Acute High-Intensity Exercise Impairs Skeletal Muscle Respiratory Capacity : Medicine & Science in Sports & Exercise

Journal Logo

BASIC SCIENCES

Acute High-Intensity Exercise Impairs Skeletal Muscle Respiratory Capacity

LAYEC, GWENAEL1,2,3; BLAIN, GREGORY M.4; ROSSMAN, MATTHEW J.2; PARK, SONG Y.2; HART, COREY R.2; TRINITY, JOEL D.1,2,3; GIFFORD, JAYSON R.1,2,3; SIDHU, SIMRANJIT K.1,5; WEAVIL, JOSHUA C.1; HUREAU, THOMAS J.1,3,6; AMANN, MARKUS1,2,3; RICHARDSON, RUSSELL S.1,2,3

Author Information
Medicine & Science in Sports & Exercise 50(12):p 2409-2417, December 2018. | DOI: 10.1249/MSS.0000000000001735
  • Free

Abstract

Skeletal muscle mitochondria play a critical role in maintaining cellular homeostasis while also providing the energy required for activities of daily living and exercise. There have been a host of both cross-sectional (1,2) and longitudinal studies documenting that increased physical activity and chronic endurance exercise enhance muscle mitochondrial content (3,4) and respiratory capacity (5–7). However, in contrast, the effect of acute exercise on mitochondrial respiratory function is still equivocal, which is interesting in light of the many, potentially deleterious, intracellular perturbations associated with acute exercise, particularly when performed at a high intensity. Specifically, after acute high-intensity exercise, skeletal muscle mitochondrial oxidative phosphorylation may be hindered by exercise-induced reactive oxygen (O2) species as well as the accumulation of inorganic phosphate, lactate, and protons (8), all either within or near the mitochondria.

Initial animal studies reported that high-intensity exercise to exhaustion induced acute mitochondrial swelling (9) and transiently impaired mitochondrial respiration in skeletal muscle (10–12). However, later studies in humans documented unchanged or even increased complex I–specific ADP-stimulated respiration (state 3CI) and mitochondrial efficiency (P/O) in isolated mitochondria and permeabilized muscle fibers immediately after prolonged exercise at a moderate intensity [60%–75% of maximal oxygen uptake (V˙O2max)] (13–16) or intermittent supramaximal cycling exercise (130% of V˙O2max) (17). However, a caveat of these prior studies is that the mitochondrial respiration attained during state 3CI yields only 65% to 75% of the maximal respiratory rate that can be measured without the additional substrates necessary to facilitate convergent electron flow through complexes I and II (state 3CI + CII) (18). The tricarboxylic acid (TCA) cycle is, indeed, not complete using NADH-specific substrates (e.g., pyruvate and malate, or glutamate and malate) such that citrate, isocitrate, 2-oxoglutarate, and succinate are depleted, which, in turn, prevents both the electron flow through complex II and the attainment of true maximal muscle respiratory capacity (18). Consequently, the results of these studies do not constitute unequivocal evidence of an uncompromised electron transport chain (ETC) after acute exercise. Furthermore, the acute effects on mitochondrial respiratory function of a short-duration high-intensity exercise that ultimately yields V˙O2max and elicits improvements in muscle aerobic capacity when performed chronically (19,20) have yet to be assessed.

Consequently, the purpose of this study was to comprehensively examine respiratory flux through the complexes of the ETC in skeletal muscle mitochondria, both before and immediately after high-intensity aerobic exercise. We used a 5-km cycling time trial, an exercise modality that increases aerobic metabolism to V˙O2max for several minutes and induces considerable intracellular metabolic perturbations and fatigue (8). On the basis of the large intracellular perturbations associated with this form of exercise (8) and the apparently preserved complex I–specific ADP-stimulated respiration immediately after exercise (13–16), we hypothesized that complex I + II state 3 (state 3CI + CII) respiration, assessed in permeabilized muscle fibers obtained from the vastus lateralis, would be significantly diminished immediately after exercise as a consequence of compromised electron flow through complex II.

METHODS

Subjects

Following informed consent procedures, eight recreationally active, healthy, nonmedicated, and nonsmoking men participated in this study (mean ± SD: weight, 83 ± 14 kg; height, 180 ± 6 cm; age, 26 ± 2 yr). Intramuscular metabolites and neuromuscular function results have previously been published for these subjects (8). Subjects were instructed to refrain from exercise for 48 h and caffeine for 24 h before each trial. The study was approved by the institutional review boards of both the University of Utah and the Salt Lake City Veterans Affairs Medical Center.

Exercise protocol

During preliminary visits, all participants performed two to three practice 5-km cycling time trials and a maximal incremental exercise test to exhaustion to determine V˙O2peak on a computer-controlled electromagnetically braked cycle ergometer (Velotron, Elite Model; Racer Mate, Seattle, WA). On the day of the experiment, with the subject lying on a bed, a percutaneous biopsy of the vastus lateralis muscle, approximately 3.5 cm deep, 15 cm proximal to the knee, and slightly distal to the ventral midline of the muscle, was obtained from the right leg before exercise. The 5-mm diameter biopsy needle (Bergstrom) was attached to sterile tubing and a syringe to apply a negative pressure to assist in the muscle sample collection (21). Immediately after the muscle sample (~100 mg) was removed from the leg, a portion of the sample (~20 mg) was immersed in ice-cold biopsy preservation fluid BIOPS (in mM: 2.77 CaK2EGTA, 7.23 K2EGTA, 20 imidazole, 50 K+MES, 20 taurine, 0.5 dithiothreitol, 6.56 MgCl2, 5.77 ATP, 15 phosphocreatine, pH 7.1) for respiratory analysis, whereas the remaining sample was immediately frozen and stored at −80°C for later analysis (8). Subjects then moved to the cycle ergometer to complete a 5-km cycling time trial with freedom to alter power output by changing the gear ratio and/or pedaling frequency (8). Immediately after the exercise, a cuff, placed on the upper part of the thigh, was rapidly inflated to suprasystolic pressure (250 mm Hg) to clamp the metabolic milieu until muscle sampling was complete (<30 s after exercise cessation). Given the relatively large O2 store of the muscle (22,23) and the relatively short duration of the ischemia in comparison to the time documented to induce mitochondrial defects (24), it is unlikely that this procedure elicited an anoxic state and, in turn, altered mitochondrial respiratory capacity. The right leg was used for baseline sampling, whereas the left leg was used for postexercise sampling.

Pulmonary gas exchange measurements

At rest and throughout exercise, pulmonary gas exchange and ventilation were measured continuously using an open-circuit calorimetry system (True Max 2400; Parvo Medics, Salt Lake City, UT).

Preparation of permeabilized muscle fibers and mitochondrial respiration measurements

The tissue preparation and respiration measurement techniques were adapted from established methods (25,26) and have been previously described (27). Briefly, BIOPS-immersed fibers were carefully separated with fine-tip forceps and subsequently bathed in a BIOPS-based saponin solution (50 μg saponin·mL−1 BIOPS) for 30 min. After saponin treatment, muscle fibers were rinsed twice in ice-cold mitochondrial respiration fluid (MIR05, in mM: 110 sucrose, 0.5 EGTA, 3 MgCl2, 60 K-lactobionate, 20 taurine, 10 KH2PO4, 20 HEPES, BSA 1 g·L−1, pH 7.1) for 10 min each. After the muscle sample was gently dabbed with a paper towel to remove excess fluid, the wet weight of the sample was measured using a standard, calibrated scale (2–4 mg). The muscle fibers were then placed in the respiration chamber (Oxytherm; Hansatech Instruments, King’s Lynn, UK) with 2 mL of MIR05 solution and warmed to 37°C. After allowing the permeabilized muscle sample to equilibrate for 5 min, mitochondrial respiratory function was assessed in duplicate. To assess the function of each mitochondrial complex, O2 consumption was assessed with the addition of a series of respiratory substrates and inhibitors in the following order and final concentrations in the chamber: glutamate-malate (10 and 2 mM), ADP (5 mM), succinate (10 mM), cytochrome c (10 μM), rotenone (0.5 μM), antimycin-A (2.5 μM), oligomycin (2 μM), and N,N,N,N-tetramethyl-p-phenylenediamine (TMPD)–ascorbate (2 and 0.5 mM). Pilot studies indicated that the concentrations of the substrates and inhibitors used were at saturating levels (27). This allowed the determination of 1) state 2 respiration, the nonphosphorylating resting state, which provides an index of proton leak, assessed in the presence of malate + glutamate; 2) complex I–driven state 3 respiration (state 3CI), the ADP-activated state of oxidative phosphorylation, assessed in the presence of glutamate + malate + ADP; 3) complex I + II–driven state 3 respiration (state 3CI + CII), assessed in the presence of glutamate + malate + ADP + succinate; 4) complex II–driven state 3 respiration (state 3CII), assessed in the presence of glutamate + malate + ADP + succinate + rotenone; and 5) uncoupledCIV respiration, where the link between the ETC and ATP synthesis has been abolished, assessed by inhibiting complex III (antimycin A) and complex V (oligomycin) followed by the addition of TMPD + ascorbate.

Respirometry data analysis

Only one sample demonstrated impaired mitochondrial membrane integrity (a >10% increase in respiration in response to cytochrome c) and was therefore excluded from the analysis. In each condition, the respiration rate was recorded for at least 3 min until a steady state was reached, and the average of the last minute was used for data analysis. Inhibition of respiration with combined antimycin A and oligomycin allowed for the determination and correction for residual O2 consumption, indicative of nonmitochondrial O2 consumption (i.e., from chemical and instrument-related respiration in the chamber). Given the within-subject comparison design, the rate of O2 consumption was simply expressed relative to muscle sample mass (in picomoles per second per milligram of wet weight) as mitochondrial content is unlikely to change within such a short time frame (28). The leak control ratio (i.e., the ratio between state 2 and uncoupledCIV respiration) was calculated as an index of uncoupling due to electron leak across the membrane or slip of protons in the respiratory chain (25). The ratio between state 3CI + CII and uncoupledCIV respiration was also calculated as an index of the excess capacity of cytochrome c oxidase (25,29,30).

Citrate synthase and succinate dehydrogenase activity

After the respiration measurements, the same muscle samples (2–4 mg wet weight) were homogenized with homogenization buffer containing 250 mM sucrose, 40 mM KCl, 2 mM EGTA, and 20 mM Tris·HCl (Qiagen, Hilden, Germany). The citrate synthase (CS) activity assay was performed as previously described (27) using a spectrophotometer with light absorbance set at 412 nm (Synergy 4; Biotek Instruments, Winooski, VT). Complex II enzyme activity, succinate dehydrogenase (SDH) of homogenized tissue sample was measured using a microplate assay kit and performed according to the manufacturer’s recommendations (Abcam ab109908). In this assay, the production of ubiquinol by complex II is coupled to the reduction of the dye 2,6-diclorophenolindophenol, and SDH activity was measured using a spectrophotometer with light absorbance set at 600 nm.

Statistical analysis

Because of the limited sample size, differences between baseline and postexercise were evaluated using a nonparametric Wilcoxon test (Statsoft, version 5.5; Statistica, Tulsa, OK). Statistical significance was accepted at P < 0.05. Results are presented as mean ± SD in figures.

RESULTS

V˙O2peak and time-trial performance

During the maximal and graded cycling exercise, the group mean V˙O2peak was 3.6 ± 0.5 L·min−1 corresponding to 43.8 ± 6.8 mL·min−1·kg−1, which was reached at 296 ± 37 W. The mean power output over the entire 5-km time trial was 220 ± 24 W for an exercise duration of 8.75 ± 0.38 min. Changes in pulmonary V˙O2 throughout the 5-km time trial are illustrated in Figure 1. The time spent over 80% of V˙O2peak averaged 5.1 ± 2.5 min.

F1
FIGURE 1:
Pulmonary oxygen consumption (V˙O2) during the 5-km cycling time trial. The dashed line indicates the oxygen consumption corresponding to 80% of the group V˙O2peak, whereas the solid upper line represents the group V˙O2peak. Values are presented as mean ± SD.

Mitochondrial respiration

The rates of O2 consumption for state 2, state 3I, 3II, 3I + II, and uncoupledCIV respiration are summarized in Figure 2. State 3II and state 3I + II respiration rates were decreased immediately after the 5-km cycling time trial (P < 0.05). In contrast, state 2 (P = 0.7), state 3I (P = 0.27), and uncoupledCIV respiration (P = 0.89) were not significantly different between baseline and immediately after the exercise. As illustrated in Figure 3, the leak control ratio was not significantly changed (P = 0.90), whereas the ratio between state 3CI+ CII and uncoupledCIV was diminished immediately after the exercise (P < 0.05).

F2
FIGURE 2:
Skeletal muscle respiratory fluxes (O2 consumption rates per milligram of tissue) at resting baseline and immediately after a 5-km time trial. The following substrates were used: state 2, malate + glutamate; state 3 CI, malate + glutamate + ADP; state 3 CI + II, malate + glutamate + succinate + ADP; state 3 CII, malate + glutamate + succinate + ADP + rotenone (complex I inhibitor); and uncoupledCIV respiration, corresponding to ETC capacity, malate + glutamate + succinate + ADP + rotenone + antimycin A (complex III inhibitor) + oligomycin (complex V inhibitor) + ascorbate + TMPD. Values are presented as mean ± SD after correction for nonmitochondrial O2 consumption. *P < 0.05, significantly different from baseline.
F3
FIGURE 3:
Respiratory control ratio for leak (state 2/uncoupledCIV respiration) and the ratio between state 3CI + CII and uncoupledCIV respiration, an index of the relative excess capacity of cytochrome c oxidase, at resting baseline and immediately after a 5-km time trial. Values are presented as mean ± SD. *P < 0.05, significantly different from baseline.

CS and SDH activity

Although the CS activity increased by approximately 41% from baseline to end-exercise (baseline, 17.2 ± 12.1 au; end-exercise, 24.3 ± 8.8 au), likely due to between-subject variability, this did not reach statistical significance (P = 0.18). Also, SDH activity was not significantly different between baseline and immediately after the exercise (baseline, 1.15 ± 0.6 au; end-exercise, 0.97 ± 0.7 au; P = 0.62).

DISCUSSION

This study sought to determine the impact of high-intensity, aerobic, whole-body cycling exercise on respiratory flux through the ETC of locomotor muscle mitochondria. In agreement with our hypothesis, state 3CI + CII respiration was significantly diminished immediately after completion of a 5-km cycling time trial. This decreased muscle respiratory capacity was predominantly the consequence of attenuated state 3CII respiration, as state 3CI respiration, CS activity, state 2 respiration, and uncoupledCIV respiration were not significantly affected by the exercise. Collectively, these findings reveal that the metabolic challenge imposed by acute high-intensity cycling exercise, likely transiently, compromises skeletal muscle state 3CII and 3CI + CII respiration and therefore oxidative phosphorylation capacity. This attenuated mitochondrial respiratory function may amplify the exercise-induced development of fatigue and could also play an important role in initiating exercise-induced mitochondrial adaptations.

Acute high-intensity exercise and skeletal muscle oxidative phosphorylation capacity

The two key and novel findings of this study are that the completion of a 5-km cycling time trial, eliciting a sustained metabolic demand close to V˙O2peak (Fig. 1), resulted in 1) a lower maximal respiratory rate, as assessed in vitro with substrates for convergent electron flow through complexes I and II of the ETC (Fig. 2), and 2) a decrease in the phosphorylation system control ratio (Fig. 3). Thus, the oxidative phosphorylation capacity of the skeletal muscle, a major determinant of endurance exercise performance (31), was acutely compromised by a bout of high-intensity, whole-body exercise. This lower state 3CI + CII respiration seemed to be predominantly mediated by attenuated complex II–specific respiration (50%, Fig. 2) and, to some extent, by an approximately 30% (nonsignificant) decline in complex I–specific respiration.

Consistent with the animal literature (11,12), the present study suggests that acute high-intensity aerobic whole-body exercise can, likely transiently, inhibit mitochondrial respiratory capacity. Somewhat in contrast to the current findings, it has previously been reported in humans that skeletal muscle state 3CI respiration is unaffected or even increased immediately after prolonged cycling exercise at a moderate intensity (60%–75% of V˙O2max) (13–16) or after intermittent supramaximal cycling exercise (130% of V˙O2max for an exercise duration of ~2.5 min) (17). In addition, Rasmussen et al. (32) also reported that, in isolated mitochondria from the vastus lateralis, state 3CI and state 3CII, assessed separately, were not significantly altered after five bouts of 1-min cycling exercise to exhaustion. The reason for the discrepancy between the current findings and previous studies may be attributable to several factors including the preparation used (isolated mitochondria vs permeabilized muscle fibers), the titration protocol (the separate assessment of the mitochondrial complexes vs convergent electron flow through complexes I and II), and/or the exercise protocol used. Indeed, mitochondrial isolation may result in a biased selection of intact organelles (33), which might have precluded the observation of a detrimental effect of exercise on mitochondrial respiratory function. Also, the attenuated state 3CI + CII respiration was, to a large extent, the consequence of compromised respiration through complex II, such that our findings are consistent with previous studies indicating that state 3CI respiration is unaffected by exercise. With regard to the exercise protocol, by design, it is likely that the metabolic challenge in the current study differed from prolonged moderate exercise, which requires substantial energy supply from β-oxidation, used in previous investigations (13–16,34,35). The current subjects during the 5-km time trial demonstrated a sustained high-intensity aerobic effort culminating near V˙O2max (Fig. 1), which requires that mitochondrial oxidative phosphorylation is fueled by carbohydrate substrates and operates at or near maximal capacity (36) for several consecutive minutes. This type of effort might also represent a greater metabolic challenge for the mitochondria than short-duration intermittent supramaximal cycling exercise (17,32), during which anaerobic metabolism (glycolysis and phosphocreatine) contributes significantly to ATP production.

High-intensity exercise-induced attenuation in oxidative phosphorylation: putative mechanisms

There are multiple putative mechanisms that may explain the attenuated oxidative phosphorylation documented in this study. Among them, diminished substrate availability due to the impaired activity of some of the key enzymes of the TCA cycle is likely to have played a role. Indeed, although oxidative phosphorylation capacity decreases as a consequence of high-intensity whole-body exercise by nearly 40%, mitochondrial permeability to protons (state 2 respiration, Fig. 2), leak control ratio (Fig. 3), and cytochrome c oxidase (uncoupledCIV respiration, Fig. 2) all seem to be well preserved. These findings imply that, despite saturating substrate concentrations in the buffer solution (glutamate, malate, and succinate), factors upstream of the respiratory complexes, such as the impaired activity of some enzymes of the TCA cycle, might have been responsible for the decrease in state 3CI + CII respiration by limiting the amount of FADH provided to complex II. This interpretation is further supported by the lower phosphorylation system control ratio immediately after the exercise (Fig. 3), which indicates a lower phosphorylative constraint relative to ETC capacity, assessed at the level of complex IV, considering that the ETC is ultimately limited by its terminal enzyme, cytochrome c oxidase, in human skeletal muscle (29,30). In addition, although respirometry measurements indicated attenuated state 3CII respiration after exercise, SDH activity was not significantly different between baseline and immediately after the exercise. Therefore, in the conditions generated in the current study, the observed attenuation in oxidative phosphorylation capacity may be a consequence of decreased substrate availability, although a lower ATP synthase activity is also a possible explanation. Somewhat conflicting with the hypothesis of a deficit in substrate availability, CS activity, an enzyme of the TCA cycle, was unchanged or even tended to increase (P = 0.18) after exercise. However, previous studies documented divergent effects of exercise on several enzymes of the TCA cycle (7,32). For instance, after five bouts of 1-min cycling exercise to exhaustion, some enzymes have been reported to be unchanged (e.g., cytochrome oxidase and CS), decreased (pyruvate dehydrogenase, α-ketoglutarate dehydrogenase, glutamate dehydrogenase), or even increased (α-glycerophosphate dehydrogenase, NADH oxidase) (32). Therefore, CS activity could have been maintained in the current scenario, whereas other enzymes of the TCA such as α-ketoglutarate dehydrogenase or aconitase may well have been decreased and in turn attenuated the production of reducing equivalents.

High-intensity cycling exercise greatly increases free radical levels (37), and therefore, reactive O2 species in the muscle bed and in particular hydrogen peroxide (38) may be the link connecting exercise to the acute changes in mitochondrial respiratory function observed in the current study. Indeed, there is growing evidence that high-intensity exercise or in vitro administration of physiological concentrations of hydrogen peroxide can inhibit certain redox-sensitive enzymes of the Krebs cycle (α-ketoglutarate, aconitase, and SDH) (7,32,39,40). For instance, during exercise, reactive oxygen species (ROS)–mediated inhibition of aconitase activity has been suggested to result in citrate accumulation to protect the mitochondria from oxidative damage while inhibiting mitochondrial respiration through the attenuation of reducing equivalents generation (36). According to this scenario, the fall in substrate availability for oxidative phosphorylation would diminish mitochondrial membrane potential and ADP-stimulated mitochondrial respiration (41,42) and in turn trigger the dynamic remodeling of the mitochondrial network, promoting the morphological changes documented after chronic exercise interventions. Consistent with this scenario, energetic stress has been suggested to activate AMP-activated protein kinase, which stimulates mitochondrial biogenesis via peroxisome proliferator–activated receptor-γ coactivator upregulation and simultaneously triggers the destruction of existing defective mitochondria through mitophagy (43), resulting in improved mitochondrial quantity and quality. However, in conditions with excessive ROS production, the inhibition of aconitase may abolish citrate accumulation and upregulate the endogenous antioxidant system to restore the redox balance rather than stimulating mitochondrial biogenesis (36).

High-intensity exercise is commonly associated with marked metabolic disturbances that can also influence mitochondrial function. Specifically, during high-intensity exercise, inorganic phosphate, lactate, and protons accumulate in the exercising muscle (8) and have the potential to inhibit mitochondrial respiration (44). However, experimental evidence in permeabilized skeletal muscle fibers or isolated mitochondria suggests that the effects of these metabolic by-products are limited to certain nonphysiological conditions (nonphosphorylating mitochondria) (45) or only seem to influence the sensitivity of respiration to some substrates without affecting maximal ADP-stimulated respiration (46). It is therefore unclear to what extent the large metabolic disturbance occurring during a 5-km cycling time trial (8) would contribute to the impaired oxidative phosphorylation capacity observed here.

Muscle contraction generates heat, which has also the potential to modulate mitochondrial respiration. For instance, heat stress before muscle sampling decreased state 2 and state 3CI respiration without affecting muscle respiratory capacity (state 3CI + CII) in Notolabrus celidotus (47). Also, increasing the temperature of isolated mitochondria or permeabilized skeletal muscle fibers in vitro augmented state 2 respiration to a larger extent than state 3CI + CII, resulting in a diminished respiratory control ratio and suggesting structural damage to the mitochondria (47–49). These previous findings diverge from the decreased muscle respiratory capacity and state 3CII observed in the current study (Fig. 2). In addition, in this study, state 2 respiration was not significantly altered after the exercise, and only one sample (not included in the analysis) demonstrated an accelerated respiration rate with cytochrome c, implying that mitochondrial membrane integrity was preserved in these experimental conditions. Together, these findings suggest that it is unlikely that an exercise-induced increase in muscle temperature could explain the detrimental effect of high-intensity aerobic exercise on mitochondrial function.

A transient mitochondrial defect?

It should be noted that using the current experimental design, it cannot be determined if the apparent exercise-induced attenuation in muscle respiratory capacity was transient or permanent. However, on the basis of the animal literature (12), it is reasonable to assume that muscle respiratory capacity was restored within several hours. Also, it is well established that, across several days, multiple high-intensity exercise sessions ultimately improve muscle respiratory capacity (6,50). Interestingly, although the time course for the restoration of respiratory capacity after exercise in humans is still unknown, the molecular processes involved in mitochondrial functional recovery/improvement after exercise have been the focus of many studies. Specifically, after a single exercise session, a rapid (<4 h) upregulation of transcriptional factors such as peroxisome proliferator–activated receptor-γ coactivators 1α and 1β (PGC-1α and PGC-1β) and peroxisome proliferator–activated receptors β/δ (PPARβ/δ) occurs transiently. Thereafter (<24 h), this is followed by an upregulation of PGC-1α and PPARα protein expression, suggesting a rapid activation of the pathways responsible for mitochondrial biogenesis (51). It is, however, important to note that this effect is thought to be cumulative, such that successive sessions are required to increase muscle mitochondrial content (50,51). Concurrently, exercise stimulates the degradation of damaged cellular components from the mitochondria through autophagy. Some proteins involved in this process (light chain 3 I and II, and the autophagy adaptor protein p62) have been documented to be modulated within 2 h after an exercise bout (52). This exercise-induced autophagy is intensity dependent (53) and seems to be targeting oxidatively damaged proteins (52). Besides mitochondrial biogenesis and autophagy, exercise can also rapidly (~10 min) increase mitochondrial membrane interactions (28). However, changes in the abundance of key regulatory proteins involved in mitochondrial dynamic remodeling, again, seem to require multiple exercise sessions (51). Future studies investigating in parallel the time course of the restoration of mitochondrial function and these molecular mechanisms (biogenesis, mitochondrial dynamics, and autophagy) could provide valuable information that would improve our understanding of the adaptations in muscle in response to acute and chronic exercise.

Physiological and practical implications for muscle performance

Despite the large magnitude of the decrease in state 3CI + CII (~40%) immediately after high-intensity exercise, it is unlikely that the fall in mitochondrial respiratory capacity observed herein limited exercise performance during a 5-km cycling time trial. As illustrated in Figure 1, subjects maintained their V˙O2peak and even increased power output (8) during the last kilometer of the time trial, suggesting that the acute decrease in mitochondrial capacity was not a limiting factor for V˙O2max and exercise performance. This interpretation is also supported by several pieces of evidence collected in untrained individuals, suggesting that muscle mitochondrial respiratory capacity is in excess of peripheral O2 delivery during cycling (36). For instance, in agreement with the present results, it was recently reported that prolonged low-intensity skiing in the arctic for 6 wk decreased muscle mitochondrial respiratory capacity by approximately 20%, whereas peak leg and pulmonary V˙O2 remained unaltered (54). Therefore, taken together, these findings suggest that it is unlikely that exercise performance during the time trial was limited by a shortage in ATP generated by the mitochondria.

It also remains to determine whether the transient mitochondrial defect, recognized herein, may have indirectly contributed to the exercise-induced development of fatigue by exaggerating the accumulation of metabolites (e.g., inorganic phosphates, ADP, or protons); by increasing the firing of type III/IV afferent fibers (8), which are involved in the regulation of peripheral fatigue; or by potentiating other stress signals originating from the myocyte. In this regard, albeit using a very different experimental paradigm, it has recently been suggested that mitochondria have the potential to perturb whole-body physiological responses to stress and endocrine signals between organs via mechanisms that remain to be elucidated (55). Although still speculative at this point, this potential role of the mitochondria in the development of exercise-induced fatigue and as a mediator of the stress response to exercise is of interest and warrants further investigation.

Finally, the concept of “mitohormesis” postulates that stress induces an adaptive response and triggers defense mechanisms that prevent mitochondrial damage during a subsequent similar stress (56). With this concept in mind, the impairment in mitochondrial respiratory function observed immediately after high-intensity cycling exercise could trigger the dynamic remodeling of the mitochondrial network and promote the morphological changes reported after chronic exercise interventions. Therefore, although additional studies are warranted, mitochondrial hormesis may be one mechanism by which high-intensity exercise seems to be a suitable exercise training method when specifically targeting peripheral muscle adaptations.

EXPERIMENTAL CONSIDERATIONS

It is noteworthy that in this study the inhibition of respiration with combined antimycin A and oligomycin was used to correct for residual O2 consumption, indicative of nonmitochondrial O2 consumption (i.e., from chemical and instrument-related respiration in the chamber), which resulted in apparently lower respiration rates compared with previously published values. Without such a correction, the respiration rates for complex I + II substrates at baseline are within the range of values previously reported by our group for healthy adults using a similar protocol and the same instrument (27,57,58). We also conducted some additional experiments in permeabilized skeletal muscle fibers to compare the respiratory rates obtained with the Hansatech system and a widely used apparatus (O2k-core; Oroboros, Innsbruck, Austria) to determine whether the lower rates consistently observed by our group could be attributed to the instrumentation used. The muscle fibers for both experiments were obtained from the same sample of tissue (gastrocnemius muscle) and prepared similarly for the test by the same investigator. Other than initial [O2] (O2k: ~300 μM·mL−1; Hansatech: 180 μM·mL−1), the experimental conditions (temperature, 37°C) and substrate concentrations were the same. We found that the respiratory rate (state 3CI + II) was 1.7-fold higher with the O2k-core (64°pM·mg−1·s−1) compared with the Hansatech (37°pM·mg−1·s−1). This result strongly suggests that different instrumentation rather than muscle preparation contributed to the lower rates consistently reported by our group. Furthermore, using a similar preparation, we have previously reported a close correlation between the PCr recovery time constant measured by 31P-MRS and state 3CI + II (58), a significant correlation between uncoupledCIV respiration and CS activity (57), and an excess mitochondrial capacity in vitro compared with whole-body V˙O2peak (59). We are therefore very confident that the respiratory rates reported in this study suitably reflect muscle respiratory capacity in these in vitro experimental conditions.

CONCLUSIONS

The present study revealed that, largely as a consequence of attenuated complex II–specific respiration, skeletal muscle oxidative phosphorylation capacity is diminished immediately after high-intensity aerobic whole-body exercise in healthy active adults. This transient mitochondrial defect induced by high-intensity cycling exercise might amplify the exercise-induced development of fatigue and play an important role in initiating exercise-induced mitochondrial adaptations.

The authors would like to express their gratitude to subjects who gave their time and effort so generously to partake in this study.

This study was funded in part by the National Institutes of Health Heart, Lung, and Blood Institute (K99HL125756, HL-103786, HL-116579, HL-091830), National Institute of Aging (F32AG053009), the Flight Attendant Medical Research Institute, the Veterans Affairs Rehabilitation Research and Development Service (merit awards: E6910-R, E1697-R), SPiRe Awards (E1433-P), Senior Research Career Scientist (grant E9275-L), Career Development award (IK2RX001215), the American Heart Association (1850039, 14POST17770016), and the French Ministry of Higher Education (grant CIFRE 2012/0445).

No conflicts of interest, financial or otherwise, are declared by the authors. The results of the study are presented clearly, honestly, and without fabrication, falsification, or inappropriate data manipulation and do not constitute endorsement by the American College of Sports Medicine.

REFERENCES

1. Zoll J, Sanchez H, N’Guessan B, et al. Physical activity changes the regulation of mitochondrial respiration in human skeletal muscle. J Physiol. 2002;543(Pt 1):191–200.
2. Jacobs RA, Lundby C. Mitochondria express enhanced quality as well as quantity in association with aerobic fitness across recreationally active individuals up to elite athletes. J Appl Physiol. 2013;114(3):344–50.
3. Holloszy JO, Coyle EF. Adaptations of skeletal muscle to endurance exercise and their metabolic consequences. J Appl Physiol. 1984;56(4):831–8.
4. Hoppeler H, Howald H, Conley K, et al. Endurance training in humans: aerobic capacity and structure of skeletal muscle. J Appl Physiol. 1985;59(2):320–7.
5. Pesta D, Hoppel F, Macek C, et al. Similar qualitative and quantitative changes of mitochondrial respiration following strength and endurance training in normoxia and hypoxia in sedentary humans. Am J Physiol Regul Integr Comp Physiol. 2011;301(4):R1078–87.
6. Daussin FN, Zoll J, Dufour SP, et al. Effect of interval versus continuous training on cardiorespiratory and mitochondrial functions: relationship to aerobic performance improvements in sedentary subjects. Am J Physiol Regul Integr Comp Physiol. 2008;295(1):264–72.
7. Larsen FJ, Schiffer TA, Ortenblad N, et al. High-intensity sprint training inhibits mitochondrial respiration through aconitase inactivation. FASEB J. 2016;30(1):417–27.
8. Blain GM, Mangum TS, Sidhu SK, et al. Group III/IV muscle afferents limit the intramuscular metabolic perturbation during whole body exercise in humans. J Physiol. 2016;594(18):5303–15.
9. Gollnick PD, King DW. Effect of exercise and training on mitochondria of rat skeletal muscle. Am J Physiol. 1969;216(6): 1502–9.
10. Chen J, Gollnick PD. Effect of exercise on hexokinase distribution and mitochondrial respiration in skeletal muscle. Pflugers Arch. 1994;427(3–4):257–63.
11. Dohm GL, Huston RL, Askew EW, Weiser PC. Effects of exercise on activity of heart and muscle mitochondria. Am J Physiol. 1972;223(4):783–7.
12. Gollnick PD, Bertocci LA, Kelso TB, Witt EH, Hodgson DR. The effect of high-intensity exercise on the respiratory capacity of skeletal muscle. Pflugers Arch. 1990;415(4):407–13.
13. Ydfors M, Hughes MC, Laham R, Schlattner U, Norrbom J, Perry CG. Modelling in vivo creatine/phosphocreatine in vitro reveals divergent adaptations in human muscle mitochondrial respiratory control by ADP after acute and chronic exercise. J Physiol. 2016;594(11):3127–40.
14. Perry CG, Kane DA, Herbst EA, et al. Mitochondrial creatine kinase activity and phosphate shuttling are acutely regulated by exercise in human skeletal muscle. J Physiol. 2012;590(21):5475–86.
15. Madsen K, Ertbjerg P, Djurhuus MS, Pedersen PK. Calcium content and respiratory control index of skeletal muscle mitochondria during exercise and recovery. Am J Physiol. 1996;271(6 Pt 1):E1044–50.
16. Tonkonogi M, Harris B, Sahlin K. Mitochondrial oxidative function in human saponin-skinned muscle fibres: effects of prolonged exercise. J Physiol. 1998;510(Pt 1):279–86.
17. Tonkonogi M, Walsh B, Tiivel T, Saks V, Sahlin K. Mitochondrial function in human skeletal muscle is not impaired by high intensity exercise. Pflugers Arch. 1999;437(4):562–8.
18. Gnaiger E. Capacity of oxidative phosphorylation in human skeletal muscle: new perspectives of mitochondrial physiology. Int J Biochem Cell Biol. 2009;41(10):1837–45.
19. Helgerud J, Høydal K, Wang E, et al. Aerobic high-intensity intervals improve VO2max more than moderate training. Med Sci Sports Exerc. 2007;39(4):665–71.
20. Roels B, Thomas C, Bentley DJ, Mercier J, Hayot M, Millet G. Effects of intermittent hypoxic training on amino and fatty acid oxidative combustion in human permeabilized muscle fibers. J Appl Physiol (1985). 2007;102(1):79–86.
21. Bergstrom J. Muscle-biopsy needles. Lancet. 1979;1(8108):153.
22. Tevald MA, Lanza IR, Befroy DE, Kent-Braun JA. Intramyocellular oxygenation during ischemic muscle contractions in vivo. Eur J Appl Physiol. 2009;106(3):333–43.
23. Bendahan D, Chatel B, Jue T. Comparative NMR and NIRS analysis of oxygen-dependent metabolism in exercising finger flexor muscles. Am J Physiol Regul Integr Comp Physiol. 2017;313(6):R740–R53.
24. Rouslin W. Mitochondrial complexes I, II, III, IV, and V in myocardial ischemia and autolysis. Am J Physiol. 1983;244(6):H743–8.
25. Pesta D, Gnaiger E. High-resolution respirometry: OXPHOS protocols for human cells and permeabilized fibers from small biopsies of human muscle. Methods Mol Biol. 2012;810:25–58.
26. Kuznetsov AV, Veksler V, Gellerich FN, Saks V, Margreiter R, Kunz WS. Analysis of mitochondrial function in situ in permeabilized muscle fibers, tissues and cells. Nat Protoc. 2008;3(6):965–76.
27. Gifford JR, Trinity JD, Layec G, et al. Quadriceps exercise intolerance in patients with chronic obstructive pulmonary disease: the role of altered skeletal muscle mitochondrial respiration. J Appl Physiol (1985). 2015. jap 00460 2015.
28. Picard M, Gentil BJ, McManus MJ, et al. Acute exercise remodels mitochondrial membrane interactions in mouse skeletal muscle. J Appl Physiol (1985). 2013;115(10):1562–71.
29. Kunz WS, Kudin A, Vielhaber S, Elger CE, Attardi G, Villani G. Flux control of cytochrome c oxidase in human skeletal muscle. J Biol Chem. 2000;275(36):27741–5.
30. Kadenbach B, Huttemann M, Arnold S, Lee I, Bender E. Mitochondrial energy metabolism is regulated via nuclear-coded subunits of cytochrome c oxidase. Free Radic Biol Med. 2000;29(3–4):211–21.
31. Jacobs RA, Rasmussen P, Siebenmann C, et al. Determinants of time trial performance and maximal incremental exercise in highly trained endurance athletes. J Appl Physiol (1985). 2011;111(5):1422–30.
32. Rasmussen UF, Krustrup P, Bangsbo J, Rasmussen HN. The effect of high-intensity exhaustive exercise studied in isolated mitochondria from human skeletal muscle. Pflugers Arch. 2001;443(2):180–7.
33. Piper HM, Sezer O, Schleyer M, Schwartz P, Hütter JF, Spieckermann PG. Development of ischemia-induced damage in defined mitochondrial subpopulations. J Mol Cell Cardiol. 1985;17(9):885–96.
34. Gollnick PD, Piehl K, Saltin B. Selective glycogen depletion pattern in human muscle fibres after exercise of varying intensity and at varying pedalling rates. J Physiol. 1974;241(1):45–57.
35. Gollnick PD, Armstrong RB, Sembrowich WL, Shepherd RE, Saltin B. Glycogen depletion pattern in human skeletal muscle fibers after heavy exercise. J Appl Physiol. 1973;34(5):615–8.
36. Boushel R, Gnaiger E, Calbet JA, et al. Muscle mitochondrial capacity exceeds maximal oxygen delivery in humans. Mitochondrion. 2011;11(2):303–7.
37. Bailey DM, Lawrenson L, McEneny J, et al. Electron paramagnetic spectroscopic evidence of exercise-induced free radical accumulation in human skeletal muscle. Free Radic Res. 2007;41(2):182–90.
38. Saitoh S, Zhang C, Tune JD, et al. Hydrogen peroxide: a feed-forward dilator that couples myocardial metabolism to coronary blood flow. Arterioscler Thromb Vasc Biol. 2006;26(12):2614–21.
39. Nulton-Persson AC, Szweda LI. Modulation of mitochondrial function by hydrogen peroxide. J Biol Chem. 2001;276(26):23357–61.
40. Zini R, Berdeaux A, Morin D. The differential effects of superoxide anion, hydrogen peroxide and hydroxyl radical on cardiac mitochondrial oxidative phosphorylation. Free Radic Res. 2007;41(10):1159–66.
41. Walsh B, Tonkonogi M, Sahlin K. Effect of endurance training on oxidative and antioxidative function in human permeabilized muscle fibres. Pflugers Arch. 2001;442(3):420–5.
42. Sanz A, Caro P, Gomez J, Barja G. Testing the vicious cycle theory of mitochondrial ROS production: effects of H2O2 and cumene hydroperoxide treatment on heart mitochondria. J Bioenerg Biomembr. 2006;38(2):121–7.
43. Barbieri E, Agostini D, Polidori E, et al. The pleiotropic effect of physical exercise on mitochondrial dynamics in aging skeletal muscle. Oxid Med Cell Longev. 2015;2015:917085.
44. Jubrias SA, Crowther GJ, Shankland EG, Gronka RK, Conley KE. Acidosis inhibits oxidative phosphorylation in contracting human skeletal muscle in vivo. J Physiol. 2003;553(Pt 2):589–99.
45. Tonkonogi M, Sahlin K. Actively phosphorylating mitochondria are more resistant to lactic acidosis than inactive mitochondria. Am J Physiol. 1999;277(2 Pt 1):C288–93.
46. Walsh B, Tiivel T, Tonkonogi M, Sahlin K. Increased concentrations of P(i) and lactic acid reduce creatine-stimulated respiration in muscle fibers. J Appl Physiol. 2002;92(6):2273–6.
47. Iftikar FI, Hickey AJ. Do mitochondria limit hot fish hearts? Understanding the role of mitochondrial function with heat stress in Notolabrus celidotus. PLoS One. 2013;8(5):e64120.
48. Brooks GA, Hittelman KJ, Faulkner JA, Beyer RE. Temperature, skeletal muscle mitochondrial functions, and oxygen debt. Am J Physiol. 1971;220(4):1053–9.
49. Jarmuszkiewicz W, Woyda-Ploszczyca A, Koziel A, Majerczak J, Zoladz JA. Temperature controls oxidative phosphorylation and reactive oxygen species production through uncoupling in rat skeletal muscle mitochondria. Free Radic Biol Med. 2015;83:12–20.
50. Daussin FN, Rasseneur L, Bouitbir J, et al. Different timing of changes in mitochondrial functions following endurance training. Med Sci Sports Exerc. 2012;44(2):217–24.
51. Perry CG, Lally J, Holloway GP, Heigenhauser GJ, Bonen A, Spriet LL. Repeated transient mRNA bursts precede increases in transcriptional and mitochondrial proteins during training in human skeletal muscle. J Physiol. 2010;588(Pt 23):4795–810.
52. Halling JF, Ringholm S, Nielsen MM, Overby P, Pilegaard H. PGC-1α promotes exercise-induced autophagy in mouse skeletal muscle. Physiol Rep. 2016;4(3).
53. Brandt N, Dethlefsen MM, Bangsbo J, Pilegaard H. PGC-1α and exercise intensity dependent adaptations in mouse skeletal muscle. PloS One. 2017;12(10):e0185993.
54. Boushel R, Gnaiger E, Larsen FJ, et al. Maintained peak leg and pulmonary VO2 despite substantial reduction in muscle mitochondrial capacity. Scand J Med Sci Sports. 2015;25(4 Suppl):135–43.
55. Picard M, McManus MJ, Gray JD, et al. Mitochondrial functions modulate neuroendocrine, metabolic, inflammatory, and transcriptional responses to acute psychological stress. Proc Natl Acad Sci U S A. 2015;112(48):E6614–23.
56. Ristow M, Schmeisser K. Mitohormesis: promoting health and lifespan by increased levels of reactive oxygen species (ROS). Dose Response. 2014;12(2):288–341.
57. Park SY, Gifford JR, Andtbacka RH, et al. Cardiac, skeletal, and smooth muscle mitochondrial respiration: are all mitochondria created equal? Am J Physiol Heart Circ Physiol. 2014;307(3):H346–52.
58. Layec G, Gifford JR, Trinity JD, et al. Accuracy and precision of quantitative 31P-MRS measurements of human skeletal muscle mitochondrial function. Am J Physiol Endocrinol Metab. 2016;311(2):E358–66.
59. Gifford JR, Garten RS, Nelson AD, et al. Symmorphosis and skeletal muscle V˙O2 max: in vivo and in vitro measures reveal differing constraints in the exercise-trained and untrained human. J Physiol. 2016;594(6):1741–51.
Keywords:

OXIDATIVE PHOSPHORYLATION CAPACITY; ELECTRON TRANSPORT CHAIN; STATE 3 RESPIRATION; CYCLING TIME TRIAL; MITOHORMESIS

Copyright © 2018 by the American College of Sports Medicine