We examined the regulation of glycogen phosphorylase (Phos) and pyruvate dehydrogenase (PDH) in white muscle of rainbow trout during a continuous bout of high-intensity exercise that led to exhaustion in 52 s. The first 10 s of exercise were supported by creatine phosphate hydrolysis and glycolytic flux from an elevated glycogenolytic flux and yielded a total ATP turnover of 3.7 μmol · g wet tissue−1 · s−1. The high glycolytic flux was achieved by a large transformation of Phos into its active form. Exercise performed from 10 s to exhaustion was at a lower ATP turnover rate (0.5 to 1.2 μmol · g wet tissue−1 · s−1) and therefore at a lower power output. The lower ATP turnover was supported primarily by glycolysis and was reduced because of posttransformational inhibition of Phos by glucose 6-phosphate accumulation. During exercise, there was a gradual activation of PDH, which was fully transformed into its active form by 30 s of exercise. Oxidative phosphorylation, from PDH activation, only contributed 2% to the total ATP turnover, and there was no significant activation of lipid oxidation. The time course of PDH activation was closely associated with an increase in estimated mitochondrial redox (NAD+-to-NADH concentration ratio), suggesting that O2 was not limiting during high-intensity exercise. Thus anaerobiosis may not be responsible for lactate production in trout white muscle during high-intensity exercise.
- adenosine 5′-triphosphate turnover
- mitochondrial redox
- cytoplasmic redox
- oxygen limitation
- rainbow trout
the classical scheme of substrate use during high-intensity exercise involves the temporally separate utilization of creatine phosphate (CrP) followed by activation of glycolysis and then oxidative phosphorylation (16). In this scheme, the initial depletion of CrP at the onset of exercise results in the accumulation of Pi, free ADP (ADPf), free AMP (AMPf), inosine monophosphate (IMP), and NH3, which are thought to activate glycolysis for further ATP production. Subsequently, the accumulation of pyruvate and lactate from glycolysis stimulates pyruvate dehydrogenase (PDH) for ATP production via oxidative phosphorylation. Trout white muscle is known (14) to rely primarily on CrP and glycogen to support high-intensity exercise. However, the majority of research in trout muscle metabolism has focused on elucidating the pattern of metabolite recovery after high-intensity exercise (2,14, 17, 18, 28, 39) and the factors that regulate recovery metabolism (30). As pointed out in numerous studies (e.g., 21, 35, 39), little information is available on the dynamics of substrate selection and the integrated mechanisms that regulate fuel use in trout white muscle during high-intensity exercise.
Dobson et al. (5) examined the dynamics of “anaerobic” ATP production in trout swum at exercise intensities ranging from a 10-s sprint (∼150% critical swimming speed; Ucrit) to 30 min of burst swimming leading to exhaustion. Ten-second sprints were solely supported by CrP hydrolysis. During longer (>10 min) and slower swimming (∼120% Ucrit), glycogen was the primary fuel, utilized through the activation of glycolysis. Control points in glycolysis were identified primarily at phosphofructokinase-1 (PFK-1). Evidence from mammalian studies (3, 22) further implicates the rate-limiting enzyme glycogen phosphorylase (Phos) in regulating glycolytic flux by setting the upper limit for glycogen entry into glycolysis. Phos is regulated at the transformational level by reversible phosphorylation events and at the posttransformational level through substrate availability (Pi), product inhibition (glucose 6-phosphate; G-6-P), and changes in allosteric modulators (ADPf and AMPf; 10, 27). Phosphorylation of Phos by Phos kinase transforms the low-activity form of Phos (Phosb) into the high-activity form (Phosa), whereas the dephosphorylation by Phos phosphatase converts Phosa to Phosb (15). The transformation between Phosb and Phosa is regulated at the contractile level through Ca2+ release from the sarcoplasmic reticulum. Decreases in intracellular pH (pHi) affect the transformation of Phos by inhibiting Phos kinase (4) and affect substrate availability by shifting the speciation of Pi (12). To our knowledge, the transformation of Phos has not been examined in trout white muscle during high-intensity exercise.
PDH is the rate-limiting enzyme that sets the rate at which glycolytically derived pyruvate is decarboxylated for entry into the tricarboxylic acid (TCA) cycle and oxidative phosphorylation. The catalytic rate of PDH is regulated by covalent modification (phosphorylation and dephosphorylation) and product inhibition (37). Dephosphorylation of PDH by PDH phosphatase transforms the inactive PDH (PDHb) into the active PDH (PDHa) whereas phosphorylation of PDH by PDH kinase transforms PDHa into PDHb. PDH kinase is allosterically stimulated by acetyl-CoA, NADH, and ATP and is inhibited by ADP, free CoA (CoA-SH), NAD+, and pyruvate. In mammalian muscle, PDH phosphatase is stimulated by Ca2+ release from the sarcoplasmic reticulum and hormonally by insulin (26). Elevations in mitochondrial NADH inhibit PDH phosphatase and reduce its transformation. In trout white muscle, Richards et al. (28) demonstrated that PDH was maximally activated at exhaustion. Richards et al. suggested that this pathway could have contributed up to 14% of the total ATP production during a bout of high-intensity exercise. Furthermore, the relative transformation patterns and catalytic rates of Phos and PDH have recently been implicated (22, 31) in lactate production in human muscle.
The goal of the present study was to determine the role of the rate-limiting enzymes, Phos and PDH, in regulating white muscle metabolism during a continuous bout of high-intensity exercise. To accomplish these objectives, we measured the transformation states of Phos and PDH, as well as their allosteric regulators (e.g., ADPf, AMPf, Pi, and pHi), in white muscle of trout during a typical high-intensity exercise regime. Insights into cellular O2limitations during high-intensity exercise were made through estimates of cytoplasmic and mitochondrial redox states (NAD+-to-NADH concentration ratio; [NAD+]/[NADH]).
Adult rainbow trout (Oncorhynchus mykiss, Walbaum; 211 ± 8 g, n = 40) were obtained from Humber Springs Trout Hatchery (Orangeville, ON, Canada). Trout were transported to our freshwater holding facility and held in 800-liter tanks supplied with aerated, dechlorinated tap water (from the city of Hamilton) at 15°C for 2 mo before experimentation. Fish were fed daily to satiation with commercial trout pellets. One day before an experiment, fish were placed individually into dark, aerated, 2.5-l acrylic boxes supplied with 100 ml/min freshwater at 15°C. All experimental procedures fully complied with the “Guiding Principles for Research Involving Animals and Human Beings” of the American Physiology Society and the guidelines of the Canadian Council of Animal Care.
To exercise fish, we transferred individual fish without air exposure from the acrylic box to a 150-liter circular tank filled with aerated freshwater at 15°C. Fish were chased with almost constant body contact to avoid burst-glide swimming. White muscle samples were terminally sampled at rest (see below), at 10.5 ± 0.3, 20.4 ± 0.3, and 30.0 ± 0.2 s (n = 8) during high-intensity exercise, and at exhaustion (51.8 ± 3.2 s;n = 8). Fish were considered exhausted when they did not attempt to escape from manual chase. To sample muscle as quickly as possible, fish were removed from the tank by hand (with no struggle by the fish) at a prespecified time and killed by concussion. A 0.5- to 1-cm-thick cross section of the fish was taken posterior to the dorsal fin and immediately freeze clamped between two aluminum blocks cooled in liquid N2. This sampling position is consistent with that used previously in other studies (e.g., 5, 18, 30, and 35). The entire sampling procedure took <5 s.
To obtain resting white muscle samples, we terminally anesthetized trout in their boxes by adding 0.5 g/l 2-aminobenzoic acid ethyl ester (methanesulfonate salt; MS-222) to the surrounding water from a neutralized stock solution. At complete anesthesia (∼1 min), the fish was removed from the water and two muscle samples were taken. The first muscle sample was immediately freeze clamped as described above. The second muscle sample was freeze clamped after a 1-min delay to obtain resting Phos activities (27). All muscle tissues were stored under liquid N2 until analyzed.
The frozen muscle was broken into pieces (50–100 mg) in an insulated mortar and pestle cooled in liquid N2. As in other studies (e.g., 5, 18, 30, and 35), only white muscle dorsal of the lateral line was used. Several aliquots of muscle were stored separately in liquid N2 for determination of PDHa and total PDH activity as previously described by Richards et al. (28). A portion of the frozen muscle (∼50 mg) was ground into a fine powder under liquid N2and used for pHi measurements as described by Pörtner et al. (24). Total muscle NH3 was determined on frozen muscle by the glutamate dehydrogenase method as described by Wang et al. (36). The remaining muscle was lyophilized for 72 h, dissected free of connective tissue, powdered, and stored dry at −80°C for subsequent analysis.
An aliquot of lyophilized muscle was used for determination of Phos activity. Briefly, a known weight (5–10 mg) of dry muscle was homogenized at −25°C in 200 μl of buffer containing 100 mM Tris, 50 mM KCl, and 10 mM EDTA in 60% glycerol at pH 7.5. Homogenates were then diluted with 800 μl of the above buffer without glycerol and homogenized further at 0°C. Total Phos activity (in the presence of 3 mM AMP) and Phosa (in the absence of AMP) were measured by following the production of glucose 1-phosphate (G-1-P) spectrophotometrically at 15°C. Maximal velocity (V max) and the mole fraction of Phosa were calculated as described by Chasiotis et al. (3).
For the determination of muscle glycogen, ∼20 mg of lyophilized muscle were digested in 1 ml 30% KOH at 100°C. Glycogen was isolated as described by Hassid and Abraham (9), and free glucose was determined after digestion with amyloglucosidase (1).
For the extraction of metabolites from white muscle, aliquots of lyophilized muscle (∼20 mg) were weighed into borosilicated tubes, with 1 ml of ice-cold 1 M HClO4 added, and homogenized at the highest speed of a Virtis hand-held homogenizer for 20 s at 0°C. Homogenates were transferred to 1.5-ml bullet tubes and centrifuged for 5 min at 20,000 g at 4°C, and the supernatant was neutralized with 3 M K2CO3. These extracts were assayed spectrophotometrically for ATP, CrP, creatine, lactate, pyruvate, glucose, G-6-P, fructose 6-phosphate (F-6-P), G-1-P, glycerol 3-phosphate (Gly-3-P), glycerol, l-glutamate, and α-ketoglutarate, using the methods previously described by Bergmeyer (1). Muscle acetyl-CoA, CoA-SH, and acetyl-, free-, short-chain fatty acyl- (SCFA), long-chain fatty acyl- (LCFA), and total carnitine were determined on neutralized extracts by radiometric methods previously described in Richards et al. (28).
[ADPf] and [AMPf] were calculated according to Dudley et al. (7) using constants calculated from Schulte et al. (30). [Pi] was calculated as the difference between [resting PCr] and [exercise PCr], less the accumulation of G-6-P, F-6-P, G-1-P, and Gly-3-P. Values for [resting Pi] were estimated from Wang et al. (35) by subtracting [resting PCr] and [ATP] from measured total phosphate and were ∼0.75 μmol/g wet tissue, a value that agrees well with the low resting Pi values reported by van den Thillart et al. (33).
The rates of glycogenolysis and glycolysis (in μmol glycosyl units · g wet weight−1 · s−1) were estimated as described by Spriet et al. (32) from the accumulation of glycolytic intermediates plus the flux of pyruvate through PDHa during each time interval. The equations were as follows where Δ represents a change in tissue metabolite concentration over a specified time period and PDH represents the catalytic rate of PDHa (in μmol glycosyl units · g wet weight−1 · s−1). The catalytic rate of PDHa has been demonstrated to equal flux of pyruvate into the TCA cycle in mammalian muscle (8, 11).
ATP turnover from CrP was calculated from the breakdown of CrP assuming that 1 mol of ATP is produced per mole of CrP consumed. ATP turnover from glycolysis was calculated from the accumulation of lactate and the flux of pyruvate through PDHa assuming 1.5 mol of ATP produced per lactate. The rate of ATP turnover from oxidative phosphorylation was calculated as the total acetyl-CoA production from the PDHa catalytic rate for each time period, assuming 1 mol of acetyl-CoA from glycogenolysis would yield 15 mol of ATP.
The redox state (i.e., [NAD+]/[NADH]) of the cytoplasm was estimated from the apparent equilibrium of the lactate dehydrogenase reactions using measurements of pHi, lactate, and pyruvate and the equilibrium constant (K eq) from Wang et al. (35). Mitochondrial redox was estimated from the glutamate dehydrogenase reaction using whole cell measurements of NH3, glutamate, α-ketoglutarate,K eq from Williamson et al. (38), and estimates of mitochondrial pH from Moyes et al. (19) for carp muscle.
Data presentation and statistical analysis.
All data are presented as means ± SE (n). All metabolite concentrations determined on lyophilized tissues were converted back to wet weights by taking into account a wet-to-dry ratio of 4:1 (35). Statistical analysis consisted of a one-way ANOVA followed by a least significant difference method of pairwise multiple comparisons. Results were considered significant at P< 0.05.
In response to manual chasing, trout swam vigorously for the first 20 s of exercise, then slowed but maintained burst activity until exhaustion at 51.8 ± 3.2 s (n = 8).
Adenylates, CrP, and cellular energy status.
White muscle [ATP] decreased by 28% during the first 10 s of exercise, then remained stable until 20 s and further decreased to 23% of resting [ATP] at exhaustion (Fig.1). During the initial 10 s of exercise, there was a 63% decrease in [CrP] that remained lower than resting values and more or less stable for the duration of the exercise (Fig. 1). These decreases in [CrP] were matched by stoichiometric increases in [Cr], and these relative differences account for the majority of the calculated increase in [Pi] (Table1). The calculated [ADPf] and [AMPf] both increased rapidly within the first 10 s of exercise and then gradually decreased until exhaustion at which point [ADPf] and [AMPf] were not significantly higher than resting values (Table 1). As a result, the ATP-to-ADPf ratio (ATP/ADPf) decreased during the first 10 s of exercise and remained depressed until exhaustion. pHi decreased during the initial 10 s of exercise, stabilized, and then further decreased at exhaustion (Table1). White muscle total NH3 increased gradually over the entire time of exercise (Table 1).
The maximum total Phos activity (V max) was 0.85 ± 0.09 μmol · g wet tissue−1 · s−1 (n = 8) at rest and did not change during exercise. In contrast, Phosa activity increased from 0.16 ± 0.02 μmol · g wet tissue−1 · s−1(n = 8) at rest to 0.42 ± 0.02 μmol · g wet tissue−1 · s−1 (n= 8; P < 0.05) at 10 s and then decreased significantly to 0.34 ± 0.04 μmol · g wet tissue−1 · s−1 at exhaustion (n = 8; P < 0.05). As a result, the calculated transformation of Phosb into Phosaincreased rapidly during the first 10 s of exercise, remained elevated until 30 s of exercise, and then decreased at exhaustion to levels that remained elevated compared with resting values (Fig.2).
The total PDH activity in white muscle of trout was 2.9 ± 0.3 nmol · g wet tissue−1 · s−1(n = 8). In contrast, PDHa activity increased gradually from resting values of 0.3 ± 0.1 (n = 8) to 3.0 ± 0.3 nmol · g wet tissue−1 · s−1 (n = 8) at 30 s and remained elevated until exhaustion. Consequently, the transformation of PDHb into PDHa increased gradually during the first 20 s of exercise, reaching 100% transformation at 30 s and exhaustion (Fig. 2).
White muscle [glycogen] decreased by 35% during the first 10 s of exercise, remained more or less stable until 30 s, and then decreased at exhaustion to levels that were 16% of resting values (Fig. 3). These changes in [glycogen] were matched by reciprocal increases in [lactate] and [pyruvate] (Fig. 3). During the entire bout of exercise, there was a gradual accumulation of the glycolytic intermediates, G-6-P, F-6-P, and G-1-P, all of which peaked in concentration at 20 s (Table 2). Muscle [Gly-3-P] remained stable for the first 30 s of exercise and then increased at exhaustion (Table 2). Muscle [glycerol] increased over the first 20 s of exercise and then remained unchanged until exhaustion (Table 2). There were no changes inl-glutamate or α-ketoglutarate during exercise (Table 2). The calculated rates of glycogenolysis and glycolysis were both high during the first 10 s of exercise then decreased by ∼60% during subsequent exercise (Fig. 4). There was higher glycogenolytic flux compared with glycolytic flux between 10 and 20 s, a trend that reversed between 20 and 30 s. Between 30 s and exhaustion, calculated glycogenolytic and glycolytic fluxes were equal (Fig. 4).
White muscle [acetyl-CoA] did not change during high-intensity exercise, except at 30 s when it decreased slightly (Table3). However, muscle [CoA-SH] decreased by ∼60% during the first 10 s of exercise and remained lower than resting values until exhaustion (Table 3). There was no effect of exercise on muscle [acetyl-carnitine] (Table 3). Muscle [total carnitine] and [free carnitine] were variable, with a trend for a gradual decrease over the exercise period (Table 3). [SCFA-carnitine] and [LCFA-carnitine] did not change during exercise (Table 3).
The present study examined the regulation of substrate use in white muscle of rainbow trout during a continuous bout of high-intensity exercise leading to exhaustion. During the first 10 s of burst activity, high ATP turnover rates (Fig.6) were supported primarily by CrP hydrolysis and glycolysis (Fig. 1 and 4), yielding dramatic decreases in muscle CrP (Fig. 1) and glycogen with corresponding increases in lactate and pyruvate (Fig. 3). The entry of glycogen into glycolysis was the consequence of a very large and rapid transformation of Phosb into Phosa (Fig. 2). Oxidative phosphorylation through PDH made only minor contributions to total ATP turnover during the first 10 s of exercise. Total ATP turnover rate during the first 10 s of exercise was 3.7 μmol · g wet tissue−1 · s−1, which is in close agreement with the maximum ATP turnover rates estimated by Dobson et al. (5) for trout white muscle (3.1 μmol · g wet tissue−1 · s−1).
During longer periods of exercise (>10 s), Phos remained significantly transformed compared with resting values (Fig. 2), but glycogenolytic and glycolytic fluxes were reduced (Fig. 4), therefore leading to a lower ATP turnover from glycolysis (Fig. 6) and likely a correspondingly reduced power output. However, glycolysis still supported the majority of the total ATP turnover throughout the exercise period (Fig. 6). CrP hydrolysis made only minor contributions to total ATP turnover after the initial 10 s of exercise. As exercise duration increased, there was a gradual transformation of PDHb into PDHa that was complete by 30 s of exercise; however, oxidative phosphorylation from carbohydrate only contributed ∼ 2% to the total ATP turnover during the bout of exercise (Fig. 6).
Adenylates and CrP.
Considerable debate surrounds the sequence of substrate utilization by muscle during maximal contraction. It is generally believed that CrP hydrolysis is tightly linked to myofibrilliar ATPase activity through high-affinity, high-activity creatine kinase (16). Once the ATP-buffering capacity of CrP is exceeded, endogenous ATP is utilized, yielding increases in ADPf, AMPf, IMP, NH3, and Pi, all of which act to stimulate glycogenolysis and ATP production via glycolysis. Indeed, during the first 10 s of exercise CrP was depleted by 63% whereas ATP only decreased by 28% (Fig. 1), indicating that a greater proportion of CrP was hydrolyzed before endogenous ATP. However, during the first 20 s of exercise, muscle [ATP] remained elevated compared with [ATP] at exhaustion, suggesting that complete depletion of ATP was not required to activate glycolysis. Decreasing [ATP] was associated with stoichiometric increases in NH3 (Table 1), indicating the deamination of adenylates during high-intensity exercise and the formation of IMP (30).
Regulation of Phos.
It has long been recognized (2, 14, 17, 28, 35) that glycogen is an important substrate for ATP production in fish white muscle during bouts of burst activity. During the initial 10 s of exercise, glycogenolytic flux and glycolytic flux supported ∼40% of the ATP turnover (Fig. 6). Subsequently, glycogenolytic and glycolytic flux both decreased by about two-thirds but continued to contribute the largest portion to ATP turnover from 10 s of exercise to exhaustion.
The high glycogenolytic flux observed during the first 10 s of exercise (Fig. 4) was achieved primarily by a large transformation of Phosb into Phosa (Fig. 2), probably caused by Ca2+ release from the sarcoplasmic reticulum activating Phos kinase (15). The calculatedV max of Phosa during the initial 10 s of exercise was 0.42 μmol · g wet tissue−1 · s−1; therefore the increase in activity due to Phos transformation was nearly sufficient to completely explain the observed glycogenolytic flux (Fig. 4). In addition to increased Phos activity due to the transformational modification, posttransformational modification of Phosaalso contributed to increased glycogenolysis. Accumulation of Pi from CrP hydrolysis (Table 1; Fig. 1) increases substrate availability during the first 10 s of exercise, and the accumulation of AMPf (Table 1) is thought to decrease theK m of Phosa for Pi and thus increase the catalytic rate (27). In addition, the accumulation of the allosteric activators (ADPf; Table 1), decreasing cellular energy status (ATP/ADPf; Table 1), and lack of product accumulation (G-6-P; Table 2) would further allosterically increase Phosa activity to allow for the observed glycogenolytic flux.
Glycogenolytic flux decreased linearly during the first 30 s of high-intensity exercise (Fig. 4). During this period of decreased glycogenolysis, Phos remained significantly transformed into Phosa (Fig. 2), and the calculatedV max of Phosa was two- to threefold larger than the calculated glycogenolytic flux. As a result, after the initial 10 s of exercise, Phos activity was primarily mediated via posttransformational modification. The decreased flux through Phosa was likely mediated by an increase in product, G-6-P (Table 2), acting to inhibit Phosa in addition to the decrease in AMPf observed at 20 s compared with 10 s (Table 1), which would reduce the activation of Phosa. G-6-P remained elevated and AMPf remained at lower than peak levels until exhaustion. Decreasing pHi inhibits Phos kinase and would reduce the transformation of Phos (4). In addition, decreasing pHi would shift the chemical form of Pi from the monoprotonated form, which is the active substrate for Phosa, to the diprotonated form, which does not act as a substrate for Phosa (12). Changes in muscle NH3 suggest that at >10 s of exercise IMP was accumulating from the deamination of AMP, which is also thought to allosterically stimulate Phosa (3). However, it appears that G-6-P and AMPf are the two important allosteric regulators of Phos activity in trout white muscle.
In general, the calculated fluxes through Phos and glycolysis were reasonably well matched throughout the exercise period (Fig. 4), although during the initial 20 s of exercise, there was greater glycogenolytic flux than glycolytic flux and this pattern was reversed between 20 and 30 s. This match between Phos and glycolysis suggests that Phos transformation and catalytic rate directly set the upper limit for glycolytic flux and therefore pyruvate and lactate production. The slightly lower glycolytic rate observed during the first 20 s of exercise suggests that PFK-1 modifies the glycolytic rate set by Phos, a conclusion reinforced by the accumulation of F-6-P and G-6-P during this period (Table 2). Dobson et al. (5) identified PFK-1 as a point of control in trout white muscle during exercise but did not critically examine the role of Phos. Considering the very close match between glycogenolytic flux and glycolytic flux observed in Fig. 4, it appears that the transformation state of Phos sets the upper limit for glycogenolytic and glycolytic flux in trout muscle, and flux is further modified by posttransformational modification of Phos and allosteric regulation of PFK-1 activity (6, 23).
Regulation of PDH.
During exercise, the transformation of PDHb into PDHa was slow to occur, taking 30 s of intense activity before PDHb was fully transformed into PDHa (Fig. 2). The initial cue for the transformation of PDH is generally thought to be Ca2+ release from the sarcoplasmic reticulum (26). However, Ca2+does not appear to be a potent stimulator of PDH phosphatase in trout white muscle because of the considerable delay in PDH transformation (Fig. 2). Within 10 s of exercise, the energy status (ATP/ADPf) of the cells was substantially reduced with a large and significant increase in ADPf and pyruvate (Table1; Fig. 3), which should inhibit PDH kinase (37) and further allow Ca2+-mediated activation of PDH phosphatase and thus PDH transformation. However, in trout white muscle there must be factors that override the stimulatory effects of Ca2+ on PDH and slow its transformation. Thus the reason for the delayed transformation of PDH appears, at first glance, to be paradoxical.
Two possible explanations exist to explain the delayed activation of PDH (Fig. 2) in trout muscle. First, PDH is located within the mitochondrial matrix and is therefore subject to regulation by changes in mitochondrial metabolites and energy status. During the transition from rest to 10 s of exercise there is a significant decrease in CoA-SH in the absence of any change in acetyl-CoA; thus the acetyl-CoA-to-CoA-SH ratio (acetyl-CoA/CoA-SH) increases substantially during the first 10 s of exercise. This increase in acetyl-CoA/CoA-SH could stimulate PDH kinase (10) and slow the transformation of PDH at the onset of exercise. The reason for decreasing CoA-SH is also paradoxical, but likely CoA-SH is bound by TCA cycle intermediates or fatty acids (34). In human muscle, acetyl-CoA/CoA-SH is thought to contribute to PDH regulation only at rest, and its role in regulating PDH during exercise is thought to be minimal (25). However, in fish muscle the importance of acetyl-CoA/CoA-SH maybe greater.
The second plausible reason for the delayed PDH activation in trout muscle is related to the estimated redox state of the mitochondrial matrix. The pattern of PDH transformation (Fig. 2) closely approximates the estimated changes in muscle mitochondrial redox (Fig. 5). This suggests that increases in mitochondrial [NAD+] act to stimulate PDH phosphatase and inhibit PDH kinase in trout muscle. Therefore, it appears that PDH transformation may be more closely related to the redox state of the cellular compartment within which PDH is found than to the general energy status of the cell or Ca2+ release from the sarcoplasmic reticulum.
In all studies that have examined PDH activation to date (e.g., 8, 11), the activity of PDH is very closely matched to the total flux through the TCA cycle and thus oxidative phosphorylation. In previous work examining recovery metabolism in trout white muscle, we (28) observed that immediately after high-intensity exercise PDH was fully transformed into PDHa. In that study (28), we assumed an immediate transformation of PDHb to PDHa at the onset of exercise and calculated that oxidative phosphorylation from carbohydrate could contribute up to a maximum of 14% of the total ATP turnover during a 5-min bout of high-intensity exercise. However, in the present study, we have demonstrated that PDH transformation is slow to occur over a 52-s bout of exercise, therefore oxidative phosphorylation only contributes ∼2% to total ATP turnover during high-intensity exercise in trout muscle. Despite the lack of ATP production via oxidative phosphorylation of carbohydrate in trout muscle, the pattern of PDH transformation has implications toward lactate metabolism.
Lactate metabolism in trout white muscle.
Classically, lactate production was thought to occur during high-intensity exercise because of a limitation of O2supply to the mitochondria preventing oxidative phosphorylation, thus activating “anaerobic” glycolysis for ATP production. However, recent evidence (10, 31) suggests that O2 may not be limiting in mammalian muscle during high-intensity exercise but rather that lactate production may be due to metabolic inertia whereby pyruvate production exceeds pyruvate oxidation. The present study contributes to the accumulating evidence (e.g., 22) that suggests lactate accumulation is due to an imbalance between the transformation pattern and catalytic rates of Phos and PDH. The catalytic rate of Phosa in trout white muscle is about 140 times greater than the catalytic rate of PDHa (Phos, ∼0.42 μmol · g wet tissue−1 · s−1; cf. PDH, 0.003 μmol · g wet tissue−1 · s−1; seeresults). The low mitochondrial content present within trout white muscle and likely low copy number of PDH will accentuate the mismatch between glycolytic flux and pyruvate oxidation and will therefore yield lactate accumulation via lactate dehydrogenase.
Considerable controversy surrounds the issue of whether an O2 limitation in muscle cells plays a role in lactate formation during high-intensity exercise (13). Measurements of whole cell [NADH] and [NAD+] suggest that the cell becomes more reduced with increases in exercise intensity (29). However, as with the adenylates (ADP and AMP), changes in total [NADH] and [NAD+] do not provide information regarding the regulation of metabolism; only changes in free NADH and NAD+ provide insight into metabolic control. On the basis of the lactate dehydrogenase reaction, we have demonstrated that cytoplasmic redox remains constant during high-intensity exercise. This is in direct agreement with previous work from our laboratory (35) and further agrees with Dobson et al. (5) in trout swum for 10 min at 120% Ucrit.
Estimates of cytoplasmic redox do not yield information regarding the redox state of the mitochondrial matrix. Our estimates of mitochondrial NAD+/NADH in trout muscle during exercise suggest that the mitochondrial matrix becomes more oxidized as exercise progresses (Fig.5) and that this is associated with the progressive activation of PDH (Fig. 2), supporting the idea that the electron transport chain is functioning. However, it should be noted that the use of the glutamate dehydrogenase equilibrium to estimate mitochondrial redox state remains highly controversial (13), especially in white muscle with low mitochondrial number (20). Therefore, the conclusion that O2 is not limiting at the mitochondrion during high-intensity exercise in trout muscle must be considered tentative and awaits confirmation by other methods.
If O2 is not limiting in white muscle mitochondria during high-intensity exercise, it is possible that other substrates are also used for oxidative ATP production. Because lipid oxidation is very important during postexercise recovery (28), we wished to determine whether lipid oxidation could contribute significantly to ATP production in trout white muscle during high-intensity exercise. There was a small, transient decrease in free carnitine within the first 10 s of exercise, but this decrease did not correspond to an increase in either SCFA or LCFA-carnitine and there were no increases in acetyl-CoA (Table 3). These data indicate that there was not an increase in fatty acid transport into the mitochondria by carnitine palmitoyltransferase I.
In conclusion, we have demonstrated that during the first 10 s of high-intensity exercise CrP hydrolysis and glycolysis support ATP turnover. Subsequent exercise through to exhaustion is supported primarily by glycolytic ATP production. Entry of glycogen into glycolysis is regulated by the transformational and posttransformational modification of Phos. As exercise duration increased there was a near maximal activation of PDH, and the mismatch between Phos and PDH transformation can explain lactate production in fish muscle. Estimates of cytoplasmic and mitochondrial NAD+/NADH, in addition to the activation pattern of PDH, suggest that O2 is not limiting in white muscle cells during high-intensity exercise and is not the primary cause of lactate formation.
Many studies have examined the effects of exhaustive exercise on trout white muscle metabolism and looked at the pattern of metabolite recovery. However, the present study is the first to document changes in muscle metabolites during a continuous bout of exercise. Measurements of the transformation state of regulating enzymes (Phos and PDH) and their allosteric modulators have added evidence to support the hypothesis that lactate production during exercise is not due to an O2 limitation but rather to an imbalance between Phos and PDH transformation. The slow activation pattern of PDH in trout white muscle indicates that the major fate of lactate is not oxidation. Instead, lactate production in white muscle could be considered integral to white muscle design in fish, so that accumulated lactate is conserved as the substrate for glycogen synthesis during recovery (17, 28, 30). Further research should focus on isolating the role of PDH in white muscle lactate production through manipulation of PDH activity with dichloroacetate (a pharmacological analog of pyruvate), hyperoxia, and hypoxia.
We gratefully acknowledge E. Fitzgerald for the excellent technical assistance.
This work was supported by the Natural Sciences and Engineering Research Council of Canada (NSERC) (C. M. Wood) and the Medical Research Council of Canada (G. J. F. Heigenhauser). J. G. Richards was supported by an NSERC postgraduate scholarship, and C.M. Wood is supported by the Canada Research Chair Program.
Address for reprint requests and other correspondence: J. G. Richards, Dept. of Biology, McMaster Univ., 1280 Main St. West, Hamilton, Ontario, Canada L8S 4K1 (E-mail:).
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.
- Copyright © 2002 the American Physiological Society