Regulation of glycogen phosphorylase and PDH during exercise in human skeletal muscle during hypoxia

Michelle L. Parolin1, Lawrence L. Spriet2, Eric Hultman3, Melanie G. Hollidge-Horvat1, Norman L. Jones1, and George J. F. Heigenhauser1

1 Department of Medicine, McMaster University, Hamilton, Ontario L8N 3Z5; 2 Department of Human Biology and Nutritional Sciences, University of Guelph, Guelph, Ontario N1G 2W1, Canada; and 3 Department of Clinical Chemistry, Huddinge University Hospital, Karolinska Institute, S-141 Stockholm 86, Sweden


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

The present study examined the acute effects of hypoxia on the regulation of skeletal muscle metabolism at rest and during 15 min of submaximal exercise. Subjects exercised on two occasions for 15 min at 55% of their normoxic maximal oxygen uptake while breathing 11% O2 (hypoxia) or room air (normoxia). Muscle biopsies were taken at rest and after 1 and 15 min of exercise. At rest, no effects on muscle metabolism were observed in response to hypoxia. In the 1st min of exercise, glycogenolysis was significantly greater in hypoxia compared with normoxia. This small difference in glycogenolysis was associated with a tendency toward a greater concentration of substrate, free Pi, in hypoxia compared with normoxia. Pyruvate dehydrogenase activity (PDHa) was lower in hypoxia at 1 min compared with normoxia, resulting in a reduced rate of pyruvate oxidation and a greater lactate accumulation. During the last 14 min of exercise, glycogenolysis was greater in hypoxia despite a lower mole fraction of phosphorylase a. The greater glycogenolytic rate was maintained posttransformationally through significantly higher free [AMP] and [Pi]. At the end of exercise, PDHa was greater in hypoxia compared with normoxia, contributing to a greater rate of pyruvate oxidation. Because of the higher glycogenolytic rate in hypoxia, the rate of pyruvate production continued to exceed the rate of pyruvate oxidation, resulting in significant lactate accumulation in hypoxia compared with no further lactate accumulation in normoxia. Hence, the elevated lactate production associated with hypoxia at the same absolute workload could in part be explained by the effects of hypoxia on the activities of the rate-limiting enzymes, phosphorylase and PDH, which regulate the rates of pyruvate production and pyruvate oxidation, respectively.

pyruvate dehydrogenase; lactate metabolism; glycogenolysis; glycolysis; oxidative phosphorylation; phosphocreatine


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

EXERCISE AT A GIVEN SUBMAXIMAL power output during hypoxia is associated with elevated blood and muscle lactate concentrations relative to normoxia (7, 24, 36). This elevated lactate production has been attributed to increased rates of glycogenolysis and glycolysis secondary to the reduced phosphorylation potential of hypoxic tissue. Classically, this was ascribed to a reduction in O2 supply, with lactate production supplementing the reduced ATP provision from oxidative phosphorylation (for review see Ref. 39). There is general agreement that an O2 limitation may not occur under normoxic conditions at power outputs less than ~50% maximal O2 uptake (VO2 max). But when O2 supply was compromised by lowering the fraction of inspired O2 (FIO2) during submaximal exercise in humans at ~50-55% of the maximum work rate, whole body O2 uptake (VO2) (1) and VO2 across the exercising leg (41) were maintained at normoxic levels. Thus the adjustment in phosphorylation potential may simply represent an adaptation to a lower ambient PO2 in an attempt to maintain O2 utilization and ATP homeostasis by shifting to a more O2-efficient fuel in the presence of sufficient O2 delivery (27, 61).

Lactate accumulates when the rate of glycolytic pyruvate production exceeds the rate of pyruvate oxidation. As pyruvate accumulates, it is converted to lactate by the near-equilibrium enzyme lactate dehydrogenase, whose equilibrium favors lactate formation. In terms of the rate-limiting enzymes capable of exerting metabolic control, glycogen phosphorylase, which catalyzes the rate-limiting step in glycogenolysis, has the potential to set the upper limit for the rates of glycolysis and pyruvate production, whereas pyruvate dehydrogenase (PDH), which catalyzes the oxidative decarboxylation of pyruvate to acetyl-CoA, regulates the rate of entry of pyruvate into the tricarboxylic acid (TCA) cycle. Thus the balance between the activities of glycogen phosphorylase and PDH will determine lactate production under conditions of acute hypoxia, when the muscle relies more heavily on glycogen as a fuel source (see Ref. 40 for review). The data available to date on the effects of hypoxia during exercise in humans has mainly been limited to measurements of lactate accumulation.

The present comprehensive study was designed to determine the role of these rate-limiting enzymes in lactate metabolism under conditions of acute hypoxia. The transformation of phosphorylase and the activation of PDH were measured, in addition to the regulatory factors that modulate their activities. We hypothesized that acute hypoxia would result in an increased rate of glycogenolysis through posttransformational regulation of phosphorylase, exceeding the maximum flux through PDH, to account for the greater lactate accumulation observed in hypoxia. The effects of hypoxia were examined relative to normoxia at rest, at 1 min during the transition from rest to exercise, and after 15 min of submaximal exercise at the same power output.


    METHODS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Subjects

Six healthy nonsmoking male subjects volunteered to participate in the present study. Their mean (± SE) age, height, weight, and VO2 max (determined during normoxia) were 22.3 ± 1.0 yr, 176.1 ± 2.1 cm, 72.2 ± 4.2 kg, and 55.8 ± 3.0 ml · kg-1 · min-1, respectively. Three of the subjects were well trained (one was a triathlete, and the other two cycled competitively), and the other three each participated in gymnastics, hockey, or soccer 3-4 times/wk. Informed consent was obtained from each subject after a verbal and written explanation of the experimental protocol and its attendant risks. The study was approved by the McMaster University Ethics Committee.

Preexperimental Protocol

VO2 max during normoxia was determined on a cycle ergometer (Excalibur, Quinton Instruments, Seattle, WA) and metabolic cart (Quinton Q-plex 1, Quinton Instruments) by use of a continuous incremental exercise protocol. Approximately 1 wk before the study, a practice trial was conducted to familiarize subjects with the experimental protocol and hypoxic conditions of the study.

Experimental Protocol

The experimental protocol was conducted under both normoxic (control) and hypoxic conditions, on two separate occasions 1-2 wk apart. The order of the hypoxic and normoxic trials was randomized and took place at the same time of day for each subject. The subjects consumed a high-carbohydrate diet during the 48 h preceding the first trial and were asked to replicate this diet before the second trial. Subjects were also instructed to abstain from the consumption of caffeine and alcohol and to refrain from strenuous exercise for 24 h before each trial.

Before the beginning of the protocol, a venous catheter was inserted into the antecubital vein of the forearm and was maintained patent with saline. One thigh was prepared for needle biopsies of the vastus lateralis. Incisions were made through the skin to the deep fascia under local anesthesia (2% lidocaine without epinephrine) as described by Bergström (5). Four incisions were made on one thigh for the hypoxic trial, and three were made on the contralateral thigh for the normoxic trial.

The experimental protocol is summarized in Fig. 1. Before exercising, subjects rested quietly on a bed while breathing the appropriate gas mixture during a 20-min equilibration period. For both conditions, subjects breathed, with a noseclip in place, from a mouthpiece attached to a 120-liter Tissot spirometer containing either room air [normoxic control, fraction of inspired O2 (FIO2) = 21%] or a hypoxic gas mixture (FIO2 = 10.9%, balance N2). A muscle biopsy (hypoxia trial only) and venous blood sample were taken before the equilibration period. A biopsy was not taken before the equilibration period of the normoxic trial, no changes being expected with the subject at rest while breathing room air. At the end of the equilibration period, a resting venous blood sample and muscle biopsy were taken while the subject rested on the bed before he was moved to the cycle ergometer. While continuing to breathe the appropriate gas mixture, the subject cycled for 15 min at ~55% of the VO2 max determined during normoxia (154 ± 10 W). Muscle biopsies were taken after 1 and 15 min of exercise while the subject remained on the cycle ergometer. Venous blood was sampled after 5, 10, and 14 min of exercise. Throughout the protocol, expired gases were continuously sampled. O2 uptake (VO2), CO2 output (VCO2), respiratory exchange ratio (RER), ventilation (VE), tidal volume (VT), fraction of expired O2 (FEO2), fraction of expired CO2 (FECO2), and end-tidal PCO2 (PETCO2) were measured from expired gases using a Quinton metabolic cart. Data were averaged over 15-s intervals. FIO2 was also monitored to ensure that the subjects were effectively breathing the appropriate gas mixture from the Tissot spirometer. Throughout the protocol, heart rate, blood pressure, and oximetric arterial oxygen saturation were monitored.


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Fig. 1.   Experimental design. Muscle biopsies and venous blood samples were taken at times indicated by arrows and asterisks, respectively. VO2 max, maximal O2 uptake.

Blood Handling and Analysis

Venous blood samples were drawn into heparinized plastic syringes and placed on ice. A 1-ml aliquot of blood was immediately centrifuged for 1 min at 10,000 rpm. Four hundred microliters of the supernatant were added to 100 µl of 5 M NaCl, vortexed, and placed into a 56°C waterbath for 30 min to inactivate lipoprotein lipase (17). This sample was stored at -20°C for the determination of plasma free fatty acids using the Wako NEFA C assay kit (WAKO Chemicals, Osaka, Japan) at a later time. A second 400-µl aliquot of blood was deproteinized in 800 µl of 0.5 M HClO4, vortexed, and centrifuged at 10,000 rpm for 1 min. The supernatant was stored at -20°C and was later analyzed for glucose, lactate, and glycerol, as described by Bergmeyer (4).

Muscle Handling and Analysis

Muscle biopsies were immediately frozen in liquid N2, removed from the needle while frozen, and stored in liquid N2 until analyzed. A 5- to 10-mg piece of muscle was chipped from each biopsy under liquid N2 and dissected free of blood and connective tissue for the determination of the active form of PDH (PDHa), as described by Putman et al. (50). The remaining muscle was freeze-dried, dissected free of blood, connective tissue, and fat, and stored dry at -70°C for subsequent analysis.

One aliquot of powdered muscle was used for the determination of glycogen phosphorylase activity, as described by Young et al. (63). Briefly, 3-4 mg of powdered muscle were homogenized at -25°C in 200 µl of buffer containing 100 mM Tris, 60% glycerol, 50 mM KF, and 10 mM EDTA (pH 7.5). Homogenates were then diluted with 800 µl of the above buffer without glycerol and homogenized further at 0°C. Total (a + b) phosphorylase activity (in the presence of 3 mM AMP) and phosphorylase a activity (in the absence of AMP) were measured by following the production of glucose 1-phosphate spectrophotometrically at 30°C. Maximal velocity and the mole fraction of phosphorylase a and a + b were calculated from the measured activities, as described by Chasiotis et al. (12). To minimize the total number of biopsies in the study, resting measurements of phosphorylase a were not obtained. An accurate estimate of the active form of phosphorylase at rest requires that biopsies be frozen in liquid N2 after a 30-s delay at room temperature (54) and would have necessitated two additional biopsies. Previous measurements have given values of ~10% for the mole fraction of phosphorylase a at rest (13, 46).

A second aliquot of powdered muscle was assayed enzymatically for glycogen, as described by Harris et al. (25). The remaining dry powdered muscle was extracted in a solution of 0.5 M HClO4 and 1 mM EDTA and neutralized with 2.2 M KHCO3. These extracts were assayed for ATP, phosphocreatine (PCr), creatine, pyruvate, lactate, glucose, glucose 1-phosphate (G-1-P), glucose 6-phosphate (G-6-P), fructose 6-phosphate (F-6-P), and glycerol 3-phosphate (Gly-3-P), as described by Bergmeyer (4), and for acetyl-CoA, CoA, acetylcarnitine, and carnitine as described by Cederblad et al. (8). Muscle metabolites and enzyme activities were corrected to the highest total creatine content for each subject.

Calculations

Arterial PCO2 (PaCO2) was estimated from PETCO2 and VT according to Jones et al. (35). Intracellular [H+] was calculated from muscle [lactate] and [pyruvate] according to Sahlin et al. (57). The concentrations of free ADP and free AMP were calculated from the near-equilibrium reactions of creatine kinase and adenylate kinase, respectively (16). The concentration of free inorganic phosphate ([Pi]) was calculated as the difference between resting and exercise [PCr], less the accumulation of G-6-P, F-6-P, and Gly-3-P, plus the assumed resting concentration of 10.8 mmol/kg dry wt (16).

Pyruvate oxidation was estimated by the flux through PDH. Because previous studies have shown that flux through PDH is equivalent to its level of activity (31, 49), PDHa was used as a measure of pyruvate oxidation and converted to units of millimoles per kilogram dry weight per minute, with the assumption of a wet-to-dry muscle ratio of 4:1 at rest and 4.5:1 during exercise (48). The rate of pyruvate production was calculated from the accumulation of muscle lactate and pyruvate and blood lactate, plus the flux of pyruvate through PDHa. Lactate production was calculated from the accumulation of muscle and blood lactate. The distribution volume of blood lactate was assumed to be 0.64 × body weight (3). This calculation does not account for oxidation of lactate by other tissues and would result in a slight underestimation of lactate accumulation (3).

Estimates of the glycogenolytic rate during the 1st min of exercise were derived from the accumulation of muscle G-6-P, F-6-P, Gly-3-P, pyruvate, lactate and blood lactate, plus the flux of pyruvate through PDHa from rest to 1 min of exercise. The glycogenolytic rate during the subsequent 14 min of exercise was calculated from total glycogen utilization during 15 min of exercise, minus the estimated glycogen utilization in the 1st min, divided by time.

The rate of ATP turnover from PCr was calculated from the breakdown of PCr, whereas the rate of ATP turnover from glycolysis was calculated from the accumulation of muscle and blood lactate and the flux of pyruvate through PDHa. The rate of ATP turnover from oxidative phosphorylation originating from carbohydrate sources was calculated from total acetyl-CoA production as the area under the PDHa curves; 1 mmol of acetyl-CoA was equal to 15 mmol ATP.

Statistical Analysis

All data are presented as means ± SE. Data were analyzed by a two-way ANOVA with repeated measures over time. When a significant F ratio was found, the Newman-Keuls post hoc test was used to compare the means. Cardiorespiratory parameters and net changes in [glycogen] during normoxia and hypoxia were compared using a two-tailed paired dependent-samples t-test. Pre- and postequilibrium PDHa and metabolite concentrations were also compared during hypoxia by use of a two-tailed paired dependent-samples t-test. Results were considered significant at P < 0.05.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Cardiorespiratory Parameters

At rest, none of the cardiorespiratory parameters were changed by hypoxia, with the exception of a higher heart rate and lower FEO2 and PETCO2 compared with normoxia (Table 1). During the last 5 min of exercise, VO2 and VCO2 were unaffected by hypoxia, whereas RER, VE, VT, and heart rate were higher, and FEO2, FECO2, PETCO2, and PaCO2 were lower during the hypoxic condition compared with normoxia. All parameters changed significantly from rest during exercise under both conditions with the exceptions of RER, PETCO2, and PaCO2, which did not change during normoxia.

                              
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Table 1.   Cardiorespiratory parameters at rest and during exercise in normoxia and hypoxia

Equilibration Period

During the 20-min equilibration period, hypoxia had no effect on resting levels of PDHa and muscle and blood metabolite contents. Consequently, after the 20-min equilibration period, these values were similar to those in the normoxic condition.

Glycogen Phosphorylase

After 1 min of exercise, the mole fraction of phosphorylase a was unaffected by hypoxia. After 15 min of exercise, the mole fraction of phosphorylase a was unchanged in normoxia but decreased significantly during hypoxia and was lower compared with normoxia (Fig. 2).


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Fig. 2.   Muscle phosphorylase a mole fraction during exercise in normoxia and hypoxia. Dagger  Significantly different from 1 min. + Significantly different from normoxia.

PDHa

After 1 min of exercise, PDHa increased significantly during both conditions but was significantly lower during hypoxia (Fig. 3). After 15 min of exercise, PDHa did not change during normoxia but increased significantly during hypoxia and was significantly greater compared with normoxia.


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Fig. 3.   Muscle pyruvate dehydrogenase in its active form (PDHa) at rest and during exercise in normoxia and hypoxia. * Significantly different from rest. + Significantly different from normoxia. Dagger  Significantly different from 1 min.

Muscle Metabolites

Glycogen. Resting muscle [glycogen] was not significantly different between conditions (456 ± 42 vs. 503 ± 44, normoxia vs. hypoxia), but at the end of exercise [glycogen] had decreased significantly during both conditions and was significantly lower during the hypoxic condition compared with normoxia (381 ± 65 vs. 277 ± 42 mmol/kg dry wt). Total glycogen utilization was significantly greater in hypoxia compared with normoxia (226 ± 42 vs. 75 ± 30 mmol/kg dry wt).

Glycolytic intermediates. In normoxia, muscle [glucose] did not change with exercise. After 1 min of exercise, [glucose] was similar in both conditions but increased significantly after 15 min of exercise in hypoxia and was higher compared with normoxia (Table 2). [G-1-P] was unaltered by exercise and was similar between conditions (Table 2). [G-6-P] and [F-6-P] increased significantly after 1 min of exercise and were similar under both conditions (Table 2). After 15 min of exercise during normoxia, [G-6-P] and [F-6-P] declined to levels not different from rest. During hypoxia, [G-6-P] and [F-6-P] increased significantly from 1 min to 15 min of exercise and were greater compared with normoxia. In normoxia, [Gly-3-P] was unaltered by exercise, whereas during hypoxia, [Gly-3-P] increased progressively with exercise and was higher compared with normoxia at both time points (Table 2). In normoxia, [pyruvate] was unaltered by exercise, whereas in hypoxia, [pyruvate] increased significantly after 15 min of exercise and was significantly greater than normoxia at 1 and 15 min (Table 2).

                              
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Table 2.   Muscle contents of glycolytic intermediates at rest and during exercise in normoxia and hypoxia

Lactate and H+. In normoxia, muscle [lactate] did not change significantly during exercise. In contrast, in hypoxia, [lactate] increased progressively with exercise and was always greater compared with normoxia (Fig. 4). In normoxia, estimated intramuscular [H+] was unaffected by exercise. After 1 min of exercise, [H+] was similar in both conditions but increased significantly after 15 min of exercise in response to hypoxia and was higher compared with normoxia (Table 2).


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Fig. 4.   Muscle lactate concentration at rest and during exercise in normoxia and hypoxia. Symbols are as in Fig. 3.

High energy phosphates. [ATP] was unaltered by exercise and was similar between conditions (Table 3). After 1 min of exercise, [PCr] decreased significantly and was similar under both conditions (Fig. 5). [PCr] was unchanged after 15 min of exercise during normoxia but decreased further during the hypoxic condition and was significantly lower compared with normoxia. Free [ADP] and free [AMP] were similar during both conditions after 1 min of exercise. Thereafter, there was no change in normoxia, but both free [ADP] and free [AMP] increased significantly after 15 min of exercise in hypoxia and were higher compared with normoxia (Table 3). The accumulation of free Pi was reciprocal to PCr degradation and was similar at 1 min in both conditions. Free [Pi] did not change further after 15 min of exercise in normoxia but did increase further after 15 min during hypoxia and was greater compared with normoxia (Table 3).

                              
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Table 3.   Muscle contents of high energy phosphates at rest and during exercise in normoxia and hypoxia



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Fig. 5.   Muscle phosphocreatine (PCr) concentration at rest and during exercise in normoxia and hypoxia. Symbols are as in Fig. 3.

CoA, carnitine, and acetylated forms. After 1 min of exercise in hypoxia, [acetyl-CoA] increased significantly and increased further after 15 min (Table 4). During normoxia, [acetyl-CoA] increased significantly from rest only after 15 min of exercise. There were no significant differences in [acetyl-CoA] at any time between conditions. Free [CoA] tended to decrease with exercise but was not different between conditions (Table 4). The ratio of acetyl-CoA to CoA increased significantly from rest after 1 min of exercise during hypoxia and after 15 min of exercise during normoxia, but it was similar under both conditions at all time points. In normoxia, [acetylcarnitine] increased progressively with exercise at both time points (Table 4). [Acetylcarnitine] did not change from rest after 1 min of exercise in hypoxia and was similar to the normoxic condition, but it increased significantly after 15 min of exercise in hypoxia and was higher compared with normoxia. The changes in free [carnitine] were reciprocal to the changes in [acetylcarnitine], and there was a progressive decrease after 1 and 15 min of exercise in both conditions (Table 4). After 1 min of exercise, free [carnitine] was similar between conditions but was significantly lower after 15 min of exercise in hypoxia.

                              
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Table 4.   Muscle contents of CoA, carnitine, and their acetylated forms at rest and during exercise in normoxia and hypoxia

Blood Metabolites

Blood [lactate] did not change with exercise in normoxia but increased significantly at each time point during hypoxia (Table 5). Blood [glucose] did not change significantly from rest but was significantly higher during hypoxia compared with normoxia after 15 min of exercise (Table 5). Both blood [glycerol] and [free fatty acid] were unaffected by exercise and were similar between conditions (Table 5).

                              
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Table 5.   Whole blood lactate, glucose, glycerol, and plasma FFA concentrations at rest and during exercise in normoxia and hypoxia

Glycogenolysis

During the 1st min of exercise, the glycogenolytic rate was greater in hypoxia compared with normoxia. During the subsequent 14 min of exercise, glycogenolysis decreased significantly in both conditions and remained significantly greater in hypoxia compared with normoxia (Fig. 6).


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Fig. 6.   Estimated rates of muscle glycogenolysis during exercise in normoxia and hypoxia. Symbols are as in Fig. 3.

Pyruvate Production and Oxidation and Lactate Accumulation

During the 1st min and subsequent 14 min of exercise, the rate of pyruvate production was similar within each condition but was consistently greater during hypoxia (Fig. 7). The rate of pyruvate oxidation increased significantly from the 1st to the subsequent 14 min of exercise in normoxia, representing 51 and 83% of the rate of pyruvate production in the 1st min and subsequent 14 min of exercise, respectively. During the 1st min of exercise in hypoxia, the rate of pyruvate oxidation was significantly lower compared with normoxia and accounted for only 24% of the pyruvate produced. The rate of pyruvate oxidation increased significantly during the subsequent 14 min of exercise in hypoxia and was significantly greater compared with normoxia, accounting for 51% of the rate of pyruvate production (Fig. 7). The rate of lactate accumulation was similar throughout exercise in each condition but was significantly greater in hypoxia during both time periods (Fig. 7).


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Fig. 7.   A comparison of the rates of pyruvate production and oxidation and lactate accumulation during exercise in normoxia and hypoxia. Symbols are as in Fig. 3.

ATP Turnover During the 1st Min of Exercise

During the 1st min of exercise in normoxia, substrate phosphorylation accounted for 26% of the total ATP turnover, with 12 and 14% from PCr degradation and glycolysis, respectively (Fig. 8). In hypoxia, substrate phosphorylation accounted for 41% of the total ATP turnover during the 1st min of exercise, with 14 and 27% from PCr degradation and glycolysis, respectively (Fig. 8). Therefore, during the 1st min of exercise, oxidative phosphorylation contributed to the remaining 74% of the total ATP turnover in normoxia compared with only 59% in hypoxia (Fig. 8).


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Fig. 8.   ATP turnover rates from PCr degradation, glycolysis, and oxidative phosphorylation at the onset of exercise in normoxia and hypoxia.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

The present study examined the acute effects of hypoxia on the regulation of skeletal muscle metabolism at rest and during 15 min of submaximal exercise. At rest, no effects on muscle metabolism were observed in response to hypoxia. In the 1st min of exercise, during the transition from rest, glycogen utilization was greater in hypoxia despite similar phosphorylase a mole fractions and free [AMP] and [Pi] in both hypoxia and normoxia. During hypoxia, there was a delayed activation of PDH relative to normoxia and hence a greater rate of lactate accumulation in the 1st min of exercise. By the end of 15 min of exercise, glycogen utilization in hypoxia was greater than that in normoxia. The mole fraction of phosphorylase a was lower in hypoxia, and an enhanced rate of glycogenolysis was primarily due to posttransformational regulation of phosphorylase through higher free [AMP] and [Pi]. After 15 min of exercise in hypoxia, a greater PDHa contributed to a greater rate of pyruvate oxidation relative to normoxia, but lactate accumulation remained greater than in normoxia, indicating that the rates of pyruvate production remained greater than pyruvate oxidation in hypoxia. Hence, the elevated lactate production associated with hypoxia at the same absolute workload could at least in part be explained by the differing activities of the rate-limiting enzymes, phosphorylase and PDH, which regulate the rates of pyruvate production and pyruvate oxidation, respectively.

Glycogen Utilization

There is general agreement that acute hypoxia increases the reliance on carbohydrate as a fuel source during exercise. Several studies have demonstrated increases in muscle glucose uptake in humans exercising at ~40-50% VO2 max (6, 15, 38, 53), but no studies have actually demonstrated increased glycogen utilization under conditions of acute hypoxia in human skeletal muscle (24, 43, 62). Exercise at the same relative workload (85% VO2 max), but lower absolute power, during hypoxia was shown to result in a glycogen utilization similar to that observed during normoxia (62). Only two other studies have compared glycogen utilization at the same absolute workload, ~50% of the VO2 max determined during normoxia, but with differing results (24, 43). Linnarsson et al. (43) found no differences in glycogen utilization during only 4 min of exercise, although the duration of exercise may have been too brief to measure any significant differences in glycogen utilization. In contrast, Green et al. (24) observed a reduced glycogen utilization after 45 min of exercise at acute altitude, but diet was not carefully controlled in this study and the resting glycogen content after ascent to altitude was significantly lower than at sea level, which may have induced a glycogen-sparing effect (21). In the present study, identical diets were consumed for 2 days preceding each trial such that starting glycogen levels were similar for both conditions. Glycogen utilization was approximately threefold greater during exercise in hypoxia compared with normoxia. If we assume that the ATP turnover was similar under both conditions, the latter observation suggests that fat utilization was considerably reduced during hypoxia compared with normoxia.

Regulation of Glycogen Phosphorylase

The breakdown of glycogen is regulated by the rate-limiting enzyme glycogen phosphorylase. Phosphorylase itself is regulated by reversible enzymatic phosphorylation and exists in two interconvertible forms: a less active b form and a more active a form. Phosphorylase b is transformed to its a form by phosphorylase kinase, whereas dephosphorylation back to the b form is catalyzed by phosphorylase phosphatase. The transformation of phosphorylase is regulated at the hormonal level by cAMP and at the contractile level by Ca2+ release from the sarcoplasmic reticulum (10, 12). In the present study, the transformation of phosphorylase to its more active a form was similar in both conditions at 1 min of exercise (Fig. 2). This initial transformation was probably due to similar Ca2+ release from the sarcoplasmic reticulum at the onset of exercise at the same power output.

Posttransformational regulation of phosphorylase activity and glycogenolysis occurs via the availability of substrate (Pi and glycogen) and the presence of positive (free AMP and IMP) and negative (G-6-P) allosteric modulators. The Michaelis-Menten constant (Km) values of phosphorylase a and a + b for Pi are 26.2 and 6.8 mM, respectively (12). Furthermore, the presence of 0.01 mM free AMP reduces the Km of phosphorylase a for Pi to 11.8 mM (55). Phosphorylase b is also active in the presence of high levels of free AMP (>0.01 mM) and other regulators (2). Free [AMP] was similar under both conditions after 1 min of exercise (Table 3). Similar rates of PCr degradation in the 1st min of exercise (Fig. 5) resulted in significant accumulation of free Pi (Table 3), which fell within the range of the Km for phosphorylase. These results are consistent with other studies in humans that have shown no differences in PCr degradation between hypoxia and normoxia within the first 1-2 min of exercise at the same power output, representing ~50-60% of the maximum work rate (26, 28). In the present study, despite similar transformation of phosphorylase at 1 min of exercise, the greater glycogenolytic rate in hypoxia may have been due to a very small nonsignificant increase in substrate concentration (free Pi) in the hypoxic condition (Figs. 2 and 6 and Table 3).

After 1 min of exercise, the mole fraction of phosphorylase a remained constant in normoxia (Fig. 2), consistent with a previous study that showed no change in transformation after 1 min of exercise at a similar power output (31). In contrast, in hypoxia, phosphorylase had partially reverted to its less active b form (Fig. 2), suggesting that the mechanism of activation by Ca2+ was being overridden. This inhibition of phosphorylase transformation to its a form has been shown to occur in human skeletal muscle secondary to a reduction in pH to 6.6-6.7 (11, 31, 46). The elevated [H+] has an inhibitory effect on phosphorylase kinase that reduces the transformation of phosphorylase to its a form (42). In the present study, the estimated [H+] increase secondary to lactate accumulation (257 nM, pH 6.6) after 15 min of exercise in hypoxia was ~2.5-fold greater than in normoxia (106 nM, pH 7.0) and thus may have contributed to inhibition of phosphorylase transformation (Table 2).

Although after 15 min of exercise in hypoxia the mole fraction of phosphorylase a was significantly lower than in normoxia, the glycogenolytic rate was higher (Fig. 2 and 6). Free [Pi] and [AMP] did not change from 1 to 15 min in normoxia, whereas in hypoxia, there was a greater degradation of PCr accompanied by a concomitant increase in free [Pi] and [AMP] (Table 3). The present results were similar to those observed in an isolated rat muscle preparation in which hypoxia caused a 70% reduction in glycogen content in the absence of phosphorylase transformation, due to posttransformational modulation by increases in free [Pi] and [AMP] (52). Thus, in the present study, a higher rate of glycogenolysis was achieved after 15 min of exercise in hypoxia by an enhanced stimulation of phosphorylase a, through a combination of increased availability of substrate (free Pi) and positive allosteric modulator (free AMP). On the other hand, a higher concentration of G-6-P (Table 2), a known allosteric inhibitor of phosphorylase b (20), may have exerted an inhibitory effect on glycogenolysis; without sufficient levels of free AMP, the contribution from phosphorylase b was probably minor and had little effect on the rate of glycogenolysis.

Regulation of PDH

PDH is the rate-limiting enzyme that regulates the entry of glycolytically generated pyruvate into oxidative metabolism. The PDH complex is a mitochondrial enzyme that is also regulated by reversible phosphorylation. PDH kinase catalyzes the phosphorylation of PDH with concomitant inactivation, whereas PDH phosphatase dephosphorylates the enzyme, thereby activating PDH. The activity of PDH is determined by the proportion of the complex in the active form. PDH kinase is stimulated by elevated ratios of acetyl-CoA to CoA, NADH to NAD+, and ATP to ADP; conversely, it is inhibited by pyruvate (see Refs. 51 and 60 for review). PDH phosphatase is stimulated by Ca2+ (see Refs. 51 and 60 for review).

In the transition from rest to exercise, PDHa increased to a greater degree in normoxia compared with hypoxia (Fig. 3). The initial activation of PDH was probably due to Ca2+ release from the sarcoplasmic reticulum at the onset of exercise. The Ca2+ released activates PDH phosphatase in the heart (32) and may simultaneously inactivate PDH kinase (60), thus increasing the proportion of PDH in the active form. Although the expected Ca2+ release would be similar at the same power output, other factor(s) must have been responsible for the delayed activation of PDH in hypoxia. Pyruvate is a known activator of PDH and inactivates PDH kinase (32). In normoxia, [pyruvate] did not change from rest in the 1st min of exercise (Table 2). In contrast, despite a large rise in [pyruvate] after 1 min of exercise in hypoxia, a larger rise in PDHa relative to normoxia did not occur. Other regulators of PDH, such as [acetyl-CoA] and the ratios of acetyl-CoA to CoA and ATP to free ADP, were not significantly different between conditions (Tables 3 and 4). Although NADH and NAD+ were not measured in the present study, Katz and Sahlin (36) showed no differences in whole muscle [NADH] at rest in human skeletal muscle during hypoxia, suggesting that there was not a strong inhibitory force on PDH activation during the transition from rest to exercise. However, given the uncertainty in the measurement of whole muscle NADH (37), an assumption based on the redox state of the mitochondria after 1 min of exercise cannot be made. With the feed-forward effect of the elevated [pyruvate] as a substrate and positive allosteric modulator in hypoxia, and with the assumption that the same power output elicited a similar rise in cytosolic [Ca2+] during both conditions, other factor(s) must have exerted an inhibitory effect on PDH activation at the onset of exercise in hypoxia. The respiratory alkalosis at rest in hypoxia (33) may have contributed to the delay in PDH activation at 1 min, because, like phosphorylase, the regulation of PDH is sensitive to pH; phosphorylase is inhibited by an elevated [H+] and, conversely, PDH is activated by an elevated [H+] (see below). Although the estimated pH was similar at 1 min, this calculation by Sahlin et al. (57) is based on [pyruvate] and [lactate] alone and fails to take into account changes in PCO2. Later in exercise, this effect would be lost, because [H+] increases in parallel with increasing [lactate].

In normoxia, PDH reached its greatest degree of activation within 1 min of exercise, which was maintained to the same degree throughout the remaining 14 min of exercise (Fig. 3), consistent with previous observations (31). In contrast, PDHa continued to increase during exercise in hypoxia and exceeded the activation of PDH in normoxia at the end of 15 min of exercise. In normoxia, [pyruvate] did not increase significantly from rest, having no effect on PDHa after 15 min of exercise (Table 2). In agreement with a previous study in humans during exercise at 50% VO2 max (36), [pyruvate] rose in hypoxia as a consequence of the higher rates of glycogenolysis and appeared to be near the estimated range of the inhibitory constant of PDH kinase for pyruvate of 0.5-2.0 mM (60). The greater [pyruvate] in hypoxia may have exerted a feed-forward effect on PDHa as a substrate and probably also functioned as an allosteric regulator, which stimulated activation of PDH. Under the hypoxic condition, the greater levels of pyruvate generated by the higher glycogenolytic rate had a dual effect: first, to increase the oxidation of carbohydrate sources through an increase in PDHa, and second, to increase the production of lactate through interaction with lactate dehydrogenase.

The intramuscular acidification observed at the end of exercise in hypoxia is consistent with another report (24). After 15 min of exercise in hypoxia, the estimated [H+] was ~2.5-fold greater compared with normoxia and may have played a role in PDH activation (Table 2). The only available evidence has been demonstrated in cardiac muscle. A study in perfused rat heart reported that acidosis increased PDH activation (47). Hucho et al. (32) also reported pH optima of 7.0-7.2 for bovine kidney and heart PDH kinases, and 6.7-7.1 for the PDH phosphatases, suggesting that an increase in [H+] would favor the activity of the phosphatase over the kinase and increase the transformation of PDH to its active form.

The drop in the ratio of ATP to free ADP at the end of exercise in hypoxia would have relieved the stimulation of PDH kinase and thus increased PDHa. In contrast, Katz and Sahlin (36) observed an increase in the [NADH] after 5 min of exercise at 50% VO2 max, which would have exerted a stimulatory effect on PDH kinase and reduced PDHa. The increase in [acetyl-CoA] and the ratio of acetyl-CoA to CoA after 15 min of exercise were similar between conditions, with the rise in [acetyl-CoA] buffered by significant increases in [acetylcarnitine] (Table 4). This buffering effect was most pronounced in hypoxia, when the higher PDHa was associated with a greater increase in [acetylcarnitine] (14). Therefore, even with the probability of an increase in the [NADH], in the present study it appears that the increases in Ca2+ with contraction and increases in [pyruvate] and [H+], and the drop in the ratio of ATP to free ADP in hypoxia override the inhibition that may be exerted by an elevated [NADH] in support of a greater PDHa at the end of 15 min of exercise.

O2 Availability, Cellular Energetics, and Lactate Accumulation

The question of whether muscle oxygen limitation plays a dominant role in lactate accumulation has generated considerable controversy (for review see Ref. 37). Although a number of studies have shown that lactate accumulation increases in response to exercise during acute hypoxia (18, 38, 41), more recent evidence lends support to the notion that lactate production is complex and may be related to factors other than tissue hypoxia alone (56). Using proton magnetic resonance spectroscopy to determine myoglobin saturation, Richardson et al. (56) showed that the intracellular PO2 is constant during incremental exercise to maximal work rates during both hypoxia and normoxia and is unrelated to lactate efflux. Whereas PO2 was in fact lower in hypoxia compared with normoxia, at 2.3 Torr (56), it was still well above the critical PO2 at which mitochondrial respiration is compromised (0.1-0.5 Torr) (9). Meanwhile, others have found that whole body VO2 (1) and the VO2 across the exercising leg (41) are preserved during hypoxia. The results of the present study show no difference in the whole body VO2 and are therefore in agreement with these previously mentioned studies (Table 1). Although the available evidence is conflicting, until a reliable method of measuring intracellular oxygen is developed, we must depend on circumstantial evidence to formulate a potential mechanism for lactate production.

Although O2 may not be limiting, it may play a modulatory role in oxidative metabolism. Mitochondrial oxidative phosphorylation is represented by the following equation
NADH + ½ O<SUB>2</SUB> + H<SUP>+</SUP> + 3ADP

+ 3P<SUB>i</SUB> ⇄ NAD<SUP>+</SUP> + H<SUB>2</SUB>O + 3ATP
and as such is regulated by the ratios of [NAD+] to [NADH] ([NAD+]/[NADH]) and [ATP] to [ADP][Pi] ([ATP]/[ADP][Pi]), as well as the availability of O2 (61). When one of these two ratios changes, a compensatory change occurs in the other to drive oxidative phosphorylation (61). Studies have shown that a decrease in PO2 is countered by a decrease in the ratios of [NAD+]/[NADH] and [ATP]/[ADP][Pi] to provide the same driving force for oxidative phosphorylation (61).

During the transition from rest to exercise, there may be a transient mismatch between ATP utilization and aerobic ATP production. This transient mismatch in ATP provision is met by an increased rate of PCr degradation. The resulting increase in the free [Pi], [ADP], and [AMP] serves to stimulate glycogenolysis and ATP provision through substrate phosphorylation from glycolysis. In the present study, the delay in PDH activation at the onset of exercise in hypoxia would have caused a delay in the provision of carbohydrate-derived substrate to the TCA cycle and the subsequent production of reducing equivalents for oxidative phosphorylation. Therefore, in the presence of a similar ratio of [ATP]/[ADP][Pi] at the onset of exercise, a lower [NADH] and possibly a lower PO2 in hypoxia may require higher free [ADP] and [Pi] to drive oxidative phosphorylation and maintain the same ATP turnover rate as in normoxia.

Consequently, an increased PCr degradation after 15 min of exercise in hypoxia resulted in a greater accumulation of free [Pi] compared with normoxia (Fig. 5 and Table 3). The rise in free [Pi], [ADP], and [AMP] served to stimulate glycogenolysis and glycolysis ending in pyruvate production, which in turn exerted a feed-forward effect on PDHa. The increase in PDHa and subsequent substrate delivery to the mitochondrial dehydrogenases of the TCA cycle would have resulted in a rise in [NADH] at the end of exercise in hypoxia (36). In addition to the observed increases in free [ADP] and [Pi], the rise in [NADH] would provide the driving force for oxidative phosphorylation in hypoxia and allow the muscle to function at the same ATP turnover rate as normoxia through changes in the phosphorylation potential and redox state of the cell.

ATP Provision and O2 Equivalents

A study employing a protocol similar to the present study found that VO2 kinetics at the onset of exercise at ~40% VO2 max were delayed under the hypoxic condition relative to the normoxic condition (34). Because of the slower VO2 on-kinetics in the transition from rest to exercise in hypoxia, greater O2 deficits were incurred during hypoxia relative to normoxia in humans exercising at ~40-50% VO2 max (34, 43). This evidence is consistent with decreased ATP provision from oxidative phosphorylation in hypoxia in the transition from rest to exercise, which may be directly attributed to the delayed activation of PDH (Fig. 8). The balance of the ATP equivalents not accounted for by oxidative phosphorylation in hypoxia was provided from substrate phosphorylation by PCr degradation and glycolysis (Fig. 8). During the 1st min of exercise, the delayed activation of PDH in hypoxia resulted in a lower ATP production from oxidative phosphorylation compared with normoxia, which was equivalent to 24 mmol ATP, whereas the greater ATP production from substrate phosphorylation in hypoxia compared with normoxia was 32 mmol ATP (Fig. 8). These differences in ATP provision are equivalent to calculated differences of 200 and 268 ml O2, respectively, which are similar to the O2 deficits incurred in these previously mentioned studies (34, 43).

Grassi et al. (22, 23) have shown that enhanced peripheral diffusion and faster adjustment of O2 delivery do not affect VO2 on-kinetics during electrically stimulated isometric contractions at 60-70% of peak VO2 in dog muscle during normoxia. They suggest that VO2 kinetics are mainly set by an intrinsic inertia of oxidative metabolism at the onset of exercise (22). The latter is supported by the fact that the activation of PDH was delayed by hypoxia in the present study, which reduced the availability of substrate for the TCA cycle and electron transport chain and resulted in reduced O2 utilization. The hypothesis that oxidative phosphorylation is limited by a delayed activation of PDH has been tested in a study by Timmons et al. (58). Activation of PDH by infusion of dichloroacetate resulted in enhanced delivery of oxidative substrate to the TCA cycle and a concomitant reduction of substrate phosphorylation by PCr degradation and glycolysis within 3 min of the onset of ischemic exercise at 65% VO2 max in humans (58). Similar results were observed in a study from our laboratories over a shorter time course (30), in which activation of PDH by dichloroacetate resulted in an improved energy state of the cell and reduced lactate production compared with control, within both 30 s and 2 min of the onset of exercise at 65% VO2 max. This evidence suggests that oxidative phosphorylation is not limited by O2 supply but rather is limited in part by substrate availability through a delayed activation of PDH at the onset of exercise.

Epinephrine, Respiratory Alkalosis, and Lactate Accumulation

Increased lactate production at altitude has been associated with elevated levels of circulating plasma epinephrine (44). However, the same group showed that beta -blockade failed to prevent the rise in lactate production observed during 45 min of exercise at ~50% VO2 max at acute altitude (45). In another study from our laboratories, infusion of epinephrine at concentrations found at altitude failed to increase the rate of glycogenolysis during 90 min of exercise at 65% VO2 max compared with control in untrained subjects (59), suggesting that epinephrine may not be an important determinant of lactate production at altitude.

A number of studies have shown that breathing a low-oxygen gas mixture causes a hyperventilation-induced increase in blood pH at rest (33) and during submaximal exercise (1, 19) compared with normoxia. Hogan et al. (28) employed 31P magnetic resonance spectroscopy and observed a transient increase in pH at the onset of incremental exercise, followed by a significant decrease at 50% of the maximum work rate in humans. In a recent study from this laboratory (29), induced metabolic alkalosis resulted in an enhanced PCr degradation, glycogen utilization, lactate production, and an upregulation of PDHa after 15 min of exercise at 60% VO2 max in humans. These observations were similar to those after 15 min of exercise in the present study, suggesting that extracellular alkalosis may account for the increased lactate production observed in the present study. To determine whether a transient respiratory alkalosis affects the rate-limiting enzymes in carbohydrate metabolism, a study designed to induce hyperventilation during normoxia to the same degree observed during hypoxia might determine whether the acid-base changes and lactate production incurred during hypoxia are in part due to respiratory alkalosis.


    ACKNOWLEDGEMENTS

The authors thank Tina Bragg, Sarah Donaldson, and George Obminski for excellent technical assistance.


    FOOTNOTES

This study was supported by operating grants from the Medical Research and Natural Sciences and Engineering Research Councils of Canada. M. L. Parolin was supported by a Natural Sciences and Engineering Research Council scholarship and by the Ontario Thoracic Society. M. G. Hollidge-Horvat was supported by a Medical Research Council Studentship. G. J. F. Heigenhauser is a Career Investigator of the Heart and Stroke Foundation of Ontario (no. I-2576).

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

Address for reprint requests and other correspondence: G. J. F. Heigenhauser, Department of Medicine, McMaster University Medical Centre, 1200 Main Street West, Hamilton, Ontario, Canada L8N 3Z5 (E-mail: heigeng{at}fhs.csu.mcmaster.ca).

Received 19 July 1999; accepted in final form 22 October 1999.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
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