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Exercise Effects on Mitochondrial Function and Lipid Metabolism during Energy Balance


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Medicine & Science in Sports & Exercise: April 2020 - Volume 52 - Issue 4 - p 827-834
doi: 10.1249/MSS.0000000000002190
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Aerobic exercise training (AET) is well known to confer numerous cardiovascular and metabolic health benefits. These beneficial effects may be mediated in part by improvements in mitochondrial biogenesis and oxidative capacity (1,2) and upregulation of proteins related to lipid metabolism (3). These findings have implications in the treatment of chronic cardiometabolic diseases, as improved mitochondrial function has been linked to improved insulin sensitivity (4) and lower oxidative stress (5). However, AET can promote both acute and chronic energy deficits, and there are several independent effects of negative energy balance (EB) on mitochondrial function, including lower ROS emission and increased respiratory capacity (6,7). Thus, it remains to be determined if improvements in mitochondrial function observed after AET occur independent of negative EB.

Numerous studies have provided evidence to suggest the mechanisms by which AET can induce improvements in mitochondrial function. AET improves the activity of the electron transport system in rodents (8) and humans (9,10). In addition, a key adaptation to AET is increased mitochondrial content, which is thought to occur to meet the energetic demands of endurance training (1) and contribute to increased mitochondrial capacity. Mitochondrial biogenesis is induced in part by peroxisome proliferator-activated receptor-γ coactivator-1α (PGC1α) (11), a key protein implicated in mitochondrial function (12) and dynamics (13,14). AET is also known to enhance mitochondrial capacity to oxidize fatty acids (FA) (3) and can increase the storage of intramyocellular triacylglycerol (15). mRNA levels of key regulatory proteins in FA transport and metabolism, FA translocase (CD36) and carnitine palmitoyltransferase 1B (CPT1B), have been shown to be elevated after a short exercise program (16), and CD36 appears to be an important mediator in the increase in FA metabolism after exercise (17). However, many of these studies are confounded by samples obtained ≤24 h after the last exercise bout (16,18) or after weight loss during the AET (10). Given these limitations, it is unknown whether changes in mitochondrial oxidative capacity due to AET persist when measured in carefully controlled EB.

The acute and chronic energetic deficits created by AET interventions may play a role in the improvements observed in mitochondrial function attributed to exercise. Chronic caloric restriction is known to promote greater coupled respiration (19), likely to maximize efficiency of ATP production despite limited substrate availability. The acute depletion of energy reserves from a bout of aerobic exercise is known to promote increased O2 consumption, increased ATP synthesis (20), and decreased uncoupled mitochondrial respiration, likely driven by decreased uncoupling protein 3 (UCP3) (21). Whether weight loss alone is sufficient to alter mitochondrial oxidative capacity is debated (10); however, weight loss is well understood to decrease energy expenditure (22).

The purpose of this study was to assess the effects of AET on skeletal muscle mitochondrial function and markers of lipid metabolism when assessed 72 h removed from the last exercise bout and EB has been restored. We hypothesized that exercise-induced improvements in skeletal muscle mitochondrial function reported in previous literature would persist when EB is controlled. A secondary aim was to assess proteins associated with mitochondrial bioenergetics and lipid metabolism to gain insight into potential mechanisms that mediate changes in skeletal muscle mitochondrial function after AET.



This is a secondary analysis of a study designed to evaluate insulin sensitivity, resting energy expenditure, and blood pressure after a bout of high-intensity exercise (23–25). We recruited 14 participants between the age of 21 and 45 yr who self-identified as either of European American (EA) or African American descent (AA). Participants were not taking oral contraceptives or any medication known to influence blood pressure or metabolism. All participants were normotensive, nonsmokers, and normoglycemic, and they did not engage in routine physical activity (participating in less than one exercise activity per week). All testing was conducted in the first 10 d of the follicular phase of the menstrual cycle. All participants provided written, informed consent, and the Institutional Review Board at the University of Alabama at Birmingham approved this study.

Study design

The study design is depicted in Figure 1, and each method is outlined in further detail below. A skeletal muscle biopsy of the vastus lateralis was collected from each participant before beginning an AET program. All food was provided to the participants for 72 h before the tissue collection and was made up of ~60% carbohydrates, ~15% protein, and ~25% fat by calories. The participants spent the 23 h immediately before the tissue collection in our whole-room indirect calorimeter. Skeletal muscle tissue was used to measure mitochondrial oxidative function and protein levels. Posttraining evaluations took place after 8 to 16 wk of exercise training. Training duration varied because of the experimental design of the parent study in which there was a random assignment of test order over 8–16 wk exercise training. Only the results of the baseline tests (before initiation of training) and posttraining condition in which the subjects participated in no exercise for 72 h before evaluation are included. The other two conditions, either a bout of moderate-intensity or a bout of high-intensity cycle exercise 22 h before evaluation, are not included in this article (25). There were neither differences between the three posttraining conditions for V˙O2peak nor a test order effect for any of the cycling economy, or any of the mitochondrial respiration values, suggesting that duration of training did not affect any variable of interest. The same protocol was used before the collection of post-AET muscle biopsies. Again, participants were provided all food for 72 h, and the 23 h before tissue collection was spent in the whole-room calorimeter. Post-AET tissue was collected at least 72 h removed from the last exercise bout to assess the chronic effect of AET on mitochondrial oxidative function and protein levels in EB.

Study design.

EB and room calorimetry

Participants were provided with all food for 72 h before tissue collection. We used multiple regression to develop equations designed to maintain EB before and during the room calorimeter stay. These equations have previously been used in previous articles (23,25). Caloric intake for the 48 h before the room calorimetry visit was based on estimates generated from 330 doubly labeled water estimates of free-living energy expenditure of sedentary premenopausal women collected in our laboratory (26,27). The first equation was as follows: equation 1 = 750 kcal + [(31.47 × fat-free mass) – (0.31 × fat mass) – (155 × race, race coded 1 for AA and 2 for EA)]. An equation for estimating the room calorimeter energy intake was developed from over 200 room calorimeter visits of premenopausal women (28,29): equation 2 = 465 kcal + [(27.8 × fat-free mass) – (2.4 × fat mass) – (188 × race, race coded 1 for AA and 2 for EA)]. However, we recognized that the estimates may result in overfeeding or underfeeding individual subjects. Therefore, we developed a correction equation for the room calorimeter visit that was based on energy expenditure during the room calorimeter stay up to 5:30 pm. This equation was as follows: [equation 3 = 9(390 kcal + average energy expenditure in kilocalories per minute between 8:00 am and 5:30 pm) × 925 kcal) – equation 2 estimate of energy expenditure]. We then adjusted the food intake of the evening meal to match the results of equation.

AET program

Each participant trained 3 d·wk−1 for 8–16 wk and was monitored by trained exercise physiologists during each training session. Each exercise session included a 3- to 5-min warm-up and cooldown period of light activity. Participants were able to complete the AET using a treadmill, recumbent bike, or stationary bike (at least 50% of the training was performed by cycle ergometry, and the remainder of training was optional to the participants). During the first week, participants began training at 67% heart rate max (HRM) for 20 min, increasing time and intensity each week until the beginning of the fifth week, when each participant was training at 80% HRM for 40 min. This intensity was maintained for the duration of the AET program. We chose this volume of exercise, as we did not want participants to lose a significant amount of weight, to control for confounding effects that can occur with significant weight loss.


This protocol has been adapted from previously published work (27,30). The testing was done on a cycle ergometer with each participant pedaling at 50 W for 3 min. Every minute thereafter, the resistance was increased 20 W until the subject reached volitional exhaustion. Oxygen uptake and carbon dioxide production were continuously monitored using a MAXX-II metabolic cart (Physio-Dyne Instrument Corp., Massapequa, NY). Heart rate was measured using a POLAR Vantage XL heart rate monitor (Gays Mills, WI). The criteria for achieving V˙O2peak were heart rate within 10 bpm of estimated maximum, RER of at least 1.10, and plateauing of V˙O2. All subjects achieved at least one criteria, and all but three subjects achieved at least two criteria during each of the four tests.

Body fat quantification

Body composition (fat-free mass [kg] and fat mass [kg]) was assessed by dual-energy x-ray absorptiometry using a Lunar iDXA densitometer (GE-Lunar Corporation, Madison, WI). Participants wore light clothing and remained supine in compliance with normal testing procedures. Scans were analyzed with enCORE 2011 software (GE Healthcare Lunar, Madison, WI).

Laboratory analyses

Sera were analyzed by the Core Laboratory of the UAB Diabetes Research Center and Center for Clinical and Translational Science. Fasting glucose, total cholesterol, HDL cholesterol, triglycerides, and circulating free FA were measured using a SIRRUS analyzer (Stanbio Laboratory, Boerne, TX). LDL cholesterol was calculated using the Friedewald method (31). Fasting insulin was measured using a TOSOH AIA-II analyzer (TOSOH Corp., South San Francisco, CA).

Mitochondrial respiration measures

This technique has been adapted from previously published methods (32). Immediately after each stay in the room calorimeter (baseline and post-AET), a skeletal muscle biopsy of the vastus lateralis was collected under local anesthesia from each participant. The tissue was immediately cleaned of adipose and connective tissue, and a bundle of approximately 20 mg was selected for mitochondrial respirometry. This tissue was transferred to the laboratory on ice in Buffer X buffer containing 50 mM MES, 7.23 mM K2EGTA, 2.77 mM CaK2EGTA, 20 mM imidazole, 0.5 mM DTT, 20 mM taurine, 5.7 mM ATP, 14.3 mM PCr, and 6.56 mM MgCl2-6 H2O (pH 7.1, 290 mOsm). The tissue was then dissected into several smaller muscle bundles (of approximately 3–5 mg wet weight), and each was gently separated with a pair of antimagnetic needle-tipped forceps under magnification. Bundles were treated with 30 μg·mL−1 saponin in Buffer X and incubated on a rotator for 30 min at 4°C. The tissue bundles were washed of saponin for 15 min in Buffer Z containing 105 mM K-MES, 30 mM KCl, 1 mM EGTA, 10 mM K2HPO4, and 5 mM MgCl2-6 H2O, 5 μM glutamate, 2 μM malate, and 5.0 mg·mL−1 bovine serum albumin (pH 7.4, 290 mOsm). The samples were transferred to Buffer Z containing 20 mM creatine hydrate and 5 μM of blebbistatin for 10 min before respirometry experiments (32).

High-resolution respirometry was performed using an Oroboros Oxygraph O2K (Oroboros Instruments, Innsbruck, Austria) containing 2 mL of Buffer Z with creatine and blebbistatin, constantly stirred at 37°C. We assessed mitochondrial O2 consumption using two substrate protocols: 4 mM malate, 9 mM pyruvate, and 2.5 mM succinate (PMS) to drive convergent electron input to complexes I and II of the electron transport system or 2 mM malate and 40 μM palmitoyl carnitine (PC) to examine mitochondrial FA oxidation. State 3 in each substrate condition was measured after the addition of 1 mM ADP. Cytochrome c (10 μM) was added to assess mitochondrial membrane integrity after the dissection of the myofibers. There was no significant increase in respiration after the addition of cytochrome c. Uncoupled respiration (state 4) was induced by oligomycin (2 μg·mL−1). Maximal complex IV activity was assessed after the addition of ascorbate (2 mM) and tetramethyl-p-phenylenediamine (0.5 mM), providing a measure of mitochondrial content. The respiratory control ratio (RCR) was calculated as state 3/state 4. The RCR is a commonly used metric for assessing mitochondrial integrity and is highly correlated with coupling efficiency. Oxygen flux was normalized to either the wet weight or the complex IV activity of each fiber bundle.


A portion of muscle tissue collected at baseline and after the completion of the AET protocol was snap frozen in liquid N2 and stored at −80°C until analysis. Muscle samples of approximately 30 mg were powdered in a liquid N2-cooled mortar and pestle and homogenized in 6 μL·mg−1 muscle of ice-cold lysis buffer (150 mM NaCl, 50 mM Tris–HCl, 5 mM EDTA, 1% Triton X-100, 1% deoxycholate, 0.1% SDS, and 0.5% NP-40 at pH 7.4) with protease (Sigma P2714) and phosphatase (Sigma P0044) inhibitors. The samples were centrifuged at 15,000g for 15 min at 4°C, and the supernatant was collected. Protein content of the supernatant was quantified using a BCA Protein Assay Kit (Pierce Biotechnology, Rockford, IL). Thirty-seven micrograms of protein was treated with 4× NuPAGE LDS Sample Buffer (Novex, Carlsbad, CA) and 10× NuPAGE Reducing Agent (Novex) and incubated at 70°C for 10 min. The samples were electrophoresed in an SDS polyacrylamide gel (4%–20%) at 100 V on ice. The gels were blotted to polyvinylidene fluoride membranes using a semidry transfer method at 25 V for 12 min using a Pierce Power Blotter (Thermo Fisher Scientific, Waltham, MA). The membranes were blocked under conditions optimized for each antibody (2%–5% milk and/or 2%–5% bovine serum albumin in phosphate-buffered saline with 0.1% Tween 20 [PBST]) for 1 h at room temperature with gentle agitation. An appropriate dilution of primary antibody (Ab) was added for incubation overnight at 4°C with gentle agitation. We probed for the following: rabbit polyclonal Ab against CD36 (1:1000, sc-9154; Santa Cruz Biotechnology, Dallas, TX) and UCP3 (1:500, ab3477; Abcam, Cambridge, MA); rabbit monoclonal Ab against α-tubulin (1:1000, Cell Signaling, #2125) and CPT1B (1:1000, Abcam, ab134135); and a goat polyclonal Ab against PGC1α (1:1000, Abcam, ab106814). Horseradish peroxidase-conjugated secondary Abs were used at 1:50,000 in 0.5% of selected blocking agent in PBST for 1 h at room temperature with gentle agitation. Bands were visualized by chemiluminescent detection using ECL Western Blotting Substrate (Pierce Biotechnology) in a ChemicDoc XRS (Bio-Rad Laboratories, Hercules, CA). Band densitometry was quantified by Image Lab software (version 4.1) (Bio-Rad Laboratories). Values shown were obtained after normalization to the reference protein α-tubulin.

Statistical analyses

Not all data were available for all participants. Each table and figure accurately represents the number of participants assessed for each measure. Descriptive characteristics are reported as mean ± SD. Changes in variables between baseline and post-AET were assessed using two-tailed t-tests for paired samples and depicted as mean ± SD. An alpha level of 0.05 was used to determine statistical significance. All statistical analyses were conducted using SPSS Statistics for Macintosh version 22.0 (IBM Corp., Armonk, NY).


Demographic, physical, and metabolic characteristics of the study participants at baseline and after AET are shown in Table 1. Body mass index (BMI) ranged from 20.1 to 35.0 kg·m−2 and age from 21 to 40 yr. There were 8 EA participants and 6 AA participants. There were no significant changes in weight, BMI, or body fat between baseline and post-AET conditions. There was a 0.6 kg increase in fat-free mass (P < 0.05). V˙O2peak improved significantly after the AET protocol (P < 0.05). There were no differences in energy intake and energy expenditure during the stay in the room calorimeter (Fig. 2).

Clinical characteristics of participants (n = 14).
Energy intake, energy expenditure, and difference calculated during room calorimetry. n = 12. No significant differences between baseline and posttraining.

After AET, there were no changes in mitochondrial respiration normalized to fiber weight when supported by PMS (Fig. 3A), but there were significant increases in both state 3 and state 4 respiration rates supported by PC (Fig. 3B). There were no changes in the RCR after AET supported by PMS or PC (Fig. 3C). When mitochondrial respiration measures were normalized to maximal complex IV activity (a marker of mitochondrial content), there were no changes in mitochondrial function after AET supported by either PMS (Fig. 4A) or PC (Fig. 4B).

Oxygen consumption under state 3 and state 4 conditions normalized to fiber weight supported by PMS (A) and PC (B). C, RCR supported by PMS or PC. n = 11, *P < 0.05.
Oxygen consumption under state 3 and state 4 conditions normalized to maximal complex IV activity supported by PMS (A) and PC (B), and maximal complex IV activity supported respiration PMS (C) and PC (D). n = 12.

There were no changes in any proteins measured after the AET program when assessed under energetic balance (PGC1α, UCP3, CD36, or CPT1B) (Fig. 5).

Skeletal muscle protein expression at baseline and post-AET was measured using Western blotting. Values are normalized to α-tubulin and are presented as mean ± SEM. PGC1α, n = 11 (A); UCP3, n = 11 (B); CD36, n = 11 (C); CPT1B, n = 5 (D).


Mitochondrial oxidative capacity and content are known to improve after AET (1,2). This study tested the hypothesis that these improvements would persist 72 h after the last exercise bout when EB had been restored. There were no differences between caloric intake and energetic expenditure in the 23 h before each tissue collection and no changes in weight or REE during the AET. We demonstrated that mitochondrial respiratory capacity supported by PMS was unchanged when measured in an energetically balanced state after AET. However, when mitochondrial oxidation was supported by an FA substrate, we observed an enhanced mitochondrial oxidative capacity and elevated uncoupled respiration. These changes were no longer apparent when these data were normalized for a marker of mitochondrial content, suggesting that the improvements in mitochondrial capacity were due to mitochondrial biogenesis induced by AET, and not intrinsic changes to existing mitochondria. Finally, there were no changes in proteins known to mediate mitochondrial biogenesis or lipid transport and metabolism.

The results from this study suggest that improvements in mitochondrial FA oxidation may be due to primarily mitochondrial biogenesis. However, we cannot rule out other potential mechanism that may be responsible for changes in FA oxidation. Chronic AET is well understood to induce mitochondrial biogenesis. Improvements in mitochondrial content have been observed after only 7 wk of training (33) but can be reversed in as little as 6 wk after the conclusion of an exercise program (34). Acute exercise is known to promote enhanced expression of p38 MAPK and, thus, activation of PGC1α, an important signaling cascade for mediating mitochondrial biogenesis (35). Despite this, we found no increase in PGC1α protein levels after AET. However, this is likely because increases in PGC1α are transient, and protein levels in this study were measured in EB and more than 72 h after the last exercise bout. This is supported by two previous studies in which transient increases were observed immediately after successive exercise bouts (36) and 24 h after a single exercise bout (37). We are aware of no study that has observed increased mitochondrial content after an acute bout of exercise. We hypothesize that mitochondrial biogenesis due to exercise is regulated in a more chronic manner and that transient increases in PGC1α from acute AET are likely sufficient to induce mitochondrial biogenesis associated with chronic AET.

All other proteins measured in the present study are known to be induced by AET; however, like PGC1α, no changes were apparent when assessed in EB and 72 h removed from the last exercise bout. UCP3 expression is increased after AET in rodents, but only as a function of increased mitochondrial content (38). This study did not consider the effect of EB on UCP3 expression. Elevated expression of mRNA for both CD36 and CPT1B has been observed when measured after a short exercise training program (16). In addition, previous investigators have shown that CD36 is an important mediator in the increase in FA oxidation after exercise, independent of mitochondrial biogenesis (17). However, it is possible that the timing of sample collection plays an important role in mediating these results, as others have also found (in agreement with our findings) no changes in CD36 or CPT1B protein expression when measured 24 h after an exercise bout (39).

The increase in mitochondrial oxidative capacity (state 3) measured under an FA substrate may reflect a greater capacity to use FA substrate prompted by AET. In addition, respiration uncoupled from ATP synthesis (state 4) was increased when supported by FA substrate after AET. Uncoupled respiration is commonly thought to be a mechanism to limit the production of mitochondrial-derived reactive oxygen species (ROS), and it is known to be induced by both FA substrate (40) and ROS production (41). Given that there were no changes in the RCR, the overall proportion of coupled to uncoupled respiration remained similar before and after AET, and thus overall efficiency was unchanged. Greater mitochondrial FA oxidative capacity may have the potential to decrease cellular storage of FA in the form of intramyocellular triglyceride (42) or ceramides and diacylglycerols (43) and prevent the cardiometabolic complications associated with them.

There were several strengths associated with the present study design. The AET program required participants to attend three sessions per week in our training facility, and trained personnel monitored the participants during each session. The provision of all food consumed by the participants for 72 h before the collection of each biopsy sample and 23-h energy expenditure measured by room calorimeter ensured a high degree of control over EB. We acknowledge that a primary limitation within this study was the lack of a true control group for which EB was not controlled. In addition, there are several methods with which to quantify mitochondrial content not used in the present study (e.g., mtDNA copy number, citrate synthase activity). These measures may have provided additional insight into the mechanisms of mitochondrial biogenesis that appear to be responsible for improvements in mitochondrial capacity. In addition, these data may only reflect the population in which they were collected (generally healthy premenopausal women), and not other subgroups, including men, aging persons, or those with cardiometabolic disease.

In conclusion, an 8- to 16-wk AET program was sufficient to improve mitochondrial respiratory capacity under an FA substrate load even when measured in EB, as evidenced by no weight loss during the AET. After normalization of respiratory capacity to a marker for mitochondrial content, these changes were no longer apparent, suggesting that the improvements in mitochondrial capacity after AET were primarily due to mitochondrial biogenesis.

This work was supported by the National Institutes of Health (2R01DK049779-11A1) and the UAB Center for Exercise Medicine (5T32HD071866-04). The results of this study are presented clearly, honestly, and without fabrication, falsification, or inappropriate data manipulation. The results of this study do not constitute endorsement by the American College of Sports Medicine. The authors gratefully acknowledge the work of the UAB Bioanalytical Redox Biology Core and the Clinical and Translational Science and the Clinical Research Unit for their assistance with the muscle biopsies. In addition, they thank Mr. David Bryan and Mr. Brandon Kane for their work coordinating this study.

Author contributions: J. W. drafted the manuscript, participated in data collection, and assisted with data analysis. G. H. designed the study and conducted statistical analysis. S. W. conducted the muscle biopsies. B. G. assisted with study design and data analysis. D. R. M. assisted with the study design and participated in data collection. G. F. drafted the manuscript, assisted with the study design, participated in data collection, and conducted statistical analysis. All authors reviewed and critically revised the submitted manuscript. The authors declared no conflict of interest.


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