Journal of Applied Physiology Fuel your research with LabChart
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

J Appl Physiol 91: 1638-1644, 2001;
8750-7587/01 $5.00
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF) Free
Right arrow Submit a response
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via ISI Web of Science (20)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Lees, S. J.
Right arrow Articles by Williams, J. H.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Lees, S. J.
Right arrow Articles by Williams, J. H.
Vol. 91, Issue 4, 1638-1644, October 2001

Glycogen and glycogen phosphorylase associated with sarcoplasmic reticulum: effects of fatiguing activity

Simon J. Lees, Peter D. Franks, Espen E. Spangenburg, and Jay H. Williams

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


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

The purpose of the present study was to investigate the effects of fatiguing muscular activity on glycogen, glycogen phosphorylase (GP), and Ca2+ uptake associated with the sarcoplasmic reticulum (SR). Tetanic contractions (100 ms, 75 Hz) of the gastrocnemius and plantaris muscles, elicited once per second for 15 min, significantly reduced force to 26.5 ± 4.0% and whole muscle glycogen to 23% of rested levels. SR glycogen levels were 415.4 ± 76.6 and 20.4 ± 2.1 µg/mg SR protein in rested and fatigued samples, respectively. The optical density of GP from SDS-PAGE was reduced to 21% of control, whereas pyridoxal 5'-phosphate concentration, a quantitative indicator of GP content, was significantly reduced to 3% of control. GP activity after exercise, in the direction of glycogen breakdown, was reduced to 4% of control. Maximum SR Ca2+ uptake rate was also significantly reduced to 81% of control. These data demonstrate that glycogen and GP associated with skeletal muscle SR are reduced after fatiguing activity.

skeletal muscle; calcium handling; calcium-adenosinetriphosphatase


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

SKELETAL MUSCLE FATIGUE MANIFESTS as a result of repetitive or sustained muscle contraction and can be characterized by a temporary reduction in the capacity to generate force. It affects people in many ways such as decreased athletic performance and increased susceptibility to injury, and, more seriously, it can be a debilitating symptom of certain clinical conditions such as congestive heart failure (12, 32). Although the changes in muscle performance are well documented in the literature, the exact mechanisms that mediate these changes in muscle force are not fully understood.

The sarcoplasmic reticulum (SR) acts as an intracellular storage site for Ca2+ and also controls cytoplasmic Ca2+ concentration, which in turn regulates the force of muscle contraction. Diminished SR function may play a critical role in the development of fatigue (39, 40). Specifically, decreased rate of Ca2+ uptake may be directly responsible for slowing of relaxation as well as indirectly responsible for decreased force during fatigue. A fraction of muscle glycogen appears to be specifically associated with the SR in skeletal muscle (9, 15, 13). The binding of glycogen particles to the SR membrane is achieved by the hydrophobic tail of the glycogen-associated form of protein phosphatase 1 (20, 21). The estimated amount of glycogen associated with the SR varies greatly in the literature. Cuenda et al. (9) measured 32 µg/mg SR protein isolated from rat skeletal muscle, whereas Entman et al. (15) reported a range of 300-700 µg glycogen/mg SR membrane protein isolated from dog cardiac muscle. Because prolonged muscle contraction results in decreased muscle glycogen concentration (22), it seems logical that prolonged muscle contraction would also result in decreased glycogen associated with the SR.

Glycogen phosphorylase (GP), an enzyme involved in glycogenolysis, is also associated with the SR (11, 14, 38). Interestingly, GP may be associated with the SR via its binding to the glycogen particles (28, 38). Furthermore, glycogenolysis of SR glycogen may result in the release of GP from the membrane to the myoplasm. Cuenda et al. (9) showed that preparing SR from rabbits that were starved for 48 h (a treatment that causes glycogen depletion) resulted in a two- to fourfold decrease in GP activity and content. As for glycogen, GP associated with the SR might be expected to decline with exercise.

The specific purpose of this study was to investigate how prolonged muscle contraction affects glycogen concentration, GP content, and Ca2+ uptake rates associated with skeletal muscle SR.


    METHODS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Stimulation protocol. The Animal Use and Care Committee of Virginia Tech approved all procedures. Female Sprague-Dawley rats (200-225 g) were fed ad libitum (Purina rodent laboratory chow), allowed free access to water, and exposed to a 12:12-h light-dark cycle. The gastrocnemius and plantaris muscles from female Sprague-Dawley rats were used for all the experiments. One leg was stimulated (fatigued), and the contralateral leg served as the control (rested). Animals were anesthetized with pentobarbital sodium (90 mg/kg body wt). The sciatic nerve was surgically exposed so that the stimulation electrode could be attached without damaging the nerve. Surgical suture was then tied and glued at the insertion of the tendo calcaneus, and the distal portion of the calcaneus was cut with bone cutters. The soleus muscle was cut at its insertion so as to prevent it from contributing to force production. The animal was then placed in a prone position with the knee and distal end of the tibia securely clamped to prevent movement. The free end of the surgical suture was attached to an isometric transducer. A physiological saline solution (PSS) filled the chamber of the apparatus, covering the exposed portion of the leg, and a few drops of mineral oil were placed around the exposed sciatic nerve. The PSS was warmed to 37°C before the surgery and was maintained using a heat lamp. The PSS (pH 7.4) contained the following: 135 mM NaCl, 5 mM KCl, 2.5 mM CaCl2, 1 mM MgSO4, 1 mM NaH2PO4, and 13 mM NaHCO3. Tetanic contractions were elicited using a Grass S48 stimulator, and force measurements were measured, displayed, and recorded using a Harvard apparatus isometric transducer, Tecktronix 2201 oscilloscope, and Harvard chart recorder. The fatigue protocol entailed in situ stimulation for 15 min via the sciatic nerve. Contractions were elicited by 100-ms trains of pulses (75 Hz) delivered once per second.

Whole muscle glycogen. In the first set of experiments, the effect of stimulation on whole muscle glycogen was determined. Control and fatigue gastrocnemius and plantaris muscles were rapidly removed (within 60 s) and homogenized with three, 15-s bouts (VirTis VirTishear) in 5 ml of 0.6 M perchloric acid solution per gram of muscle wet weight and then frozen and stored at -80°C. Glycogen content was then determined in the homogenated using the glucoamylase (EC 3.2.1.3) method described by Keppler and Decker (23).

SR isolation. For the SR experiments, SR vesicles were prepared by the differential centrifugation method previously described by Williams et al. (41). Control and fatigued gastrocnemius and plantaris muscles were rapidly removed (within 60 s) and placed in 5 vol (wt/vol) of buffer containing 20 mM HEPES, 0.2% NaN3, and 0.2 mM phenylmethylsulfonyl fluoride (pH 6.8). They were then minced with scissors, homogenized (3 30-s bursts), and centrifuged at 8,000 g for 15 min. KCl (600 mM) was added to the supernatant to solubilize remaining actomyosin and centrifuged at 12,000 g for 45 min. The remaining supernatant was centrifuged at 49,000 g for 60 min, and the SR pellet (1-2 mg protein/ml) was resuspended in 20 mM HEPES, 0.2% NaN3, 0.2 mM phenylmethylsulfonyl fluoride, and 150 mM KCl (pH 6.8) and stored at -80°C until used. Samples used for determination of SR function were stored in the same buffer supplemented with 300 mM sucrose. The Bradford dye-binding assay adapted by Bio-Rad was used to determine protein concentration of the SR samples with BSA as a standard.

Measurement of SR glycogen. Glycogen associated with the SR was measured using a modification of the Keppler and Decker (23) approach. Resuspended SR (a volume containing 50 µg of protein) was incubated with shaking in 0.5 ml of an acetic acid (174 mmol/l) solution containing glucoamylase (8.7 kU/l) for 2 h at 40°C (pH 4.8). Fifty microliters of each glucoamylase-SR mixture was then added to 1 ml of a buffer (pH 7.5) containing the following: 0.3 M tetraethylammonium, 4 mM MgSO4, 120 mM KOH, 1 mM ATP, 0.9 mM beta -NADP+, 700 U/l glucose-6-phosphate dehydrogenase, and hexokinase (1.3 kU/l). After 10 min of incubation at room temperature, fluorescent emission of the reduced form of beta -NADPH was measured at 450 nm while the excitation wavelength was set at 365 nm. Glycogen standards were prepared using glycogen purified from rabbit liver. Approximately 99% of the standards were recovered. It is important to point out that no glucose could be detected in assays that were preformed without the glucoamylase incubation step.

Measurement of SR GP. GP content of the SR vesicles was determined using three different approaches: densitometric analysis of SDS-PAGE gels, measurement of enzyme activity, and measurement of pyridoxal 5'-phosphate (PLP) content. SDS-PAGE was performed following the method of Laemmli (25). Gels were loaded with 10 µg of protein and run on a mini-Protean II cell from Bio-Rad with a 5% acrylamide separating gel and a 4% acrylamide stacking gel. The running conditions were set at 45 mA (constant) until the tracking dye ran off the gel. Gels were stained overnight in a solution containing the following: 0.1% Coomassie blue R-250, 40% methanol, and 10% acetic acid. After staining, the gels were destained for ~1 h in a solution containing 50% H2O, 40% methanol, and 10% acetic acid. The bands corresponding to the molecular weight of SR Ca2+-ATPase and GP were scanned using MultiImage Light Cabinet from Alpha Innotech and analyzed using AlphaImager 2000 Documentation and Analysis System. From these scanned images, optical densities of these bands were determined.

Total GP activity, measured in the direction of glycogen breakdown, was done using a method adapted from Chi et al. (5). Five hundred nanograms of each SR sample were added to 100 µl of reagent (pH 7.0) containing the following: 50 mM imidazole, 50 mM glycogen, 20 mM K2HPO4, 0.5 mM MgCl2, 1 mM 5'-AMP, 2 µM glucose-1,6-bisphosphate, 0.5 mM dithiothreitol, 0.25% BSA, and 0.4 U/ml phosphoglucomutase. Because of the large difference in GP content between conditions, control samples were allowed to react for 15 min at room temperature, whereas fatigue samples were allowed to react for 1 h at room temperature. Glucose 1-phosphate production was linear over these time periods. Ten microliters of 0.5 N HCl were added to each sample to stop the reaction. Then they were then allowed to sit for 10 min at room temperature. One milliliter of a second reagent was added to the product of the first reaction. The second reagent contained the following: 50 mM Tris · HCl, 100 µM beta -NADP, 1 mM EDTA, 1 U/ml glucose-1,6-bisphosphate, and 0.35 U/ml phosphoglucoisomerase (pH 8.0). The samples were allowed to react in the second reagent for 20 min, and then the fluorescent emission of beta -NADPH was measured at 450 nm while the excitation wavelength was set at 365 nm. Standards used were 0, 5, 10, 25, and 35 nmol/5 µl of glucose 1-phosphate.

PLP associated with the SR vesicles was measured via HPLC using a method modified from those of Ubbink et al. (37), Mahuren and Coburn (27), Cordes and Jencks (7), and Gregory (17). By removing and measuring PLP, the GP content can be determined because over 95% of PLP in skeletal muscle is bound to this enzyme (4). To prevent PLP degradation, samples were prepared under yellow light. One-half milliliter of 10% trichloroacetic acid was added to 1 ml of sample (containing 200 µg of SR protein). The precipitated protein was then pelleted by centrifuging the samples at 1,000 g for 10 min. The supernatant was then removed, and 50 µl of 0.5 M semicarbazide were added while the pellet was discarded. The samples were then incubated for 15 min at 40°C. Diethyl ether (3 ml) was added to each sample, vortexed, and removed (after centrifugation at 1,000 g for 10 min). This last step was repeated. Dichloromethane (3 ml) was then added to each sample and vortexed, and then the supernatant was removed and saved (after centrifugation at 1,000 g for 10 min). One hundred microliters of this supernatant were injected into the HPLC system (Waters) and run through an ODS C18 4.6 mm × 25-cm Zorbax column from Mac-Mod Analytical with a mobile-phase buffer. The mobile-phase buffer was prepared in the following manner: 0.05 M KH2PO4 and 8% (vol/vol) acetonitrile were mixed in water, and pH was adjusted to 2.9 with o-phosphoric acid. The mobile-phase buffer was then filtered and degassed. The mobile phase was run at a pressure no greater than 2,200 psi at a rate of 1.1 ml/min. Fluorescent emission of PLP was measured at 478 nm while the excitation wavelength was 367 nm. The PLP peak on the fluorescent emission output was detected by its known retention time (~3.4 min) through this particular column using Waters Millennium 32 chromatography manager. Fluorescent emission was enhanced via a postcolumn bisulfite buffer containing 4% NaOH in HPLC water, which was filtered and degassed before use. Standards were made from a stock solution containing 10 mg of PLP in 1 liter of HPLC water and diluted to 0, 0.5, 1, 5, 10, and 20 ng/100 µl of PLP. Fluorescence emission was found to be linear over this concentration range.

SR Ca2+ uptake. The SR vesicles Ca2+ uptake rates were measured following the ratiometric method of Williams et al. (41) using a Jasco CAF-110 intracellular ion analyzer with a 75-W xenon high-pressure lamp. Extravesicular Ca2+ was measured fluorometrically using the Ca2+ indicator fura 2 (excitation wavelengths 340 and 380 nm; emission wavelength 500 nm). Free Ca2+ concentration ([Ca2+]free) was calculated by measuring the ratio (R) of the emissions at 500 nm from both excitation wavelengths and using the following equation: [Ca2+]free = Kd · beta  · (R - Rmin)/(R - Rmax), where the fura 2 dissociation constant (Kd) was 70 nM; Rmin and Rmax are the R values measured in the Ca2+ uptake buffer in a Ca2+-free medium and a Ca2+-saturated medium, respectively; and beta  is the fluorescence ratio of EGTA- and Ca2+-supplemented buffers at 340 nm (18). Fifty micrograms of SR protein were added to 1 ml of Ca2+ uptake buffer containing the following: 100 mM KCl, 20 mM HEPES, 1 mM MgCl2, 5 mM potassium oxalate, 1 mM ATP, and 2 µM fura 2. The solution was allowed to equilibrate at 37°C until the fluorescence ratio remained constant. CaCl2 (0.1 µmol/mg) was then added to initiate uptake. This resulted in a free Ca2+ concentration of ~1 µM. Fluorescence ratio values were sampled at 2 Hz using Labtech Notebook Pro and saved to disk for later analysis. Ca2+ uptake rates were computed from the steepest negative slopes after the addition of CaCl2. Figure 1 shows typical tracings of rested and fatigued samples. Our preliminary experiments showed that adding additional CaCl2 or increasing MgATP concentration did not result in noticeably higher Ca2+ uptake rates and, in some cases, slightly reduced the rate. Furthermore, we found that using antipyrylazo III as the extravesicular Ca2+ indicator resulted in uptake rates were nearly identical to those obtained using fura 2. 


View larger version (8K):
[in this window]
[in a new window]
  		Fig. 1.   Measurement of sarcoplasmic reticulum (SR) Ca2+ uptake in rested and fatigued samples. Samples were obtained from contralateral gastrocnemius and plantaris muscles. Arrow, addition of 0.1 µmol/mg CaCl2.

Statistics. Paired t-tests were used for comparisons between rested and fatigued conditions. Significance was set at the P <=  0.05 level of confidence.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Effect of stimulation on force production. Force production by the gastrocnemius and plantaris muscles was measured at the beginning and end of the 15-min stimulation protocol. Typically, force dropped off rapidly over the first minutes of stimulation, followed by a slow and steady decline over the remaining time. At the end of the 15-min protocol, muscle force production was reduced to 26.5 ± 4.0% of initial force.

Whole muscle and SR glycogen. Glycogen concentrations for whole muscle and SR preparations are shown in Fig. 2. Whole muscle glycogen concentration in the fatigued muscles was significantly decreased to 23% of rested, whereas glycogen associated with the SR was significantly reduced to 5% of rested (Fig. 2). Our measurement of SR glycogen proved to be very reproducible in that error between duplicate measurements was 3.40 ± 0.70%. However, SR glycogen in the rested samples was quite variable between animals, ranging from 117 to 652 µg/mg SR protein. This variability was much smaller in the fatigued samples (17-29 µg/mg SR protein).


View larger version (13K):
[in this window]
[in a new window]
  		Fig. 2.   Changes in whole muscle (left) and SR glycogen (right) after fatiguing stimulation of the gastrocnemius and plantaris muscles. Values are means ± SE; n = 6 muscles for each measurement. *P < 0.05 between conditions.

GP and Ca2+-ATPase associated with the SR. Optical density of the GP and Ca2+-ATPase determined via SDS-PAGE stained with Coomassie blue is shown in Fig. 3. As can be seen, there is a marked reduction in the size of the band associated with GP and little or no change in that of the Ca2+-ATPase. Figure 3, bottom, shows the mean optical densities of these bands. GP content in the rested samples was significantly reduced to 21% of rested, whereas the Ca2+-ATPase was not different between conditions. Also, the ratio of GP and Ca2+-ATPase was substantially reduced in the fatigued samples. Western blotting with an antibody to the fast Ca2+-ATPase isoform confirmed that ATPase content was not different between conditions (data not shown).


View larger version (36K):
[in this window]
[in a new window]
  		Fig. 3.   Glycogen phosphorylase (GP) and Ca2+-ATPase content in rested and fatigued samples. Top: Coomassie blue-stained 4% SDS-PAGE showing the location of the GP and Ca2+-ATPase proteins. Each lane was loaded with 10 µg of total protein and run at 45 mA until the tracking dye ran off of the gel. MW, molecular weight markers (116 and 97); R, rested; F, fatigued. Bottom: GP and Ca2+-ATPase content. Values are means ± SE; n = 8 muscles for each measurement. *P < 0.05 between conditions.

GP activity and PLP content of the rested and fatigued samples are shown in Fig. 4. Activity of GP in the direction of glycogen breakdown was significantly reduced to 4% of rested. PLP content was also reduced to a similar extent, 3% of rested. Within-sample variability for GP and PLP was 3.40 ± 1.37 and 1.74 ± 0.52%, respectively.


View larger version (12K):
[in this window]
[in a new window]
  		Fig. 4.   Changes in GP activity (left) and pyridoxal 5'-phosphate (PLP) content (right) in rested and fatigued samples. Values are means ± SE; n = 6 muscles for each measurement. G1P, glucose 1-phosphate. *P < 0.05 between conditions.

SR Ca2+ uptake. Peak Ca2+ uptake rates in the fatigued samples were significantly reduced to 81% of rested, from 2.99 ± 0.21 to 2.42 ± 0.205 µmol · mg SR protein-1 · min-1 (n = 6; P <=  0.05). Similar differences were found when the uptake rates were normalized by the SDS-PAGE optical density of the Ca2+-ATPase.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

In this study, the effects of prolonged muscle contraction on glycogen, GP, and maximum Ca2+ uptake rates associated with the SR were investigated. The sciatic nerve from one leg of animals was repetitively stimulated to fatigue the gastrocnemius and plantaris muscles. Significant reductions were found for glycogen, GP, and Ca2+ uptake rate associated with the SR between control and fatigued muscles.

Whole muscle and SR glycogen. Glycogen associated with the SR in control samples was determined to be ~400 µg/mg SR protein. Previous investigations by Cuenda et al. (9) and Entman et al. (13) reported SR glycogen concentrations to be 32 (rat skeletal muscle) and 300-700 (dog cardiac muscle) µg/mg SR protein, respectively. The discrepancy between the reported SR glycogen concentration in the present investigation and that from others may be due to the different assays used to assess SR glycogen and/or to the SR sample storage conditions. First, both Cuenda et al. (9) and Entman et al. (13) used an assay that utilized sulfuric acid to hydrolyze the glycogen and phenol for colorization, followed by measuring the absorbance change. The present investigation used amyloglucosidase to hydrolyze the glycogen and fluorometric measurement of beta -NADPH in an enzyme-linked system. We have found that the latter assay is more sensitive and more reliable at low glycogen (Lees and Williams, unpublished observations). Also, in the phenol-sulfuric acid assay, glycogen is measured in the acid precipitate. There is also a pool of acid-soluble glycogen that is typically not measured (36, 42). It is not clear whether Cuenda et al. (9) and Entman et al. (13) measured this latter pool. Second, it appears that the earlier investigators had sucrose present in the SR storage buffer. Because sucrose is measured as if it were glycogen in phenol-sulfuric acid assay, it must be subtracted from the total glycogen measurement. Because the amount of glycogen associated with the SR is extremely small compared with the sucrose content (~1%), this can be problematic. It is unclear whether sucrose had to be subtracted from their analyses or whether some other storage method was used on samples analyzed for glycogen content.

Whole muscle glycogen concentration was significantly decreased to 23% of control as a result of in situ stimulation. This reduction is similar to that reported by other investigators who used varied models to study glycogen depletion during muscular activity (1, 3, 6, 16, 22, 35). Glycogen associated with the SR was significantly reduced to a greater extent to 5.1% of control. In 1989, Fridén et al. (15) used periodic acid-thiosemicarbazide-silver proteinate staining of ultrathin muscle sections from human vastus lateralis. The staining revealed a compartmental distribution of glycogen within the muscle cell. After exercise (60 8-s sprint cycling bouts), glycogen depletion was particularly noticeable at the N2-line, or the lateral portion of the I band. Although the methods used by Fridén et al. could not directly verify that glycogen was associated with the SR and depleted with exercise, it is important to note that the SR is present at the I band where deposits of glycogen were located (34, 33). Thus our results extend those of Fridén et al. to confirm that SR glycogen is markedly reduced with muscular activity.

GP associated with the SR. GP has been shown to be associated with the SR (14, 38). Specifically, GP associated with the SR is >95% in its b (inactive, dephosphorylated) form (GPb) (11). Interestingly, GP appears to be associated to the SR via its binding to the glycogen particles (9, 14, 28, 38). Binding of glycogen particles to the SR membrane may be achieved by the hydrophobic tail of the glycogen-associated form of protein phosphatase 1 (20, 21). There is evidence to suggest that glycogenolysis of SR glycogen results in the release of glycogen phosphorylase. Cuenda et al. (9) showed that preparing SR from animals that were starved for 48 h resulted in a two- to fourfold decrease in GP activity and content. Similarly, it was shown that amylase digestion of endogenous glycogen resulted in 95% depletion of GP (14).

GP associated with the SR was measured using three different methods: optical density of the band corresponding the 97,400-molecular-weight marker on a SDS-PAGE, PLP content of the SR vesicles, and glycogen phosphorylase activity associated with the SR vesicles. Multiple approaches were used because each has individual drawbacks. Western blot analysis was not practical from the SDS-PAGE because an antibody for the skeletal muscle specific isoform of GP was not available. This reduces the confidence in measuring GP using SDS-PAGE because other proteins could be present with similar molecular weight. By removing and measuring PLP, the GP content could be determined because over 95% of PLP in skeletal muscle is bound to this enzyme (4). However, absence of PLP does not necessarily mean that GP is not present, but GP without PLP is inactive. Finally, measurement of GP activity does not allow for quantification of enzyme content, because we could not reliably determine whether the specific activity of the enzyme has been altered. Taken together, however, all three of the measurement techniques strongly suggest that the GP content of the SR is reduced with exercise. It seems unlikely that all three methods would falsely indicate similar changes in GP associated with the SR.

GP activity of the SR was significantly reduced to 4% of control. These data agree with the results from Cuenda et al. (9) in that starvation reduced GP activity. Also, Entman et al. (14) found that GP was reduced by 95% after alpha -amylase digestion of endogenous glycogen. If, in fact, GP is bound to the SR via its association with glycogen, a treatment that causes glycogen depletion should result in GP dissociation from the SR as well. Release of GP from the SR due to glycogen breakdown is compounded by the fact that GP is primarily bound to the SR as the GPb form, whereas muscle contraction mediates phosphorylation (activation) of this enzyme, which, in turn, reduces its affinity for the SR (8). Similarly, PLP content of the SR was significantly reduced to 3% of control. In contrast, optical density of GP determined via SDS-PAGE stained with Coomassie blue was only reduced to 21% of control. There may be a reasonable explanation for this discrepancy. Hirata et al. (19) have recently identified another 97,000-molecular-weight protein other than glycogen phosphorylase. This protein was reported to be involved in SR Ca2+ release; therefore, it seems likely that it may be present in the SR fraction used in the present investigation.

SR Ca2+ uptake. Decreases in SR glycogen and GP were accompanied by a reduction in maximum Ca2+ uptake rates to 81% of control. Reduced SR Ca2+ uptake rate during fatigue has been previously reported by others (for review see Ref. 40). At present, it is not clear whether there is a direct effect of glycogen and GP reduction on SR function. In our isolated SR system, it is unlikely that metabolic changes resulting from the loss of glycogen and GP affect uptake because adequate ATP is added and metabolites have been removed. Cuenda et al. (8) found that GPb status of the SR affected the conformation of the SR Ca2+-ATPase. These investigators found that, as GPb content increased, the binding of FITC to the SR proteins also increased. Because FITC binds to a specific lysine residue located in the ATP binding site of ATPase enzymes (29), it was concluded that the SR Ca2+-ATPase shifted toward an ATP binding conformation. This suggests that decreased GP concentration may directly reduce the rate of ATP hydrolysis by the SR Ca2+-ATPase and Ca2+ transport into the SR. However, additional work is needed to substantiate this notion.

Within the intact cell, there is the possibility that decreased glycogen and GP associated with the SR could alter Ca2+ handling during skeletal muscle fatigue via some metabolic effect. It has been shown that SR Ca2+ uptake can be supported solely through the enzymatic breakdown of glycogen by GP (10, 30, 31). Also, there is evidence that glycolytic, glycogenolytic, and Ca2+-accumulating enzymes are associated with the SR in close proximity to the Ca2+-ATPase (13, 14, 43). Decreased local ATP concentrations, due to glycogen depletion of the SR, may directly affect maximum SR Ca2+ uptake rates. It has been proposed that a local increase in the ADP/ATP ratio in the triads may be responsible for decreased SR Ca2+ release found in fatigue (24). Some ATPase activity, required for SR Ca2+ release, utilizes ATP in the microenvironment of the triad. This ATP is not in equilibrium with the bulk cellular ATP, probably because the Mg2+-ATP complex is large and negatively charged; therefore, a local ATP regenerating system is necessary for the maintenance of the ADP/ATP ratio (for a review see Ref. 39). On the other hand, experiments using caged ATP have shown no significant improvement on the slow decline of intracellular Ca2+ concentration late in fatigue (2). Therefore, whether or not SR glycogen depletion affects SR Ca2+ uptake, in vivo, via changes in local ATP concentration remain to be seen.

Summary. Tetanic contractions elicited once per second for 15 min reduced glycogen content, GP content, and SR Ca2+ uptake associated with the SR. GP activity and PLP content of the SR showed similar decreases. Loss of optical density of the band containing GP using SDS-PAGE was not as dramatic as seen with the other two measures; however, this may be explained by the presence of another protein (19). Reduced glycogen and GP may be involved, either directly or indirectly, in a mechanism that causes decreased SR Ca2+ uptake normally found in fatigue.


    ACKNOWLEDGEMENTS

The authors thank Drs. Robert J. Talmadge and William Newton for insightful comments and Janet Rinehart for assistant with the PLP assay.


    FOOTNOTES

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

Present address of E. E. Spangenburg: Veterinary Biomedical Sciences, University of Missouri, E102 Veterinary Medical Bldg., Columbia, MO 65211.

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

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.

Received 3 January 2001; accepted in final form 24 May 2001.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

1.   Ahlborg, B, Bergström J, Ekelund LG, and Hultman E. Muscle glycogen and muscle electrolytes during prolonged physical activity. Acta Physiol Scand 70: 129-142, 1967.

2.   Allen, DG, Lännergren J, and Westerblad H. The role of ATP in the regulation if intracellular Ca2+ release in single fibres of mouse skeletal muscle. J Physiol (Lond) 498: 587-600, 1997[ISI].

3.   Bergström, J, Hermansen L, Hultman E, and Saltin B. Diet, muscle glycogen and physical performance. Acta Physiol Scand 71: 140-150, 1967[ISI][Medline].

4.   Butler, PE, Cookson EJ, and Beynon RJ. The turnover of skeletal muscle glycogen phosphorylase studied using the cofactor, pyridoxal phosphate, as a specific label. Biochim Biophys Acta 847: 316-323, 1985[Medline].

5.   Chi, MM, Hintz CS, Coyle EF, Martin WH, III, Ivy JL, Nemeth PM, Holloszy JO, and Lowry OH. Effects of detraining on enzymes of energy metabolism in individual human muscle fibers. Am J Physiol Cell Physiol 244: C176-C287, 1983.

6.   Chin, ER, and Green HJ. Effects of tissue fractionation on exercised-induced alterations in SR function in rat gastrocnemius muscle. J Appl Physiol 80: 940-948, 1996[Abstract/Free Full Text].

7.   Cordes, E, and Jencks W. Semicarbazone formation from pyridoxal, pyridoxal phosphate, and their schiff bases. Biochemistry 1: 773-778, 1962.

8.   Cuenda, A, Centeno F, and Gutiérrez-Merino C. Modulation by phosphorylation of glycogen phosphorylase-sarcoplasmic reticulum interaction. FEBS Lett 283: 273-276, 1991[ISI][Medline].

9.   Cuenda, A, Henao F, Nogues M, and Gutiérrez-Merino C. Quantification and removal of glycogen phosphorylase and other enzymes associated with sarcoplasmic reticulum membrane preparations. Biochim Biophys Acta 1194: 35-43, 1994[Medline].

10.   Cuenda, A, Nogues M, Gutiérrez-Merino C, and de Meis L. Glycogen phosphorylysis can form a metabolic shuttle to support Ca2+ uptake by sarcoplasmic reticulum membranes in skeletal muscle. Biochem Biophys Res Commun 196: 1127-1132, 1993[ISI][Medline].

11.   Cuenda, A, Nogues M, Henao F, and Gutiérrez-Merino C. Interaction between glycogen phosphorylase and sarcoplasmic reticulum membranes and its functional implications. J Biol Chem 273: 11998-12004, 1995.

12.   Drexler, H. Skeletal muscle failure in heart failure. Circulation 85: 1621-1623, 1992[ISI][Medline].

13.   Entman, M, Kaniike K, Goldstein M, Nelson T, Bornet E, Futch T, and Schwartz A. Association of glycogenolysis with cardiac sarcoplasmic reticulum. J Biol Chem 251: 3140-3146, 1976[Abstract/Free Full Text].

14.   Entman, M, Keslensky S, Chu A, and Van Winkle B. The sarcoplasmic reticulum-glycogenolytic complex in mammalian fast twitch skeletal muscle. J Biol Chem 255: 6245-6252, 1980[Abstract/Free Full Text].

15.   Fridén, J, Seger J, and Ekblom B. Topographical localization of muscle glycogen: an untrahistochemical study in the human vastus lateralis. Acta Physiol Scand 125: 381-391, 1989.

16.   Galbo, H, Holst JJ, Chu SS, and Van Winkle WB. The effect of different diets and of insulin on the hormonal response to prolonged exercise. Acta Physiol Scand 107: 19-32, 1979[ISI][Medline].

17.   Gregory, J, III. Determination of pyridoxal 5'-phosphate as the semicarbazone derivative using high-performance liquid chromatography. Anal Biochem 201: 374-379, 1980.

18.   Grynkiewicz, G, Poenie M, and Tsien RY. A new generation of Ca2+ indicators with greatly improved fluorescence properties. J Biol Chem 260: 3440-3450, 1985[Abstract/Free Full Text].

19.   Hirata, Y, Nakahata N, and Ohizumi Y. Identification of a 97-kDa mastoparan-binding protein involving in Ca2+ release from the skeletal muscle sarcoplasmic reticulum. Mol Pharmacol 57: 1235-1243, 2000[Abstract/Free Full Text].

20.   Hubbard, MJ, and Cohen P. The glycogen-binding subunit of protein phosphatase-1G from rabbit skeletal muscle. Further characterization of its structure and glycogen-binding properties. Eur J Biochem 180: 457-465, 1989[ISI][Medline].

21.   Hubbard, MJ, Dent P, Smythe C, and Cohen P. Targetting of protein phosphatase 1 to the sarcoplasmic reticulum of rabbit skeletal muscle by a protein that is very similar or identical to the G subunit that directs the enzyme to glycogen. Eur J Biochem 30: 243-249, 1990.

22.   Kelso, T, Hodgson D, Visscher A, and Gollnick P. Some properties of different skeletal muscle fiber types: comparison of reference bases. J Appl Physiol 62: 1436-1441, 1987[Abstract/Free Full Text].

23.   Keppler, D, and Decker K. Methods of Enzymatic Analysis (3rd ed.). Weinheim, Germany: Verlag Chemie, 1984.

24.   Korge, P, and Campbell KB. Local ATP regeneration is important for sarcoplasmic reticulum Ca2+ pump function. Am J Physiol Cell Physiol 267: C357-C366, 1994[Abstract/Free Full Text].

25.   Laemmli, U. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227: 680-685, 1970[Medline].

26.   Lees, SJ, Spangenburg EE, and Williams JH. Measurement of glycogen associated with skeletal muscle sarcoplasmic reticulum (Abstract). FASEB J 14: A284, 2000.

27.   Mahuren, J, and Coburn S. B6 vitamers: cation exchange HPLC. J Nutr Biochem 1: 659-663, 1990.

28.   Meyer, F, Heilmeyer L, Haschke R, and Fisher E. Control of phosphorylase activity in a muscle glycogen particle. J Biol Chem 245: 6642-6648, 1970[Abstract/Free Full Text].

29.   Mitchinson, C, Wilderspin A, Trinnaman B, and Green M. Identification of a labelled peptide after stoichiometric reaction of fluorescein isothiocyanate with the Ca2+ dependent adenosine triphosphatase of sarcoplasmic reticulum. FEBS Lett 146: 87-92, 1984.

30.   Montero-Lomeli, M, and de Meis L. Glucose-6-phosphate and hexokinase can be used as an ATP-regenerating system by the Ca2+-ATPase of sarcoplasmic reticulum. J Biol Chem 267: 1829-1833, 1992[Abstract/Free Full Text].

31.   Nogues, M, Cuenda A, Henao F, and Gutiérrez-Merino C. Ca2+ uptake coupled to glycogen phosphorolysis in the glycogenolytic-sarcoplasmic reticulum complex from rat sketetal muscle. Z Naturforsch [C] 51: 591-598, 1996.

32.   Perreault, CL, Gonzalez-Serratos H, Litwin S, Sun X, Franzini-Armstrong C, and Morgan JP. Alterations in contractility and intracellular Ca2+ transients in isolated bundles of skeletal muscle fibers from rats with chronic heart failure. Circ Res 73: 405-412, 1993[Abstract].

33.   Pette, D. Some aspects of supramolecular organization of glycogenolytic and glycolytic enzymes in muscle. Acta Histochem Suppl 14: 47-68, 1975[Medline].

34.   Sigel, P, and Pette D. Intracellular localization of glycogenolytic and glycolytic enzymes in white and red rabbit skeletal muscle. J Histochem Cytochem 17: 225-237, 1969[Abstract].

35.   Stephenson, DG, Nguyen LT, and Stephenson GMM Glycogen content and excitation-contraction coupling in mechanically skinned muscle fibres of the cane toad. J Physiol (Lond) 519: 177-187, 1999[Abstract/Free Full Text].

36.   Stetten, D, and Stetten M. Glycogen metabolism. Physiol Rev 40: 505-537, 1960[Free Full Text].

37.   Ubbink, J, Serfontein W, and De Villiers L. Stability of pyridoxal-5-phosphate semicarbazone: applications in plasma vitamin B6 analysis and population surveys of vitamin B6 nutritional status. J Chromatogr 342: 277-284, 1985[ISI][Medline].

38.   Wanson, J, and Drochmans P. Binding of phosphorylase, phosphorylase kinase, and primer complexes to the sarcovesicles of rabbit skeletal muscle. J Cell Biol 54: 206-224, 1972[Abstract/Free Full Text].

39.   Westerblad, H, Allen DG, Bruton JD, Andrade FH, and Lännergren J. Mechanisms underlying the reduction of isometric force in skeletal muscle fatigue. Acta Physiol Scand 162: 253-260, 1998[ISI][Medline].

40.   Williams, JH, and Klug GA. Calcium exchange hypothesis of skeletal muscle fatigue: a brief review. Muscle Nerve 18: 421-434, 1995[ISI][Medline].

41.   Williams, JH, Ward CW, Spangenburg EE, and 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].

42.   Willstaetter, R, and Rhodewald M. Protein binding of physiologically important substances. The condition of glycogen in liver, muscle and leukocytes. Z Phys Chem 225: 103-124, 1934.

43.   Xu, K, Zweier J, and Becker L. Functional coupling between glycolysis and sarcoplasmic reticulum Ca2+ transport. Circ Res 77: 88-97, 1995[Abstract/Free Full Text].

J APPL PHYSIOL 91(4):1638-1644
8750-7587/01 $5.00 Copyright © 2001 the American Physiological Society


This article has been cited by other articles:


Home page
 Exp PhysiolHome page
S. Matsunaga, T. Mishima, T. Yamada, S. Inashima, and M. Wada
Alterations in in vitro function and protein oxidation of rat sarcoplasmic reticulum Ca2+-ATPase during recovery from high-intensity exercise
Exp Physiol, March 1, 2008; 93(3): 426 - 433.
[Abstract] [Full Text] [PDF]

Home page
 J. Appl. Physiol.Home page
T. A. Duhamel, H. J. Green, R. D. Stewart, K. P. Foley, I. C. Smith, and J. Ouyang
Muscle metabolic, SR Ca2+-cycling responses to prolonged cycling, with and without glucose supplementation
J Appl Physiol, December 1, 2007; 103(6): 1986 - 1998.
[Abstract] [Full Text] [PDF]

Home page
 J. Appl. Physiol.Home page
R. D. Stewart, T. A. Duhamel, K. P. Foley, J. Ouyang, I. C. Smith, and H. J. Green
Protection of muscle membrane excitability during prolonged cycle exercise with glucose supplementation
J Appl Physiol, July 1, 2007; 103(1): 331 - 339.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Physiol. Cell Physiol.Home page
T. A. Duhamel, H. J. Green, J. G. Perco, and J. Ouyang
Comparative effects of a low-carbohydrate diet and exercise plus a low-carbohydrate diet on muscle sarcoplasmic reticulum responses in males
Am J Physiol Cell Physiol, October 1, 2006; 291(4): C607 - C617.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Physiol. Regul. Integr. Comp. Physiol.Home page
T. A. Duhamel, J. G. Perco, and H. J. Green
Manipulation of dietary carbohydrates after prolonged effort modifies muscle sarcoplasmic reticulum responses in exercising males
Am J Physiol Regulatory Integrative Comp Physiol, October 1, 2006; 291(4): R1100 - R1110.
[Abstract] [Full Text] [PDF]

Home page
 J. Appl. Physiol.Home page
T. A. Duhamel, H. J. Green, J. G. Perco, and J. Ouyang
Effects of prior exercise and a low-carbohydrate diet on muscle sarcoplasmic reticulum function during cycling in women
J Appl Physiol, September 1, 2006; 101(3): 695 - 706.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Physiol. Endocrinol. Metab.Home page
C. C. Greenberg, M. J. Jurczak, A. M. Danos, and M. J. Brady
Glycogen branches out: new perspectives on the role of glycogen metabolism in the integration of metabolic pathways
Am J Physiol Endocrinol Metab, July 1, 2006; 291(1): E1 - E8.
[Abstract] [Full Text] [PDF]

Home page
 J. Biol. Chem.Home page
R. Sacchetto, E. Bovo, A. Donella-Deana, and E. Damiani
Glycogen- and PP1c-targeting Subunit GM Is Phosphorylated at Ser48 by Sarcoplasmic Reticulum-bound Ca2+-Calmodulin Protein Kinase in Rabbit Fast Twitch Skeletal Muscle
J. Biol. Chem., February 25, 2005; 280(8): 7147 - 7155.
[Abstract] [Full Text] [PDF]

Home page
 J. Appl. Physiol.Home page
T. A. Duhamel, H. J. Green, J. G. Perco, S. D. Sandiford, and J. Ouyang
Human muscle sarcoplasmic reticulum function during submaximal exercise in normoxia and hypoxia
J Appl Physiol, July 1, 2004; 97(1): 180 - 187.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Physiol. Cell Physiol.Home page
S. J. Lees and J. H. Williams
Skeletal muscle sarcoplasmic reticulum glycogen status influences Ca2+ uptake supported by endogenously synthesized ATP
Am J Physiol Cell Physiol, January 1, 2004; 286(1): C97 - C104.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF) Free
Right arrow Submit a response
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via ISI Web of Science (20)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Lees, S. J.
Right arrow Articles by Williams, J. H.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Lees, S. J.
Right arrow Articles by Williams, J. H.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Visit Other APS Journals Online