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Autophagy and Protein Turnover Signaling in Slow-Twitch Muscle during Exercise


Medicine & Science in Sports & Exercise: July 2014 - Volume 46 - Issue 7 - p 1314–1325
doi: 10.1249/MSS.0000000000000237
Basic Sciences

Purpose: The aim of this study was to characterize skeletal muscle protein breakdown and mitochondrial dynamics markers at different points of endurance exercise.

Methods: Mice run at 10 m·min−1 during 1 h, and running speed was increased by 0.5 m·min−1 every minute during 40 min and then by 1 m·min−1 until exhaustion. Animals were killed by cervical dislocation at 30, 60, 90, and 120 min; at time to exhaustion (Te); and at 3 and 24 h during recovery. The soleus and the deep red regions of the quadriceps muscles were pooled.

Results: AMPK phosphorylation (Thr172) increased from 30 min to Te, and FoxO3a phosphorylation (Thr32 and Ser253) decreased from 120 min to 3 h after exercise. FoxO3a-dependent E3 ligases Mul1 and MuRF1 proteins increased from 30 min to Te and at Te and 3 h after exercise, respectively, whereas MAFbx/atrogin-1 protein expression did not change significantly. The autophagic markers LC3B-II increased at 120 min and Te, and p62 significantly decreased at Te. The AMPK-dependent phosphorylation of Ulk1 at Ser317 and Ser555 increased from 60 min to Te and at 30 and 60 min, respectively. Akt (Ser473), MTOR (Ser2448), and 4E-BP1 (Thr37/46) phosphorylation decreased from 90 min to Te, and the MTOR-dependent phosphorylation of Ulk1 (Ser757) decreased from 120 min to Te. Ser616 phosphorylation of the mitochondrial fission marker DRP1 increased from 60 min to Te, but protein expression of the fusion markers mitofusin-2, a substrate of Mul1, and OPA1 did not significantly change.

Conclusions: These results fit with a regulation of protein breakdown triggered by FoxO3a and Ulk1 pathways after AMPK activation and Akt/MTOR inhibition. Furthermore, our data suggest that mitochondrial fission is quickly increased, and mitochondrial fusion is unchanged during exercise.

1Faculty of Sport Sciences, University of Montpellier 1, Montpellier, FRANCE; and 2INRA, UMR866 Dynamique Musculaire Et Métabolisme, University of Montpellier 1, Montpellier, FRANCE

Address for correspondence: Anthony M.J. Sanchez, Ph.D., Faculty of Sport Sciences, University of Montpellier 1, 700 avenue du Pic Saint Loup, 34090 Montpellier, France; E-mail:

Submitted for publication August 2013.

Accepted for publication November 2013.

Acute and chronic exercises represent stimuli able to generate profound modulations of cellular signaling mechanisms to promote the accomplishment of specific metabolic and morphological adaptations. According to the intensity considered, acute exercise can induce an alteration in skeletal muscle protein turnover and cause damage to cell constituents. Two major signaling pathways are mainly involved: the PI3K/Akt/MTORC1 pathway associated with protein synthesis and the forkhead box O (FoxO)–related pathway implicated in the control of protein breakdown (30,35).

The mechanistic target of rapamycin complex 1 (MTORC1) modulates protein synthesis by regulating its downstream effectors, the eukaryotic translation initiation factor 4E-binding protein 1 (4E-BP1) (1), and the ribosomal protein S6 kinase 1 (S6K1) (10). Phosphorylation of 4E-BP1 at Thr37/46 by MTOR induces its dissociation from the eukaryotic translation initiation factor 4E (eIF4E), providing the assembly of the preinitiation complex (39). In addition, after an initial phosphorylation by MTOR at Thr389, S6K1 phosphorylates substrates involved in the regulation of protein translation, such as the ribosomal protein S6 (rpS6) and eIF4B (13,29).

Conversely to this pathway, FoxO transcription factor–related pathway plays a major role in the control of protein breakdown through the regulation of the ubiquitin–proteasome and autophagy–lysosomal systems (24,34). FoxO1 and FoxO3a are required for the transcription of the E3 ubiquitin ligases MAFbx/atrogin-1 and MuRF1 (16,34), leading to the ubiquitination and proteasomal degradation of several proteins involved in skeletal muscle maintenance. MAFbx/atrogin-1 catalyzes the breakdown of the myogenic transcription factors MyoD, myogenin, the eukaryotic initiation factor of translation eIF3f, as well as some sarcomeric proteins (i.e., desmin and vimentin) (22,33). MuRF1 seems essentially implicated in the degradation of myofibrillar proteins like myosin heavy chain protein (6), myosin-binding protein C, and myosin light chain 1 and 2 to date (8). The second system is the autophagy–lysosomal pathway that constitutes a fundamental mechanism for cell maintenance through the degradation of cytoplasmic organelles, soluble proteins and macromolecules, and the recycling of the breakdown products (21). It implicates firstly the sequestration of substrates into a vacuolar system (autophagosome) and secondly their degradation by lysosomal hydrolases. Autophagy requires the Atg (autophagy-related gene) proteins involved in the formation of autophagosomes (7) and two ubiquitin-like conjugation systems. The first one is the Atg12/Atg5/Atg16 complex, which is essential for the formation of autophagosome membrane (38). The second one involves Atg8, also known as microtubule-associated protein 1 light chain 3 (LC3) in mammals. Pro-LC3 is first cleaved by the cysteine protease Atg4 to its mature form, LC3I. This protein is then lapidated by Atg7 and Atg3 to form LC3II, contributing to the membrane fusion and to the substrate selection (27). Two other complexes are also important for the initiation of the autophagic process, the unc-51–like kinase (Ulk1)/Atg1 and the Beclin1/vacuole protein sorting 34 (Vps34)/PI3K complexes (17,25). Importantly, MTOR phosphorylates Ulk1 at Ser757, preventing the initiation of autophagy (15,18). MTORC1 has been proposed as the dominant regulator of autophagy initiation in skeletal muscle (5).

Endurance exercise induces muscular remodeling especially through the AMP-activated protein kinase (AMPK), a serine/threonine protein kinase that acts as a sensor of cellular energy status switch regulating several systems including glucose and lipid metabolism (31). AMPK has been implicated in the control of skeletal muscle protein turnover by decreasing MTORC1 activity and by increasing protein breakdown through regulation of ubiquitin–proteasome and autophagy pathways (32). AMPK can increase FoxO3a-dependent protein expression and also phosphorylate Ulk1 at several sites in vitro (18,32). However, little is known about the involvement of muscle protein breakdown systems that occurred in physiological conditions during endurance exercise. Concerning mitochondrial dynamics, a recent study highlights that a 24-h ultra endurance exercise rises the phosphorylation of the dynamin-related protein 1 (DRP1), a key actor of mitochondrial and peroxisomal division, without affecting the protein expression pattern of several mitophagic markers (i.e., PINK1, Parkin, and MFN1) (14). Fission and selective fusion govern mitochondrial segregation as well as elimination by autophagy and an increase in mitochondrial fission has been shown to be permissive for the induction of mitophagy (37). Recently, the involvement of FoxO3a in mitophagy has been characterized. During muscle-wasting stimuli, enhanced FoxO3a activity results in increased transcription of the mitochondrial E3 ubiquitin protein ligase 1 (Mul1), which in turn ubiquitinates and degrades the mitofusin-2 (MFN2), a mitochondrial fusion protein critical for the maintenance and genomic stability of mitochondrial DNA (23). These events promote fragmentation, depolarization, and selective elimination of damaged mitochondria through the autophagy–lysosomal pathway (23).

The present study aimed to characterize the modulation of skeletal muscle protein breakdown and mitochondrial dynamics markers during the progression of endurance exercise and to evaluate whether these regulations were coordinated with potential modulations of protein synthesis markers.

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Ethics statement

This study was approved by the Committee on the Ethics of Animal Experiments of the Languedoc Roussillon in accordance with the guidelines from the French National Research Council for the Care and Use of Laboratory Animals (Permit Number: CEEA-LR-1069). This study is in adherence to the American College of Sports Medicine animal care standards. All efforts were made to minimize suffering.

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Experiments were carried out on 8-wk-old C57BL/6 mice (n = 52; body mass = 26.2 ± 2.1 g) from our own stock. Animals were maintained on a 12-h/12-h light–dark cycle and provided with food and water ad libitum. Experiments were performed at 22°C.

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Exercise protocol and muscle collection

The endurance exercise protocol was performed on a motor-driven treadmill (Exer-6M Treadmill; Columbus Instruments). Mice were familiarized to treadmill running during three sessions in which they successively run at 5 m·min−1 during 1 min, 7 m·min−1 during 4 min, and 10 m·min−1 during 2 min. Three days after the familiarization period, mice performed, except a control group (n = 8), an endurance exercise until exhaustion. Animals exercised exactly at the same time of the day (i.e., 8:00 a.m.) in the fed state. Mice run at 10 m·min−1 during 1 h, and running speed was increased by 0.5 m·min−1 every minute during 40 min and then by 1 m·min−1 until exhaustion. Mice were killed by cervical dislocation at 30, 60, 90, and 120 min (n = 5 for each time point); time to exhaustion (Te) during exercise; and 3 and 24 h after exercise (n = 8 for each time point). The soleus and the deep red regions of the quadriceps (slow-twitch) muscles of both the left and right hind limbs were rapidly dissected out at the corresponding time and were frozen in liquid nitrogen before storage at −80°C for subsequent analyses. Mice that exercised until exhaustion ran 1957.7 ± 347.8 m, with an effective running time of 151.2 ± 14.8 min.

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Proteins isolation and immunoblot analysis

Muscles samples (i.e., soleus and deep red regions of the quadriceps) were pooled and homogenized in 10 volumes of lysis buffer (50 mM Tris–HCl pH 7.5, 150 mM NaCl, 1 mM EGTA, 100 mM NaF, 5 mM Na3VO4, 1% Triton X-100, 40 mM β-glycerophosphate, and protease inhibitor mixture [P8340; Sigma-Aldrich]) using FastPrep-24 Instrument (MP Biomedicals). Cellular debris was removed by centrifugation at 10,000g for 15 min (4°C). Protein concentrations were measured using a Pierce BCA protein assay kit (23225; Thermo Scientific). Proteins (60 μg) were denatured and loaded onto 7% and 15% SDS–polyacrylamide gels before electrophoretic transfer onto a nitrocellulose membrane (Bio-Rad). After transfer, membranes were blocked with 50 mM Tris–HCl pH 7.5, 150 mM NaCl, and 0.1% Tween 20 (TBS-T) containing 5% skimmed milk or BSA and incubated overnight at 4°C with primary antibodies. Membranes were washed three times for 10 min with TBS-T and incubated for 1 h with a peroxidase-conjugated secondary antibody (Sigma-Aldrich). Membranes were washed again three times for 5 min, and immunoblots were revealed by using a Pierce ECL kit (32106; Thermo Scientific) according to the manufacturer’s instructions.

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Anti–phospho-AMPKα (Thr172), AMPKα, phospho-FoxO3a (Thr32), phospho-FoxO3a (Ser253), FoxO3a, phospho-Ulk1 (Ser317), phospho-Ulk1 (Ser555), phospho-Ulk1 (Ser757), Ulk1, phospho-DRP1 (Ser616), DRP1, phospho-Akt (Ser473), Akt, phospho-MTOR (Ser2448), MTOR, phospho-4E-BP1 (Thr37/46), 4E-BP1, phospho-eIF2α (Ser51), eIF2α, phospho-rpS6 (Ser240/244), rpS6, and p62 were purchased from Cell Signaling; anti-Mul1 and VDAC were obtained from Abcam; anti-MFN2 and ubiquitin from Santa Cruz; anti–α-tubulin (DM1A) and LC3 from Sigma-Aldrich; and anti–MAFbx/atrogin-1 and MuRF1 from ECM Biosciences. The OPA1 antibody was a generous gift from Drs. Béatrice Chabi (INRA UMR 866, DMEM, Montpellier, France) and Guy Lenaers (INSERM U-583, Institut des Neurosciences de Montpellier, France).

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mRNAs isolation and reverse transcription-quantitative polymerase chain reaction (RT-qPCR) analysis

Muscles (i.e., soleus and deep red regions of the quadriceps) were pooled and crushed in liquid nitrogen using mortar and pestle. Total RNA was extracted from powdered muscles by using RNeasy Fibrous Tissue Kit (Qiagen), according to the manufacturer’s instructions. RNA concentration was determined by spectrophotometric analysis, and integrity was checked by OD260nm/OD280nm absorption ratio (>1.95) and by agarose gel analysis. Total RNAs were stored at −80°C until further use. Reverse transcription reaction was performed with 1 μg of total RNA using RevertAid First Strand cDNA synthesis kit (Thermo Scientific). First-strand cDNA synthesis was made by using 200 U of SuperScript II Reverse Transcriptase and Oligo(dT) 12–18 primers (Invitrogen) according to the manufacturer’s instructions. cDNAs were diluted (1:10) in water and stored at −20°C until further use. qPCR analysis was performed in a MiniOpticon detection system (Biorad, Hercules, CA) with 10 μL of KAPA SYBR Fast Universal Readymix (Thermo Scientific), 300 nM of both forward and reverse primers, 2 μL of diluted cDNA template and water to a final volume of 20 μL. The primers were designed using Universal ProbeLibrary Assay Design Center (Roche Applied System) and RT Primer Data Base. All PCRs were performed in duplicate using the following cycle parameters: 30 s at 98°C, 40 cycles of 1 s at 95°C and 15 s at 60°C. Relative mRNA expression levels were normalized to ribosomal protein S9 (RPS9) expression, which was unaffected by treatments. Primer sequences: 5′-FoxO3a GGAAATGGGCAAAGCAGA, 3′-FoxO3a AAACGGATCACTGTCCACTTG, 5′-MAFbx AGTGAGGACCGGCTACTGTG 3′-MAFbx GATCAAACGCTTGCGAATCT, 5′-MuRF1 TCCTGCAGAGTGACCAAGG, 3′-MuRF1 GGCGTAGAGGGTGTCAAAC, 5′-Mul1 AGGGCATTCTTTCAGAAGCA, 3′-Mul1 GGGGTGGAACTTCTCGTACA, 5′-RPS9 CGGCCCGGGAGCTGTTGACG, 3′-RPS9 CTGCTTGCGGACCCTAATGT.

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Densitometry analyses of immunoblots were performed using ImageJ software and were normalized to α-tubulin exposure. The statistical analyses were performed using STATISTICA 9.1 software (StatSoft France). All data are expressed as the mean ± SEM. Data were evaluated by one-way ANOVA followed by Fisher’s honestly post hoc test. Significance was declared when P< 0.05.

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Regulation of AMPK and FoxO3a

We first evaluated AMPK and FoxO3a phosphorylation at the different points of the exercise (Fig. 1). We found a quick increase by 51% ± 27% in AMPK phosphorylation on Thr172 at 30 min, and this level of phosphorylation remained elevated, i.e., +50% ± 18%, +75% ± 15%, +101% ± 14%, and +126% ± 28% at 60 min, 90 min, 120 min, and Te, respectively, when compared to non-exercised mice. AMPK phosphorylation returned to baseline at 3 h after exercise (Fig. 1A). Phosphorylation of FoxO3a at Thr32 and Ser253 was significantly decreased at 120 min (−52% ± 17% and −47% ± 13%, respectively), Te (−48% ± 11% and −30% ± 12%, respectively), and at 3 h after exercise (−48% ± 13% and −33% ± 6%, respectively; Figs. 1C and D). The total form of FoxO3a was significantly increased at 3 h after exercise (+94% ± 27%; Fig. 1E), whereas the total form of AMPK remained unchanged throughout the protocol (Fig. 1B). We also assessed FoxO3a mRNA level and found a significant increase at Te (+173% ± 55%) and at 3 h after exercise (+268% ± 139%; Fig. 2A).

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Ubiquitin–proteasome system markers

We next characterized the mRNA levels of E3 ubiquitin ligases transcriptionally regulated by FoxO3a (Fig. 2). We found an increase in MuRF1 at Te (+300% ± 113%) and at 3 h after exercise (+510% ± 90%; Fig. 2C) and an increase in MAFbx/atrogin-1 at 3 h after exercise (+179% ± 54%; Fig. 2D). Nonetheless, no change in Mul1 mRNA level was found (Fig. 2B). We also determined the protein expression of these three E3 ligases (Fig. 3) and found a significant increase in MuRF1 protein expression at Te (+104% ± 40%) and 3 h after exercise (+192% ± 74%) and an increase in Mul1 protein expression from 30 min to Te (Figs. 3B and A, respectively). These rises were +71% ± 23%, +118% ± 25%, +78% ± 7.9%, +123% ± 41%, and +99% ± 25% at 30 min, 60 min, 90 min, 120 min, and Te, respectively. However, we did not detect any change in MAFbx/atrogin-1 protein level (Fig. 3C). Furthermore, the ubiquitin-conjugated proteins levels were significantly increased by 52% ± 13%, 69% ± 22%, 48% ± 12%, 96% ± 19%, 125% ± 15%, and 52% ± 23% at 30 min, 90 min, 120 min, Te, 3 h, and 24 h after exercise (Fig. 3D).

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Autophagy system markers

The expression of several markers of autophagy pathway is depicted in Figure 4. We found an increase in LC3B-II/LC3B-I ratio at 120 min (+131% ± 57%) and Te (+100% ± 47%), suggesting an induction of autophagy during exercise (Fig. 4A). In addition, we assessed the protein level of p62, also called sequestosome 1 (SQSTM1). p62 is a ubiquitin-binding scaffold protein that binds directly to LC3 to facilitate degradation of ubiquitinated protein aggregates by autophagy–lysosomal machinery (28). As LC3, p62 is itself degraded by lysosome. Since p62 accumulates when autophagy is inhibited, and decreased when autophagy is induced, p62 is commonly used as a marker to characterize autophagic flux (3). In accordance with LC3 analysis, p62 protein expression was significantly reduced at Te (−46% ± 11%) and a tendency was observed at 120 min (P = 0.07; Fig. 4B). Moreover, a significant increase in p62 protein expression has been found at 24 h after exercise (+65% ± 23%). Next, we assessed the phosphorylation level of Ulk1 Ser317, Ser555, and Ser757 that are known to be phosphorylated by AMPK for the first two sites and by MTOR for the last one (9,18). We reported an increase in the phosphorylation of Ser317, i.e., +110% ± 31%, +77% ± 21%, +133% ± 55%, and +132% ± 36% at 60 min, 90 min, 120 min, and Te, respectively, and an increase in Ser555 phosphorylation at 30 min (+100% ± 26%) and 60 min (+76% ± 21%) of exercise (Figs. 4C and D, respectively). A decrease in Ulk1 Ser757 phosphorylation was found at 120 min of exercise (−44% ± 26%) and at Te (−59% ± 12%), whereas an increase was obtained at 24 h after exercise (+69% ± 22%; Fig. 4E). The total form of Ulk1 was significantly increased at 90 min (+33% ± 14%), 120 min (+38% ± 21%), Te (+100% ± 39%), and at 3 h after exercise (+54% ± 40%; Fig. 4F).

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Mitochondrial network remodeling markers

Next, we evaluated the modulation of major markers of mitochondrial remodeling: the phosphorylation state of DRP1 (Ser616), which is known to affect mitochondrial morphology through stimulation of mitochondrial fission, the protein levels of MFN2, a substrate of Mul1, and OPA1. The last two proteins are critical regulators of mitochondrial homeostasis because they are involved in mitochondrial fusion. Figs. 5A and B show a rise in the phosphorylation of DRP1 on Ser616 by 143% ± 63%, 178% ± 78%, 255% ± 127%, and 290% ± 108% at 60 min, 90 min, 120 min, and Te, respectively, without any change in the expression of its total form. Concerning MFN2 and OPA1, we did not find any significant variation of their protein content throughout the exercise and during recovery (Figs. 5C and D, respectively).

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Protein synthesis markers

Last, we examined several major components of the Akt/MTORC1 pathway to establish whether endurance exercise may induce a decrease in protein synthesis flux (Fig. 6). We found that exercise induces hypophosphorylation of Akt Ser473 at 90 min, 120 min, and Te (−53% ± 12%, −38% ± 12%, and −52% ± 17%, respectively); MTOR Ser2448 at 90 min, 120 min, and Te (−33% ± 13%, −36% ± 5%, and −35% ± 11%, respectively); and 4E-BP1 also at 90 min, 120 min, and Te (−43% ± 13%, −59% ± 8%, and −61% ± 8%, respectively; Figs. 6A, B, and C, respectively). In addition, we assessed the Ser240/244 phosphorylation of the direct target of S6K1, rpS6, but we did not find any significant modulation, compared to preexercise condition (Fig. 6D). In sum, these results suggest that the translational machinery could be less functional, especially for the latest points of the exercise. We also assessed the phosphorylation state of the regulatory subunit of the eukaryotic initiation factor 2 (eIF2), a heterotrimeric complex that mediates the binding of tRNAmet to the ribosome in a GTP-dependent manner (Fig. 6E). The α-subunit of this factor contains the main target for phosphorylation (Ser51) and is considered as the regulatory subunit of eIF2, with its phosphorylation leading to a stabilization of the eIF2–GDP–eIF2B complex that inhibits the turnover of eIF2B (36). Thus, phosphorylation of eIF2α at Ser51 is associated with the inhibition of protein synthesis. As expected, a significant increase in the phosphorylation of this residue occurred at 60 min, 90 min, 120 min, and Te (+137% ± 29%, +101% ± 38%, +144% ± 26%, and +195% ± 25%, respectively).

Furthermore, phosphorylation of MTOR and 4E-BP1 was significantly increased at 3 h (+37% ± 11% and +93% ± 29%, respectively) and at 24 h (+32% ± 15% and +48% ± 32%, respectively) after exercise when compared to non-exercised mice.

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In this study, we showed that autophagy and ubiquitin–proteasome markers, notably Mul1 protein expression, are upregulated during endurance exercise, coordinately with a downregulation of the protein synthesis pathway. While autophagy markers and phosphorylation of the mitochondrial fission marker DRP1 returned to basal level after exercise, ubiquitin–proteasome machinery markers, especially MuRF1 mRNA and protein levels, reached a peak at 3 h after exercise.

AMPK plays a major role in skeletal muscle homeostasis in response to energy stress conditions, including exercise (31). It was therefore not surprising to observe an increase in the phosphorylation state of AMPK on Thr172, which witnesses the kinase activation in response to the exercise. In our study, during the first 60 min corresponding to a low-intensity exercise, we were able to detect a fast increased AMPK phosphorylation. The low level of aerobic fitness of the sedentary mice used in the present study may contribute to explain the activation of the kinase from such a low intensity, since it is known that AMPK activation depends on the training status (26). Concerning FoxO3a, we found a decrease of Ser253 and Thr32 phosphorylation (i.e., two Akt-linked inhibiting phosphorylation) from 120 min of exercise and during the recovery period (i.e., at 3 h after exercise). This result suggests that FoxO3a is activated during exercise, especially when intensity is near to exhaustion, and also during the first hours of the recovery period. Accordingly, FoxO3a mRNA and protein levels were markedly increased at 3 h after exercise. Furthermore, among FoxO posttranslational modifications, phosphorylation of FoxO3a Ser413/588 by AMPK has been found to increase FoxO3a transcriptional activity in vitro (11,32). Thus, AMPK-mediated phosphorylation of FoxO3a could also be involved in FoxO3a activation during exercise, although we did not test this possibility because mouse-specific antibodies are not currently available.

Autophagic activity characterization represents a challenge because of the dynamics of the system. Among Atgs proteins, LC3B-II has been positively correlated with an increased number of autophagosomes and is therefore commonly used as a marker of autophagy activation (2). Nonetheless, an increase in LC3B-II expression strengthens the hypothesis of an enhanced number of autophagosomes but does not allow making clear whether this raise is the consequence of an increased autophagosome formation or a defect in their degradation through the lysosome. p62, a ubiquitin-binding scaffold protein that binds with LC3, may serve to link ubiquitinated substrates to the autophagic machinery and is itself degraded by the lysosome. Thus, modulation of p62 levels may be used as a marker to study autophagic flux (3,20). In our study, we used several markers involved in different steps of the autophagic process, LC3B-II and p62 levels, and the phosphorylation level of Ulk1 at several sites. We found that exercise increases LC3B-II and induces a drop in p62 expression near to exhaustion, suggesting an increase in autophagic flux at high intensities. These elements are in agreement with others emergent studies (12,14,19) and strengthen the idea that autophagy–lysosomal pathway is induced by exercise. Ulk1 is a key initiator of autophagy, and Ser317/555 has been recently found to be phosphorylated by AMPK in vitro during glucose starvation and autophagy activation (18). MTOR phosphorylates Ulk1 at Ser757, resulting in loss of Ulk1 kinase activity, thus preventing the initiation of autophagy. Here we showed that endurance exercise leads to an increase in the phosphorylation state of Ser317 and Ser555, suggesting that AMPK plays a role in the early step of autophagosome formation during exercise. However, these two phosphorylation sites are differentially regulated. Indeed, Ser555 is quickly and strongly phosphorylated at 30 and 60 min and thereafter go back to its basal phosphorylation level, whereas Ser317 stays phosphorylated from 60 min to Te. Although the dynamics of Ulk1 complex is not clearly established in skeletal muscle, some allosteric modifications could explain such differences. These results imply that autophagy machinery would be quickly initiated during exercise, and not only for high intensities, suggesting that autophagy may constitute a generic response to any kind of endurance activity. Moreover, Ulk1 Ser757 phosphorylation was decreased from 120 min to Te concurrently to MTOR hypophosphorylation, supporting a potential role of the inhibition of the MTOR signaling pathway in autophagy induction by exercise.

Furthermore, the present study shows for the first time a rise in the protein expression level of the E3 ligase Mul1 in response to exercise. Nonetheless, we did not detect any variation in Mul1 mRNA level, suggesting that exercise induces a stabilization of Mul1 protein. The molecular mechanisms underlying this stabilization remain to be clarified. Mitochondrial remodeling through fission and fusion is essential for the removal of damaged mitochondria. Mul1 has been shown to ubiquitinate the mitochondrial profusion protein MFN2, causing its degradation through the proteasome. Nevertheless, we did not observe a decrease in MFN2, suggesting that its degradation rates were unaltered. Thus, the rise in Mul1 expression that we observed during exercise was probably not sufficient to induce MFN2 degradation. In the same way, OPA1 protein level, another GTPase required for mitochondrial fusion, was not significantly altered during exercise. Mul1 has also been reported to stabilize the mitochondrial fission protein DRP1 in vitro, resulting in mitochondrial fragmentation (4), but these events remain to be tested in skeletal muscle. Phosphorylation of DRP1 at Ser616 stimulates mitochondrial fission and such an increase has been reported to be permissive for mitophagy (37). Here, we found that DRP1 phosphorylation state was progressively increased until exhaustion. All together, these results indicate that endurance exercise quickly promotes DRP1 activation, potentially stimulating mitochondrial fission, but not Mul1-dependent degradation of mitochondrial fusion markers contrary to what it has been reported during muscle wasting (23). Although we did not establish whether Mul1 regulates mitophagy in this study, this certainly warrants further investigations especially with a view to characterize other Mul1 partners or targets than MFN2 and to investigate the possible differential effect of acute exercise between slow and fast skeletal muscles.

Concerning protein synthesis pathway, we found that Akt, MTOR, and 4E-BP1 phosphorylation was inhibited from 90 min to Te. While protein flux synthesis has not been performed, these results suggest that protein synthesis is inhibited during endurance exercise. The significant increase in eIF2α phosphorylation observed from 60 min of exercise highlights these hypotheses. Furthermore, the slow but significant increase in MTOR phosphorylation and the subsequent rise in 4E-BP1 phosphorylation found at 3 and 24 h after exercise suggest an increase in protein translation and possibly protein synthesis during the recovery period. Endurance training is well known to promote a fast-to-slow muscle phenotype shift and mitochondrial biogenesis but not to induce muscle growth. Thus, a rise in protein synthesis in response to endurance exercise would be necessary for tissue repair and remodeling, especially for the synthesis of specific subcellular protein not involved in muscle hypertrophy as mitochondrial proteins. In contrast, force and resistance training are two major stimuli of muscle protein synthesis resulting in hypertrophy.

The physiological relevance of these events can be speculated. In our model, AMPK and/or Akt inhibition would modulate FoxO3a activity and Ulk1 axis to mobilize protein degradation as source of alternative nutrient production and energy substrates when exercise intensity/duration becomes high. In addition, autophagy system is known to be required for normal cellular function and for response to multiple types of stress to maintain skeletal muscle function. Thus, during prolonged exercise, THE autophagy–lysosomal pathway may represent a compensatory mechanism to prevent cellular loss function caused by constraints linked to metabolic stress.

In conclusion, the present study aimed to identify specific signaling events related to cellular component turnover that are modulated during endurance exercise. To the best of our knowledge, this study is the first to give a picture of the regulation of pathways implicated in protein balance and which consider mitochondrial dynamics markers at different points of endurance exercise. Our data support the idea that Akt/MTOR pathway inhibition and AMPK activation modulate protein turnover through FoxO3a and Ulk1 axes. Noticeably, the mitochondrial Mul1 ubiquitin ligase is quickly induced by endurance exercise, but this induction is not sufficient to trigger mitochondrial fusion markers alteration. Further investigations are needed to better understand the overall implications of AMPK or other metabolic sensors in the regulation of mitophagy and to also identify the precise role of Mul1 in these events. On the basis of these data, it is clearly conceivable that the involvement of the autophagic system in response to exercise must be considered not only in muscle homeostasis but also in disease. Because exercise is associated with improved quality of life and constitutes one of the best approaches to limit atrophy and metabolic disorders, these research directions are crucial in the fight against a wide spectrum of metabolic and muscle diseases.

The authors would like to thank the rodent animal facility of the laboratory Muscle Dynamics and Metabolism (RAM, Campus La Gaillarde, INRA, Montpellier) and the METAMUS platform dedicated to the functional exploration metabolism. The authors thank J. Glaviole for helpful discussion.

This project was supported by the Faculty of Sport Sciences of the University of Montpellier 1 and the Institut National de la Recherche Agronomique (INRA). No conflict of interest, financial or otherwise, is declared by the authors.

The results of the present study do not constitute endorsement by American College of Sports Medicine.

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