (10K): [in this window] [in a new window]   Fig. 2. Total muscle glycogen (A), macroglycogen (C), and proglycogen (B) before and during prolonged exercise, with and without glucose supplementation. Values are means ± SE (n = 15). Exercise end points represent the time to fatigue for NG and G. For NG and G, the times to fatigue were 115 ± 6 and 137 ± 7 min, respectively. A main effect (P < 0.05) of exercise was found for total, pro-, and macroglycogen content. For exercise, 0 > 30 min > 90 min > 115 ± 6 min > 137 ± 7 min.

View larger version (10K): [in this window] [in a new window]   Fig. 3. Maximal sarcoplasmic reticulum Ca2+-ATPase activity (A), Ca2+ uptake (B), and coupling ratio (C) during prolonged exercise, with and without glucose supplementation. Values are means ± SE (n = 15). Exercise end points represent the time to fatigue for NG and G. For NG and G, the times to fatigue were 115 ± 6 and 137 ± 7 min, respectively. Vmax, maximal Ca2+-ATPase activity; pro, protein. Ca2+ uptake was determined at 2,000 nM Ca2+. Apparent coupling ratio is defined as the ratio between Ca2+ uptake and Vmax. Main effects (P < 0.05) of exercise were found for Vmax and Ca2+ uptake. For exercise, rest, 30 min > 90 min > 115 ± 6 min, 137 ± 7 min.

SR Ca²⁺-uptake rates (nmol·mg protein^–1·min^–1) between NG and G conditions were assessed at 2,000 nM (Fig. 3B). No differences in resting Ca²⁺-uptake rates were observed between NG (6.4 ± 0.3) and G (6.3 ± 0.4). Exercise reduced (main effect; rest > 30 min and 90 min > fatigue) Ca²⁺-uptake rates during both NG and G conditions. During NG, Ca²⁺-uptake rates were reduced by 10.8, 15.8, and 26.9% after 30, 90, and 115 ± 6 min of exercise, respectively. Similar reductions were observed in the G condition, at comparable time points. Glucose supplementation was without effect in altering the magnitude of change induced by exercise. In addition to Ca²⁺-uptake measurements made at 2,000 nM, we also assessed Ca²⁺-uptake rates at 1,500, 1,000, and 500 nM (date not presented). At these Ca²⁺ concentrations, a similar pattern of change occurred during exercise as at 2,000 nM. Glucose supplementation did not further alter the exercise-induced reductions in Ca²⁺-uptake rates at the lower Ca²⁺ concentrations.

To assess the effects of exercise and glucose supplementation on the efficiency of Ca²⁺ transport across the SR membrane, we have calculated apparent coupling ratios (Ca²⁺ uptake rate at 2,000 nM/V_max) (Fig. 3C). No changes in the apparent coupling ratio were found with exercise for both NG and G.

No differences in resting Ca²⁺-release rates for either Phase 1 or Phase 2 were observed between NG (Phase 1, 23.4 ± 1.3; Phase 2, 7.3 ± 0.4) and G (Phase 1, 23.0 ± 1.1; Phase 2, 7.3 ± 0.5), respectively (Fig. 4). Exercise reduced both Phase 1 and Phase 2 Ca²⁺-release rates in both NG and G conditions. For Phase 1 Ca²⁺ release, rest > 30 min, 90 min, 115 ± 6 min, and 137 ± 7 min, and 30 min > 115 ± 6 min and 137 ± 7 min. For Phase 2 Ca²⁺ release, rest > 30 min, 90 min, 115 ± 6 min, and 137 ± 7 min, and 90 min > 115 ± 6 min and 137 ± 7 min. Glucose supplementation did not alter the exercise-induced reductions in Ca²⁺-release kinetics.

View larger version (12K): [in this window] [in a new window]   Fig. 4. Sarcoplasmic reticulum Ca2+-release rates for Phase 1 (A) and Phase 2 (B) during prolonged exercise, with and without glucose supplementation. Values are means ± SE (n = 15). Exercise end points represent the time to fatigue for NG and G. For NG and G, the times to fatigue were 115 ± 6 and 137 ± 7 min, respectively. For both Phase 1 and Phase 2 Ca2+-release rates, a main effect (P < 0.05) of exercise was found. For Phase 1, rest > 30 min, 90 min, 115 ± 6 min, and 137 ± 7 min; 30 min > 115 ± 6 min and 137 ± 7 min. For Phase 2, rest > 30 min, 90 min, 115 ± 6 min, and 137 ± 7 min.

View larger version (11K): [in this window] [in a new window]   Fig. 5. Representative immunoblots for phospholamban phosphorylation measured at rest and during exercise in the NG and G conditions for Ser16 (A) and Thr17 (B). Rest, before exercise; 30, 90, 115 ± 6, and 137 ± 7, time of exercise; Ser16, phospholamban phosphorylation at serine 10 and 25 kDa; Thr17, phospholamban phosphorylation at threonine 10 and 25 kDa.

	DISCUSSION

TOP ABSTRACT METHODS RESULTS DISCUSSION GRANTS REFERENCES

Contrary to our hypothesis, we found no effect of oral glucose supplementation in modifying the SR Ca²⁺-cycling responses to prolonged exercise. In our experimental design, oral glucose supplementation began at 30 min of exercise, providing a lead-in period to allow for the exercise response to become established. Differences between NG and G in the SR properties examined were expected at 90 min of exercise, the first tissue sampling point after the beginning of glucose supplementation and at the matched time point beyond 90 min of exercise, representing the point of fatigue in NG. Since our hypothesis was based on several previous studies using repetitive contractions, which demonstrated a protective effect on a SR Ca²⁺ function during increased CHO availability, at least with regard to endogenous muscle glycogen levels (7, 12, 16, 43), the failure to find an experimental effect invites further examination.

Oral glucose supplementation could promote a variety of responses, many of which could potentially affect SR function during the exercise state. To examine these possibilities, it was first necessary to determine that our glucose supplement schedule could abolish the reduction in blood glucose observed at 90 min of exercise and beyond. As expected, based on previous studies (9, 11, 55), blood glucose remained stable throughout exercise in G.

A second objective of this study was to determine whether maintenance of blood glucose concentration could alter muscle substrate utilization and metabolism. Glucose supplementation did not change the metabolic efficiency of the exercise, as indicated by the lack of differences in O₂ between conditions or the contribution of either CHO or fats in substrate oxidation. In addition, we found no effect of glucose supplementation on either glycogen content or cellular energy responses during the exercise. In this regard, our data support previous studies that have not found an effect of glucose supplementation on the magnitude of exercise-induced glycogen depletion (11, 55). Given the potential functional differences between the different forms of glycogen (50, 67), we have also measured macro- and proglycogen contents in addition to total glycogen. Our results indicate that no differences existed between NG and G conditions in the pattern of depletion that occurred, regardless of the glycogen moiety examined.

Similar to previous studies (55, 72), there was a failure of G to alter the change in the high-energy phosphates, ATP and PCr, and their metabolites, P_i and Cr, that normally occurs with exercise. We had anticipated that G would attenuate the increase in IMP, recognized as a more sensitive indication of ATP (32) observed with exercise based on a previous study (69). Although we found a clear trend toward lower IMP after glucose feeding, the difference was not significant. Our failure to find an effect on IMP accumulation between conditions, as shown previously (69), could be due to differences in exercise protocol and/or the training state of the participants, as well as the glucose supplement schedule. It should be noted that a variety of glycolytic intermediates were measured, all of which were not significantly different, either at rest or during exercise between conditions (data not presented). The lack of an effect of glucose supplementation on the metabolic properties could contribute to our negative results, since accumulation of one or more of these compounds has been shown to alter both SR Ca²⁺ release (18) and Ca²⁺ uptake (73). Since muscle glycogen levels appear to be directly involved in modifying the contractile-induced effect on SR Ca²⁺ cycling (7, 16, 43), the failure of our intervention to modify glycogen contents may also be important in our inability to realize an experimental effect on SR responses.

Another argument for glucose supplementation to effect changes in SR function during exercise is via differences in selected blood hormonal responses. Blood glucose levels have a potent effect in regulating the secretion of both Ins from the pancreas and the catecholomines from the adrenal medulla (24, 79). Our results demonstrate marked differences between conditions in the response of these hormones to exercise (25). With G, the normal reduction in serum Ins to prolonged exercise was blunted, while both the Epi and NE time-dependent increases with exercise were substantially reduced. Our findings clearly indicate that, by preventing declines in blood glucose during prolonged exercise, the concentration of these hormones can be substantially altered. Our observations are in accord with other studies, which have reported similar effects on these hormones, with glucose supplementation administered during sustained submaximal exercise (24).

The blunting of the catecholamine response with glucose phosphorylation would be expected to modify the intrinsic regulation of both the CRC and SERCA. It is generally accepted that Epi can effect increases in PLN phosphorylation through cAMP-dependent mechanisms, resulting in an increase in Ca²⁺ sensitivity in the absence of changes in V_max of the enzyme (48). Since we failed to observe changes in SR Ca²⁺-cycling sensitivity with prolonged exercise, an increase in plasma Epi concentration by itself is not effective in modifying the sensitivity of the enzyme to Ca²⁺. The failure of Epi to alter Ca²⁺ sensitivity is also supported by our measurements of site-specific phosphorylation of PLN at Ser16 and Thr17, where expected increases (59) with G were not observed.

Epi and Ins signaling can activate various protein kinase and phosphatase pathways to alter CRC phosphorylation (45, 48, 58), thereby regulating the open probability of the CRC. However, since we did not observe any changes in SR Ca²⁺-release kinetics between conditions, it appears unlikely that increases in CRC phosphorylation were induced by glucose supplementation. Since plasma Epi decreased and serum Ins increased with G during prolonged exercise, it is possible that contrasting effects of the two hormones negated a net phosphorylation.

Another possibility to explain the apparent insignificant effect of glucose supplementation on SR behavior during exercise relates to the analytic approach to measuring the SR Ca²⁺-cycling properties. Our measurements are based on in vitro techniques performed on crude homogenates under supposedly optimal conditions. As such, the increases in Ca²⁺ release and SR Ca²⁺ uptake that undoubtedly occurs "in vivo" in the transition from rest to exercise remain undetected. It is possible that changes in vivo mediated by our experimental conditions remain obscure because of analytic limitations. However, in recent work (T. A. Duhamel and H. J. Green, unpublished observations), we have been able to demonstrate that, under supposedly optimal assay conditions, the effect of changes in selected hormones and selected protein kinase and phosphatase signaling pathways on the kinetic behavior of the Ca²⁺-ATPase can be detected in vitro. These results suggest that at least some of the potential mechanisms whereby glucose can alter SR Ca²⁺ handling can be assessed in vitro under our assay conditions.

A limitation of the present study was our failure to investigate the potential role of the extraction procedure on the activation of select protein kinases and phosphatases on the phosphorylation of the proteins involved in SR Ca²⁺ cycling. It is important to note, as an example, that the lack of inclusion of protein phosphatase inhibitors within the extraction buffer may have not allowed the proper assessment of changes in PLN phosphorylation (59) and conceivably in vitro SR function. The activation of selected protein kinases and phosphatases could alter the phosphorylation status of the CRC and SERCA directly or, in the case of SERCA via the regulatory proteins, PLN and sarcolipin (SLN). Although we have examined the effect of several phosphatases on the V_max behavior of SERCA on the limited amount of tissue remaining after the initial measurements performed without any phosphatase inhibitors and found no effect, the possibility remains that the sensitivity of SERCA to Ca²⁺ may have been altered. Measurements of Ca²⁺ sensitivity would appear to be particularly important if the phosphorylation status of PLN and/or SLN were altered.

It is also possible that the isolated effects of glucose supplementation on SR responses are not conspicuous because of the effect of exercise per se on the CRC and the SERCA. Repetitive contractile activity results in a decrease in V_max, an effect that has been attributed to structural alterations in the region of the nucleotide binding site of the enzyme (47, 54). The structural alterations have been attributed to protein oxidation and nitration induced by accumulation of reactive oxygen species (39, 53). In the face of these changes, which result in reductions in V_max, it is unclear whether the mechanisms proposed to alter kinetic properties of the Ca²⁺-ATPase with glucose can act independently.

Accompanying the exercise-induced reductions in V_max and Ca²⁺-uptake rates were reductions in SR Ca²⁺-release kinetics. We have utilized a two-phase kinetic model to characterize distinct phases of Ca²⁺ release that occur following the addition of the Ca²⁺-release agent, 4-chloro-m-cresol. Phase 1 Ca²⁺ release has been characterized as the initial fast phase of Ca²⁺ release that lasts for 1–3 s in duration, while Phase 2 Ca²⁺ release has been characterized as the more prolonged, slower rate of Ca²⁺ release occurring from 4- to 10-s duration (76). In this study, exercise-induced reductions occurred for both Phase 1 and Phase 2 Ca²⁺ release rates, with the reductions being similar in both time and magnitude. Although the physiological significance of each Ca²⁺-release phase remains unclear, our previous work demonstrates that two separate phases of Ca²⁺ release occur (76). Based on previous studies employing prolonged exercise in rats (18), it would appear that the exercise-induced reductions in Ca²⁺-release kinetics can be attributed to a reduction in the number of functional CRC, possibly as a result of protein oxidation associated with the accumulation of reactive oxygen species (19).

The interpretation of the findings of this study must also consider the mixed fiber-type composition of the human vastus lateralis. Since type I and type IIa fibers (28, 78) represent 90% of the fiber population in the human vastus lateralis (62), the exercise-induced reductions in SR Ca²⁺-handling properties observed in this study represent the net change in SR function for all fiber types present in each muscle sample and, consequently, do not represent a fiber-type-specific response. We cannot discount the possibility that our data were influenced by greater reductions in one fiber type compared with another, since differences in fiber-type activation during prolonged exercise are known to exists (28, 78). Moreover, differences in SERCA protein isoform expression (83) or the expression of other SR-associated proteins known to influence the intrinsic regulation of SR properties (73) may have also caused SR properties to be affected to a greater extent in one fiber type compared with another. It would have been beneficial to determine the fiber-type-specific changes in SR properties during exercise for type I vs. type II fibers; however, this was not possible, given technical limitations in measuring SR Ca²⁺-cycling properties in single fibers.

In summary, our findings demonstrate that prolonged exercise leads to a progressive loss of SR Ca²⁺-handling properties in human skeletal muscle when assessed in vitro. Furthermore, it appears that the ingestion of oral glucose supplements initiated after the beginning of exercise does not alter the reductions in SR Ca²⁺-handling properties. We have also been able to demonstrate that, with our exercise protocol, glucose supplementation did not alter the metabolic response or the glycogen depletion patterns. However, glucose supplementation did modify both the plasma catecholamine and serum Ins responses. Although glucose supplementation did increase cycle time to fatigue, our SR Ca²⁺-cycling responses do not explain this ergogenic effect. However, in a companion paper, we have been able to determine that glucose supplementation increases Na⁺-K⁺-ATPase activity (25), which occurs in conjunction with an improved protection of membrane activity (71). It should be noted that our glucose feeding schedule was designed to prevent the reduction in blood glucose concentration that typically occurs with prolonged exercise. This we have been able to accomplish. As such, our results may differ from other studies that have used different glucose dosages and schedules designed to elevate blood glucose above resting levels during sustained exercise.

	GRANTS

TOP ABSTRACT METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
We gratefully acknowledge the financial assistance provided for this research by Quaker Oats (Gatorade Sports Science Institute) and the National Sciences and Engineering Research Council.

	FOOTNOTES

 

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

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

	REFERENCES

TOP ABSTRACT METHODS RESULTS DISCUSSION GRANTS REFERENCES

 

1. Allen DG, Kabbara AA, Westerblad H. Muscle fatigue: the role of intracellular calcium stores. Can J Appl Physiol 27: 83–96, 2002.[Web of Science][Medline]
2. Berchtold MW, Brinkmeier H, Müntener M. Calcium ion in skeletal muscle: its crucial role for muscle function, plasticity and disease. Physiol Rev 80: 1216–1265, 2000.
3. Bergström J, Hermansen L, Hultman E, Saltin B. Diet, muscle glycogen and physical performance. Acta Physiol Scand 71: 140–150, 1967.[Web of Science][Medline]
4. Booth J, McKenna MJ, Ruell PA, Gwinn TH, Davis GM, Thompson MW, Harmer AR, Hunter SK, Sutton JR. Impaired calcium pump function does not slow relaxation in human skeletal muscle after prolonged exercise. J Appl Physiol 83: 511–521, 1997.[Abstract/Free Full Text]
5. Byrd SK, Bode AK, Klug GA. Effects of exercise of varying duration on sarcoplasmic reticulum function. J Appl Physiol 66: 1383–1388, 1989.[Abstract/Free Full Text]
6. Byrd SK, McCutcheon LJ, Hodgson DR, Gollnick PD. Altered sarcoplasmic reticulum function after high intensity exercise. J Appl Physiol 67: 2072–2077, 1989.[Abstract/Free Full Text]
7. Chin ER, Allen DG. Effects of reduced muscle glycogen concentration on force, Ca²⁺-release and contractile protein function in intact mouse skeletal muscle. J Physiol 498: 17–29, 1997.[Abstract/Free Full Text]
8. Christ-Roberts CY, Mandarino LJ. Glycogen synthase: key effects of exercise on insulin action. Exerc Sport Sci Rev 32: 90–94, 2004.[CrossRef][Web of Science][Medline]
9. Coggan AR, Coyle EF. Carbohydrate ingestion during prolonged exercise: effects on metabolism and performance. In: Exercise and Sport Science Reviews, edited by Holloszy JO. Baltimore, MD: Williams and Wilkins, 1991, p. 1–40.
10. Coyle EF. Carbohydrate feeding during exercise. Int J Sports Med 13: S126–S128, 1992.[Web of Science][Medline]
11. Coyle EF, Coggan AR, Hemmert MK, Ivy JL. Muscle glycogen utilization during prolonged strenuous exercise when fed carbohydrate. J Appl Physiol 61: 165–172, 1986.[Abstract/Free Full Text]
12. Cuenda A, Nogues M, Henao F, Gutiérrez-Merino C. Interaction between glycogen phosphorylase and sarcoplasmic reticulum membranes and its functional implications. J Biol Chem 273: 11998–12004, 1995.
13. Duhamel T, Green H, Perco JG, Ouyang J. Metabolic and sarcoplasmic reticulum Ca²⁺-cycling responses in human muscle four days following prolonged exercise. Can J Physiol Pharmacol 83: 643–655, 2005.[CrossRef][Web of Science][Medline]
14. Duhamel TA, Green HJ, Perco JD, Sandiford SD, Ouyang J. Human muscle sarcoplasmic reticulum function during submaximal exercise in normoxia and hypoxia. J Appl Physiol 97: 180–187, 2004.[Abstract/Free Full Text]
15. Duhamel TA, Green HJ, Sandiford SD, Perco JG, Ouyang J. Effects of progressive exercise and hypoxia on human muscle sarcoplasmic reticulum function. J Appl Physiol 97: 188–196, 2004.[Abstract/Free Full Text]
16. Duhamel TA, Perco JG, Green HJ. Manipulation of dietary carbohydrates after prolonged effort modifies muscle sarcoplasmic reticulum responses in exercising males. Am J Physiol Regul Integr Comp Physiol 291: R1100–R1110, 2006.[Abstract/Free Full Text]
17. Dux L, Green HJ, Pette D. Chronic low-frequency stimulation of rabbit fast-twitch muscle induces partial inactivation of the sarcoplasmic reticulum Ca²⁺-ATPase and changes in its tryptic cleavage. Eur J Biochem 195: 92–100, 1990.
18. Favero TG. Sarcoplasmic reticulum Ca²⁺-release and muscle fatigue. J Appl Physiol 87: 471–483, 1999.[Abstract/Free Full Text]
19. Favero TG, Pessah IN, Klug GA. Prolonged exercise reduces Ca²⁺-release in rat skeletal muscle sarcoplasmic reticulum. Pflügers Arch 422: 472–475, 1993.[CrossRef][Web of Science][Medline]
20. Favero TG, Zable AC, Bowman MB, Thompson A, Abramson J. Metabolic end products inhibit sarcoplasmic reticulum Ca²⁺ release and [³H]ryanodine binding. J Appl Physiol 78: 1665–1672, 1995.[Abstract/Free Full Text]
21. Ferrington DA, Reijneveld JC, Bär PR, Bigelow DJ. Activation of the sarcoplasmic reticulum Ca²⁺-ATPase induced by exercise. Biochim Biophys Acta 1279: 203–213, 1996.[Medline]
22. Frayn KN. Calculation of substrate oxidation rates in vivo from gaseous exchange. J Appl Physiol 55: 628–634, 1983.[Abstract/Free Full Text]
23. Galbo H. Hormonal and Metabolic Adaptation to Exercise. New York: Thieme-Stratton, 1983.
24. Galbo H. Exercise physiology: humoral function. In: Sport Science Review: Sport Physiology. Champaign IL: Human Kinetics, 1992, p. 65–93.
25. Green HJ, Duhamel TA, Foley KP, Ouyang J, Smith IC, Stewart RD. Glucose supplements increase human muscle Na⁺-K⁺-ATPase activity in vitro during prolonged exercise. Am J Physiol Regul Integr Comp Physiol 293: R354–R362, 2007.[Abstract/Free Full Text]
26. Green HJ, Jones S, Ball-Burnett M, Fraser I. Early adaptations in blood substrates, metabolites, and hormones to prolonged exercise training in man. Can J Physiol Pharmacol 69: 1222–1229, 1991.[Web of Science][Medline]
27. Green HJ, Jones S, Ball-Burnett ME, Smith D, Livesey J, Farrance BW. Early muscular and metabolic adaptations to prolonged exercise training in man. J Appl Physiol 70: 2032–2038, 1991.[Abstract/Free Full Text]
28. Green HJ, Smith D, Murphy P, Fraser I. Training-induced alterations in muscle glycogen utilization in fibre-specific types during prolonged exercise. Can J Physiol Pharmacol 68: 1372–1376, 1990.[Web of Science][Medline]
29. Grynkiewicz G, Poenie M, Tsien RY. A new generation of Ca²⁺-indicators with greatly improved fluorescence properties. J Biol Chem 260: 3440–3450, 1985.[Abstract/Free Full Text]
30. Han JW, Thielezcek R, Varsányi M, Heilmeyer LM Jr. Compartmentalized ATP synthesis in skeletal muscle triads. Biochemistry 31: 377–384, 1992.[CrossRef][Medline]
31. Hargreaves M. Metabolic responses to carbohydrate ingestion: effects on exercise performance. In: Perspectives in Sports Medicine and Exercise Science, edited by Comb DR and Moury R. Carmel, IN: Cooper, 1999, p. 93–124.
32. Hochachka PW, Matheson GO. Regulating ATP turnover over broad dynamic work ranges in skeletal muscles. J Appl Physiol 73: 1697–1703, 1992.[Abstract/Free Full Text]
33. Holloway GP, Green HJ, Duhamel TA, Ferth S, Moule JW, Ouyang J, Tupling AR. Muscle sarcoplasmic reticulum Ca²⁺-cycling adaptations during 16 hours of heavy intermittent cycling exercise. J Appl Physiol 99: 836–843, 2005.[Abstract/Free Full Text]
34. Holloway GP, Green HJ, Tupling AR. Differential effects of repetitive activity on mechanical and sarcoplasmic reticulum responses in rat muscle of different oxidative potential. Am J Physiol Regul Integr Comp Physiol 290: R393–R404, 2006.[Abstract/Free Full Text]
35. Hughson RL, Green HJ, Sharratt MT. Gas exchange, blood lactate and plasma catecholamines during incremental exercise in hypoxia and normoxia. J Appl Physiol 79: 1134–1141, 1995.[Abstract/Free Full Text]
36. Hughson RL, Kowalchuk JM, Prime WM, Green HJ. Open-circuit gas exchange analysis in the non-steady state. Can J Appl Sport Sci 5: 15–18, 1980.[Medline]
37. Inashima S, Matsunaga S, Yasuda T, Wada M. Effect of endurance training and acute exercise on sarcoplasmic reticulum function in rat fast-and slow-twitch skeletal muscles. Eur J Appl Physiol 89: 142–149, 2003.[Web of Science][Medline]
38. Ingebretson DC, Bakken AM, Segadal L, Farstad M. Determination of adenine nucleotides and inosine in human myocardium by ion pair reversed phase high performance liquid chromatography. J Chromatogr 242: 119–126, 1982.[CrossRef][Web of Science][Medline]
39. Klebl BM, Ayoub TA, Pette D. Protein oxidation, tyrosine nitration, and inactivation of sarcoplasmic reticulum Ca²⁺-ATPase in low-frequency stimulated rabbit muscle. FEBS Lett 422: 381–384, 1998.[CrossRef][Web of Science][Medline]
40. Korge P. Factors limiting adenosine triphosphatase function during high intensity exercise. Sports Med 20: 215–225, 1995.[Web of Science][Medline]
41. Korge P. Factors limiting ATPase activity in skeletal muscle. In: Biochemistry of Exercise X, edited by Hargreaves M and Thompson M. Champaign, IL: Human Kinetics, 1998, p. 125–134.
42. Korge P, Campbell KB. The importance of the ATPase microenvironment in muscle fatigue: a hypothesis. Int J Sports Med 16: 172–179, 1995.[Web of Science][Medline]
43. Lees SJ, Franks PD, Spangenburg EE, Williams JH. Glycogen and glycogen phosphorylase associated with sarcoplasmic reticulum: effects of fatiguing activity. J Appl Physiol 91: 1638–1644, 2001.[Abstract/Free Full Text]
44. Lees SJ, Williams JH. Skeletal muscle sarcoplasmic reticulum glycogen status influences Ca²⁺-uptake supported by endogenously synthesized ATP. Am J Physiol Cell Physiol 286: C97–C104, 2004.[Abstract/Free Full Text]
45. Liu J, Brautigan DL. Insulin-stimulated phosphorylation of the protein phosphatase-1 striated muscle glycogen-targeting subunit and activation of glycogen synthase. J Biol Chem 275: 15940–15947, 2000.[Abstract/Free Full Text]
46. Lowry OH, Passonneau JV. A Flexible System of Enzymatic Analysis. New York: Academic, 1972.
47. Luckin KA, Favero TG, Klug GA. Prolonged exercise induces structural changes in SR Ca²⁺-ATPase of rat muscle. Biochem Med Metab Biol 46: 391–405, 1991.[CrossRef][Web of Science][Medline]
48. MacLennan DH, Abu-Abed M, Kang C. Structure-function relationships in Ca²⁺-cycling proteins. J Mol Cell Cardiol 34: 897–918, 2002.[CrossRef][Web of Science][Medline]
49. MacLennan DH, Asahi M, Tupling AR. The regulation of SERCA-type pumps by phosphorylation and sarcolipin. Ann NY Acad Sci 986: 472–480, 2003.[CrossRef][Web of Science][Medline]
50. Marchand I, Chorneyko K, Tarnopolsky M, Hamilton S, Shearer J, Potvin J, Graham TE. Quantification of subcellular glycogen in resting human muscle: granular size, number and location. J Appl Physiol 93: 1598–1607, 2002.[Abstract/Free Full Text]
51. Martonosi AN, Pikula S. The network of calcium regulation in muscle. Acta Biochim Pol 50: 1–30, 2003.[Web of Science][Medline]
52. Matsunaga S, Inashima S, Tsuchimochi H, Yameda T, Hazama T, Wada M. Altered sarcoplasmic reticulum function in rat diaphragm after high intensity exercise. Acta Physiol Scand 176: 227–232, 2002.[CrossRef][Web of Science][Medline]
53. Matsunaga S, Inashima S, Yamada T, Watanabe H, Hazama T, Wada M. Oxidation of sarcoplasmic reticulum Ca²⁺-ATPase induced by high intensity exercise. Pflügers Arch 446: 394–399, 2003.[Web of Science][Medline]
54. Matsushita S, Pette D. Inactivation of the sarcoplasmic reticulum ATPase in low frequency stimulated muscle results from a modification of the active site. Biochem J 285: 303–309, 1992.[Web of Science][Medline]
55. McConell G, Snow RJ, Proietto J, Hargreaves M. Muscle metabolism during prolonged exercise in humans: influence of carbohydrate availability. J Appl Physiol 89: 1083–1086, 1999.
56. O'Brien PJ. Microassay for malignant hyperthermia susceptibility: hypersensitive ligand-gating of the Ca channel in muscle sarcoplasmic reticulum causes increased amounts and rates of Ca-release. Mol Cell Biochem 93: 53–59, 1990.[CrossRef][Web of Science][Medline]
57. O'Brien PJ, Shen H, Weiler J, Ianuzzo CD, Wittnich C, Moe GW, Armstrong PW. Cardiac and muscle fatigue due to relative functional overload induced by excessive stimulation, hypersensitive excitation-contraction coupling, or diminished performance capacity correlates with sarcoplasmic reticulum failure. Can J Physiol Pharmacol 69: 262–268, 1991.[Web of Science][Medline]
58. Reiken S, Lacampagne A, Zhou H, Kerani A, Lehnart SE, Ward C, Huang F, Gaburjakova M, Gaburjakova J, Resemblit M, Warren M, He KL, Yi G, Wang J, Burkhoff D, Vassort G, Marks AR. PKA phosphorylation activates the calcium release channel (ryanodine receptor) in skeletal muscle: defective regulation in heart failure. J Cell Biol 160: 919–928, 2003.[Abstract/Free Full Text]
59. Rose AJ, Kiens B, Richter EA. Ca²⁺-calmodulin-dependent protein kinase expression and signalling in skeletal muscle during exercise. J Physiol 574: 889–903, 2006.[Abstract/Free Full Text]
60. Ruell PA, Booth J, McKenna MJ, Sutton JR. Measurement of sarcoplasmic reticulum function in mammalian skeletal muscle. Technical aspects. Anal Biochem 228: 194–201, 1995.[CrossRef][Web of Science][Medline]
61. Sahlin K, Palmskog G, Hultman E. Adenine nucleotides and IMP contents of the quadriceps muscle in man after exercise. Pflügers Arch 374: 193–198, 1978.[CrossRef][Web of Science][Medline]
62. Saltin B, Gollnick PD. Skeletal muscle adaptability: significance for metabolism and performance. In: Handbook of Physiology. Skeletal Muscle. Bethesda, MD: Am. Physiol. Soc., 1983, sect. 10, chapt. 19, p. 551–631.
63. Schacterle GR, Pollock RL. A simplified method for the quantitative assay of small amounts of protein in biologic material. Anal Biochem 51: 654–655, 1973.[CrossRef][Web of Science][Medline]
64. Schertzer J, Green H, Tupling R, Kollias H. Effects of sulphydryl oxidation on muscle sarcoplasmic reticulum Ca²⁺-ATPase activity in vitro (Abstract). Can J Appl Physiol 24: 476, 1999.
65. Schertzer JD, Green HJ, Duhamel TA, Tupling AR. Mechanisms underlying increases in SR Ca²⁺-ATPase activity following exercise in rat skeletal muscle. Am J Physiol Endocrinol Metab 284: E597–E610, 2003.[Abstract/Free Full Text]
66. Schertzer JD, Green HJ, Fowles JR, Duhamel TA, Tupling AR. Effects of prolonged exercise and recovery on sarcoplasmic reticulum Ca²⁺ cycling properties in rat muscle homogenates. Acta Physiol Scand 180: 195–208, 2004.[CrossRef][Web of Science][Medline]
67. Shearer J, Graham T. New perspectives on the storage and organization of muscle glycogen. Can J Appl Physiol 27: 179–203, 2002.[Web of Science][Medline]
68. Simonides WS, van Hardeveld C. An assay for sarcoplasmic reticulum Ca²⁺-ATPase activity in muscle homogenates. Anal Biochem 191: 321–331, 1990.[CrossRef][Web of Science][Medline]
69. Spencer MK, Yan Z, Katz A. Carbohydrate supplementation attenuates IMP accumulation in human muscle during prolonged exercise. Am J Physiol Cell Physiol 261: C71–C76, 1991.[Abstract/Free Full Text]
70. Stephenson DG, Nguyen LT, Stephenson GMM. Glycogen content and excitation-contraction coupling in mechanically skinned muscle fibres of the cane toad. J Physiol 519: 177–187, 1999.[Abstract/Free Full Text]
71. Stewart RD, Duhamel TA, Foley KP, Ouyang J, Smith IC, Green HJ. Protection of muscle membrane excitability during prolonged cycle exercise with glucose supplementation. J Appl Physiol 103: 331–339, 2007.[Abstract/Free Full Text]
72. Tsintzas OK, Williams C, Boobis LH, Greenhaff PL. Carbohydrate ingestion and single fiber glycogen metabolism during prolonged running in men. J Appl Physiol 81: 801–809, 1996.[Abstract/Free Full Text]
73. Tupling AR. The sarcoplasmic reticulum in muscle fatigue and disease: role of sarco(endoplasmic) reticulum Ca²⁺-ATPase. Can J Appl Physiol 29: 330–356, 2004.[Web of Science][Medline]
74. Tupling AR, Gramolini AO, Duhamel TA, Kondo H, Asahi M, Tsuchiya S, Borrelli M, Lepock JR, Otsu K, Hori M, MacLennan DH, Green HJ. HSP70 binds to the fast-twitch skeletal muscle sarco(endo)plasmic reticulum Ca²⁺-ATPase (SERCA1a) and prevents thermal inactivation. J Biol Chem 279: 52382–52389, 2004.[Abstract/Free Full Text]
75. Tupling R, Green HJ, Senisterra G, Lepock J, McKee N. Effects of 4-h ischemia and 1-h reperfusion on rat muscle sarcoplasmic reticulum function. Am J Physiol Endocrinol Metab 281: E867–E877, 2001.[Abstract/Free Full Text]
76. Tupling R, Green HJ. Silver ions induce Ca²⁺ release from the SR in vitro by acting on both the Ca²⁺ release channel and the Ca²⁺ pump. J Appl Physiol 92: 1603–1610, 2002.[Abstract/Free Full Text]
77. Tupling R, Green HJ, Roy BD, Grant S, Ouyang J. Paradoxical effects of prior activity on human sarcoplasmic reticulum Ca²⁺-ATPase response to exercise. J Appl Physiol 95: 138–144, 2003.[Abstract/Free Full Text]
78. Vøllestad NK, Blom PCS. Effect of varying exercise intensity on glycogen depletion in human muscle fibres. Acta Physiol Scand 125: 395–405, 1985.[Web of Science][Medline]
79. Wasserman DH, Cherrington AD. Regulation of extramuscular fuel sources during exercise. In: Handbook of Physiology. Exercise, Regulation and Integration of Multiple Systems. Bethesda, MD: Am. Physiol. Soc., 1996, sect. 12, chapt. 23, p. 1036–1074.
80. Williams JH. Contractile apparatus and sarcoplasmic reticulum function: effects of fatigue, recovery, and elevated Ca²⁺. J Appl Physiol 83: 444–450, 1997.[Abstract/Free Full Text]
81. Williams JH, Ward CW, Spangenburg EE, Nelson RM. Functional aspects of skeletal muscle contractile apparatus and sarcoplasmic reticulum after fatigue. J Appl Physiol 85: 619–626, 1998.[Abstract/Free Full Text]
82. Wolfe RR. Radioactive and Stable Isotope Tracers in Biomedicine. New York: Wiley-Liss, 1992.
83. Wu KD, Lytton J. Molecular cloning and quantification of sarcoplasmic reticulum Ca²⁺-ATPase isoforms in rat muscles. Am J Physiol Cell Physiol 264: C333–C341, 1993.[Abstract/Free Full Text]
84. Wuytack F, Raeymaekers L, Missiaen L. Molecular physiology of the SERCA and SPCA pumps. Cell Calcium 32: 279–305, 2002.[CrossRef][Web of Science][Medline]
85. Xu K, Zweier J, Becker L. Functional coupling between glycolysis and sarcoplasmic reticulum Ca²⁺-transport. Circ Res 77: 88–97, 1995.[Abstract/Free Full Text]
86. Xu KY, Becker LC. Ultrastructural localization of glycolytic enzymes on sarcoplasmic reticulum vesicles. J Histochem Cytochem 46: 419–427, 1998.[Abstract/Free Full Text]

This article has been cited by other articles:

				M. H. Fu and A. R. Tupling Protective effects of Hsp70 on the structure and function of SERCA2a expressed in HEK-293 cells during heat stress Am J Physiol Heart Circ Physiol, April 1, 2009; 296(4): H1175 - H1183. [Abstract] [Full Text] [PDF]

				M. Amann, S. M. Marcora, L. Nybo, T. A. Duhamel, T. D. Noakes, V. Jaquinandi, J. L. Saumet, P. Abraham, B. T. Ameredes, M. Burnley, et al. Viewpoint: Fatigue mechanisms determining exercise performance: integrative physiology is systems physiology. J Appl Physiol, May 1, 2008; 104(5): 1543 - 1544. [Full Text] [PDF]

				M. Hargreaves Fatigue mechanisms determining exercise performance: integrative physiology is systems biology J Appl Physiol, May 1, 2008; 104(5): 1541 - 1542. [Full Text] [PDF]

This Article Abstract Full Text (PDF) Free All Versions of this Article: 103/6/1986    most recent 01440.2006v1 Submit a response Alert me when this article is cited Alert me when eLetters are posted Alert me if a correction is posted Citation Map Services Email this article to a friend Similar articles in this journal Similar articles in Web of Science Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Citing Articles Citing Articles via HighWire Citing Articles via Web of Science (3) Citing Articles via Google Scholar Google Scholar Articles by Duhamel, T. A. Articles by Ouyang, J. Search for Related Content PubMed PubMed Citation Articles by Duhamel, T. A. Articles by Ouyang, J.