Differential response of skeletal muscles to mTORC1 signaling during atrophy and hypertrophy
- C Florian Bentzinger1Email author,
- Shuo Lin1Email author,
- Klaas Romanino1,
- Perrine Castets1, 2,
- Maitea Guridi1,
- Serge Summermatter1,
- Christoph Handschin1,
- Lionel A Tintignac1, 3,
- Michael N Hall1 and
- Markus A Rüegg1Email author
© Bentzinger et al.; licensee BioMed Central Ltd. 2013
Received: 3 October 2012
Accepted: 15 February 2013
Published: 6 March 2013
Skeletal muscle mass is determined by the balance between protein synthesis and degradation. Mammalian target of rapamycin complex 1 (mTORC1) is a master regulator of protein translation and has been implicated in the control of muscle mass. Inactivation of mTORC1 by skeletal muscle-specific deletion of its obligatory component raptor results in smaller muscles and a lethal dystrophy. Moreover, raptor-deficient muscles are less oxidative through changes in the expression PGC-1α, a critical determinant of mitochondrial biogenesis. These results suggest that activation of mTORC1 might be beneficial to skeletal muscle by providing resistance to muscle atrophy and increasing oxidative function. Here, we tested this hypothesis by deletion of the mTORC1 inhibitor tuberous sclerosis complex (TSC) in muscle fibers.
Skeletal muscles of mice with an acute or a permanent deletion of raptor or TSC1 were examined using histological, biochemical and molecular biological methods. Response of the muscles to changes in mechanical load and nerve input was investigated by ablation of synergistic muscles or by denervation .
Genetic deletion or knockdown of raptor, causing inactivation of mTORC1, was sufficient to prevent muscle growth and enhance muscle atrophy. Conversely, short-term activation of mTORC1 by knockdown of TSC induced muscle fiber hypertrophy and atrophy-resistance upon denervation, in both fast tibialis anterior (TA) and slow soleus muscles. Surprisingly, however, sustained activation of mTORC1 by genetic deletion of Tsc1 caused muscle atrophy in all but soleus muscles. In contrast, oxidative capacity was increased in all muscles examined. Consistently, TSC1-deficient soleus muscle was atrophy-resistant whereas TA underwent normal atrophy upon denervation. Moreover, upon overloading, plantaris muscle did not display enhanced hypertrophy compared to controls. Biochemical analysis indicated that the atrophy response of muscles was based on the suppressed phosphorylation of PKB/Akt via feedback inhibition by mTORC1 and subsequent increased expression of the E3 ubiquitin ligases MuRF1 and atrogin-1/MAFbx. In contrast, expression of both E3 ligases was not increased in soleus muscle suggesting the presence of compensatory mechanisms in this muscle.
Our study shows that the mTORC1- and the PKB/Akt-FoxO pathways are tightly interconnected and differentially regulated depending on the muscle type. These results indicate that long-term activation of the mTORC1 signaling axis is not a therapeutic option to promote muscle growth because of its strong feedback induction of the E3 ubiquitin ligases involved in protein degradation.
KeywordsSkeletal muscle Hypertrophy Atrophy Mammalian target of rapamycin complex 1 (mTORC1) Raptor Tuberous sclerosis complex (TSC) PKB/Akt FoxO MuRF1 Atrogin-1/MAFbx
Bovine serum albumin
Forkhead box O
Hematoxylin & eosin
Hypertrophy on recovery
Human skeletal actin
Insulin-like growth factor-1
Mammalian target of rapamycin complex 1
Nicotinamide adenine dinucleotide hydrogen-tetrazolium reductase
Protein kinase B (also called Akt)
Peroxisome proliferator-activated receptor gamma coactivator 1
Regulatory-associated protein of mTOR
Raptor muscle knockout
Ras homolog enriched in brain
Rapamycin-insensitive companion of mTOR
Surface sensing of translation
Tris-buffered saline with 0.1% Tween 20
Tuberous sclerosis complex
TSC muscle knockout
Skeletal muscle is the largest organ, accounting for 30 to 40% of the total body weight. Muscle tissue is highly plastic and adapts its size to physical demand. For example, increase in load causes hypertrophy whereas unloading causes atrophy. Importantly, muscle atrophy and subsequent wasting are also hallmarks of pathology in muscular dystrophies or in cachexia, the latter being a secondary consequence of a primary disease (for example, AIDS, cancer or sepsis). Several lines of evidence indicate that muscle mass is controlled by the balance between protein synthesis and protein degradation [1, 2]. In skeletal muscle, protein synthesis can be induced by IGF1 (insulin-like growth factor-1), which in turn, activates PI3K (phosphatidylinositol 3-kinase) and PKB (protein kinase B; also called Akt). Activated PKB/Akt inhibits the protein complex TSC1-TSC2 (tuberous sclerosis complex), which inactivates the small GTPase protein Rheb (Ras homolog enriched in brain). Rheb activates mammalian target of rapamycin complex 1 (mTORC1), which causes an increase in protein translation by phosphorylating its two best characterized targets S6K (p70S6 kinase) and 4EBP (eIF-4E-binding protein). This IGF1-PI3K-PKB/Akt-mTOR signaling pathway controls protein synthesis and cell size in several tissues [3, 4].
Activation of PKB/Akt also negatively regulates protein degradation by phosphorylating the FoxO (Forkhead box O) transcription factors. Protein degradation is mainly carried out by enzymes of the ubiquitin-proteasomal and autophagosomal-lysosomal pathways [5, 6]. Dephosphorylated FoxOs in the nuclei promote the expression of the two E3 ubiquitin ligases atrogin-1/MAFbx and MuRF1[7, 8]. FoxOs have also been described to drive expression of autophagy-related genes [6, 9]. The function of active PKB/Akt to simultaneously stimulate protein synthesis and inhibit protein degradation may explain the profound hypertrophic effect of constitutively active PKB/Akt [10, 11].
mTOR belongs to the PI3/PI4-kinase family; it is highly conserved from yeast to human and assembles into two structurally and functionally distinct multiprotein complexes, called mTORC1 and mTORC2 [12, 13]. An essential component of mTORC1 is the protein raptor (regulatory-associated protein of mTOR), whereas rictor (rapamycin-insensitive companion of mTOR) is an essential subunit of mTORC2 [3, 4]. Most functions of mTORC1 are acutely inhibited by the immunosuppressant rapamycin, whereas mTORC2 is only repressed by long-term application of rapamycin . In skeletal muscle, the function of mTORC2 seems to not be essential because mice deficient for rictor have no overt phenotype [15, 16]. In contrast, mTORC1 participates in the control of muscle size. For example, rapamycin prevents IGF1-induced growth of myotubes , inhibits compensatory hypertrophy in rat skeletal muscle  and blocks the growth-stimulating activity of clenbuterol . Moreover, transgenic overexpression of TSC1 causes muscle atrophy in mice , while acute overexpression of Rheb induces muscle hypertrophy . Finally, mice deficient for S6K1 show a reduction of muscle fiber size and a blunted response to IGF1 .
In agreement with these findings, we recently showed that mice with a skeletal muscle-specific knockout for raptor (called RAmKO for raptor muscle knockout) have a reduced muscle mass and suffer from a progressive dystrophy, which causes their death at the age of four to six months . Muscles of RAmKO mice also have a decreased oxidative capacity, which can be restored by transgenic expression of PGC-1α . In addition, RAmKO mice show sustained activation of PKB/Akt because of relieved feedback inhibition onto IRS1 (insulin receptor substrate-1) by the diminished activation of S6K .
Here we investigated the contribution of mTORC1 to muscle atrophy and hypertrophy by targeting rptor (the gene encoding raptor) or Tsc1 (encoding TSC1) specifically in mouse skeletal muscle. We show that deletion of rptor prevents muscle hypertrophy and enhances muscle atrophy. Surprisingly, sustained activation of mTORC1 by the genetic deletion of Tsc1 does not induce hypertrophy but rather causes atrophy in all but soleus muscles. While the TSC1-deficient, hypertrophic soleus muscle is also resistant to denervation-induced atrophy, tibialis anterior (TA) muscle atrophies like controls. Biochemical characterization shows that regulation of the two E3 ligases atrogin-1/MAFbx and MuRF1 differs between TA and soleus muscles. Furthermore, we demonstrate that all muscles show an increase in their oxidative capacity upon mTORC1 activation. In summary, we demonstrate that the oxidative capacity in all skeletal muscles is controlled by mTORC1, whereas the effect of sustained activation of mTORC1 on muscle size differs between muscles. Hence, our studies decipher a mechanism of biological robustness that balances the two major metabolic pathways involved in the control of skeletal muscle mass.
Mice were maintained in a conventional facility with a fixed dark–light cycle. Studies were carried out according to criteria outlined for the care and use of laboratory animals and with approval of the Swiss authorities. RAmKO mice were generated and genotyped as described before . Floxed Tsc1 mice  were obtained from The Jackson Laboratory (Bar Harbor, Maine, USA) and mated with mice expressing Cre recombinase under the human skeletal actin (HSA) promoter . Genotyping for the conditional Tsc1 allele was performed as described . TSC-RAmKO mice were generated by intercrossing mice carrying floxed rptor and Tsc1 alleles. Mice homozygous for both floxed alleles were mated with double heterozygotes, which also carried the HSA-Cre transgene. Except for overloading experiments and Western blot analysis, only male TSCmKO mice were used. Both genders were used in RAmKO and TSC-RAmKO mice. All procedures were performed in accordance with the Swiss regulations for animal experimentation and they were approved by the veterinary commission of the Canton Basel-Stadt.
Rapamycin treatment of mice
Rapamycin treatment began three days before the mice were challenged with functional overload (FO) or electroporation and continued until mice were sacrificed. Rapamycin (LC Laboratories, Woburn, MA, USA), dissolved in saline containing 2% carboxymethylcellulose (Sigma-Aldrich, St. Louis, MO, USA), was delivered once daily by i.p. (intraperitoneal) injection at a dose of 1.5 mg/kg .
The methods to construct plasmids encoding shRNA and the sequences of the Cd4 control shRNA and the NLS-GFP construct have been described elsewhere . The murine 19 nucleotide target sequences correspond to: GTT GAT GCG TAA CCT TCT G (Tsc2), GAT GGA CAC TGA TGT TGT G (Tsc1) and GAA TTT TGC TGA TTT GGA A (rptor).
Tissue culture, transfections and shRNA efficiency
Adenoviruses encoding shRNA against Tsc2 and Cd4 were created by cloning the respective shRNA sequence and H1 promoter from pSuper into pAd-DEST (Life Technologies Europe B.V., Zug, Switzerland). To test the efficiency of the Tsc2 shRNA, C2C12 myoblasts, cultured under standard conditions, were transfected with the Tsc2 and Cd4 shRNA viruses. The efficiency of the rptor shRNA was tested by co-transfection with an expression plasmid encoding HA-tagged raptor into COS7 cells using Lipofectamine 2000 (Life Technologies). For PGC1β overexpression and knockdown, myoblasts were permitted to fuse into multinucleated myotubes for 48 hr and cells were infected with adenovirus preparations for an additional 48 hr. Adenoviruses (Ad-GFP, Ad-PGC1β, Ad-scrambled or Ad-siPGC1β) were kindly provided by BM Spiegelman (Harvard University, Boston, MA, USA).
Electroporation of muscle
Plasmids encoding shRNA constructs were electroporated into muscle fibers as described before . Briefly, soleus or TA muscle of anesthetized mice was exposed and injected with 10 to 30 μl of a mixture containing the respective shRNA plasmid and a plasmid coding for NLS-GFP (2 mg/ml of each construct). The fascia and the skin were sutured and the electroporation was performed using an ECM 830 electroporation system (BTX Instruments Division, Harvard Apparatus Inc., Holliston, MA, USA). Eight pulses lasting 20 ms with the frequency of 1 Hz and the voltage set to 180 V/cm were applied. Mice were analyzed four to six weeks after electroporation.
Denervation, nerve crush and overloading
Mice were anesthetized with ketamine (111 mg/kg) and xylazine (22 mg/kg) by intra-peritoneal injection and surgery was performed under aseptic conditions. For denervation, a segment (approximately 5 mm) of the sciatic nerve at the mid-thigh level was excised . To induce muscle re-growth, the nerve was crushed with No 5 Dupont forceps (Fine Science Tools GmbH, Heidelberg, Germany) for 10 seconds at mid-thigh . To induce muscle hypertrophy, a functional overload of plantaris muscle was introduced by surgical removal of soleus and gastrocnemius muscles . Surgery was performed on one leg only. The plantaris muscle of the contralateral leg served as control.
The antibodies used were from the following sources: rabbit polyclonal antibodies directed to 4E-BP1 (Phas-I) from Zymed (Life Technologies); those recognizing Phospho-4E-BP1 (Ser65), PKB/Akt, mTOR, S6 Ribosomal Protein or Phospho-S6 Ribosomal Protein (Ser235/236) were all from Cell Signaling Technology Inc. (Danvers, MA, USA); those against FoxO1a were from Abcam plc. (Cambridge, UK); those against TSC1 were from Bethyl Laboratories (Montgomery, TX, USA). Rabbit monoclonal antibodies directed against Phospho-Akt (Ser473), IRS-1, FoxO3a (75D8) and phospho-FoxO1(Thr24)/FoxO3a(Thr32) (#9466) were from Cell Signaling Technology Inc. Mouse monoclonal antibodies to α-actinin were purchased from Sigma and antibodies against HA from Covance Inc. (Geneva, Switzerland). Rat monoclonal antibodies directed to the Laminin B2 Chain (MAB1914) were from Chemicon and sold by Millipore AG (Zug, Switzerland). The TSC2 antibodies used were described elsewhere . Mouse monoclonal antibodies against myosin heavy chain: slow (A4.840), IIa/IIx (A4.74) and IIb (BF-F3) were purchased from The Developmental Studies Hybridoma Bank (University of Iowa, Iowa City, Iowa, USA). Antibodies to puromycin  were a kind gift of Dr. Philippe Pierre (CIML Parc Scientifique de Luminy, Marseille, France).
Histology and immunohistochemistry
Muscles frozen in liquid nitrogen-cooled isopentane were cut into 12 μm cross-sections. Cross-sections were fixed with 2% paraformaldehyde (PFA) and permeabilized with 1% Triton/PBS for 5 minutes, washed with 100 mM glycine/PBS for 15 minutes, blocked with 1% BSA/PBS for 30 minutes, and incubated with the primary antibody overnight at 4°C. Samples were subsequently washed three times for 10 minutes each with 1% BSA/PBS and stained with the appropriate fluorescence labeled secondary antibodies for 1 hr at room temperature. After washing with PBS, samples were mounted with Citifluor (Citifluor Ltd. London, UK). General histology on cross-sections was performed using hematoxylin and eosin (H&E; Merck, Zug, Switzerland). NADH-TR (Nicotinamide adenine dinucleotide hydrogen-tetrazolium reductase) staining was done as described . Methods of SDH and COX staining were described elsewhere . Samples were dehydrated and mounted with DePeX mounting medium (Gurr, BDH, VWR International GmbH, Dietikon, Switzerland).
In vivo protein synthesis
Protein synthesis was measured using the surface sensing of translation (SUnSET) method  by i.p. injection of 0.040 μmol/g puromycin dissolved in 100 μl of PBS. Mice were sacrificed 30 minutes later and muscles were snap-frozen in liquid nitrogen. Muscles were lysed as described below and proteins were separated on 8 to 16% SDS-PAGE (Bio-Rad Laboratories AG, Cressier, Switzerland). After transfer to polyvinyl difluoride membranes and blocking of free binding sites with 5% milk powder in Tris-buffered saline with 0.1% Tween 20 (TBST), the mouse IgG2a monoclonal anti-puromycin antibody (clone 12D10; 1:5,000) was incubated for 1 hr at room temperature. After incubation with the appropriate HRP-coupled secondary antibody, blots were developed using enhanced chemiluminescence reagent. Coomassie Blue staining was used to verify equal loading.
Tissue homogenization, SDS-PAGE and Western blot
Muscles frozen in liquid nitrogen were powdered on dry ice and lysed in cold RIPA buffer supplemented with 1% Triton-X, 10% glycerol, protease inhibitor cocktail tablets (Roche Diagnostics AG, Rotkreuz, Switzerland), and phosphatase inhibitor cocktail I and II (Sigma). Cell lysates were incubated on ice for 2 hr, sonicated two times for 15 s and centrifuged at 13,600 g for 30 minutes at 4°C. Cleared lysates were then used to determine total protein levels (BCA Protein Assay, Pierce, Rockford, IL, USA). After dilution with sample buffer, equal protein amounts were loaded onto SDS gels.
Total RNA was isolated (SV Total RNA Isolation System, Promega AG, Dübendorf, Switzerland) from soleus muscles. RNA concentrations were adjusted between samples and reverse transcription was carried out using a mixture of oligodT and random hexamer primers (iScript cDNA Synthesis Kit, Bio-Rad Laboratories AG). Sybr Green, real-time PCR analysis (Power SYBR Green Master Mix, Life Technologies) was performed using the ABI Prism 7000 Sequence Detector. Expression levels for each gene of interest were normalized to the expression of the housekeeping protein β-actin. The following primers were used: β-actin sense primer: 5' CAG CTT CTT TGC AGC TCC TT, antisense primer: 5' GCA GCG ATA TCG TCA TCC A; atrogin-1/MAFbx sense primer: 5' CTC TGT ACC ATG CCG TTC CT, antisense primer: 5' GGC TGC TGA ACA GAT TCT CC; MuRF-1 sense primer: 5′ ACC TGC TGG TGG AAA ACA TC, antisense primer: 5′ AGG AGC AAG TAG GCA CCT CA; Pgc1α sense primer: 5’ TGA TGT GAA TGA CTT GGA TAC AGA CA, antisense primer: 5’ GCT CAT TGT TGT ACT GGT TGG ATA TG; Pgc1β sense primer: 5' GGC AGG TTC AAC CCC GA, antisense primer: 5' CTT GCT AAC ATC ACA GAG GAT ATC TTG. Quantification of mitochondrial DNA copy numbers was done as described .
Quantifications and statistics
For muscle fiber size quantification, muscle cross-sections were stained either for laminin-γ1 or fluorescence labeled wheat-germ agglutinin. Images were acquired using a Leica DM5000B fluorescence microscope with 10x objective, a digital camera (F-View; Soft Imaging System, Olympus Soft Imaging Solutions GmbH, Münster, Germany), and analySIS software (Soft Imaging System). Images of the entire soleus or tibialis anterior (TA) muscles were aligned with Adobe PhotoShop (Adobe Systems Incorporated, San Jose, CA, USA). The minimum distance of parallel tangents at opposing particle borders (minimal feret’s diameter) and cross-section area (CSA) were measured with analySIS software as described . Data are expressed as mean ± SEM. For statistical comparison of two conditions, the Student’s t- test was used. The level of significance is indicated as follows: *** P <0.001, ** P <0.01, * P <0.05.
Acute changes in mTORC1 activity affect muscle fiber size
To test the role of mTORC1 in muscle plasticity, we crushed the sciatic nerve unilaterally immediately after electroporation, which causes a transient denervation-induced atrophy, followed by fiber re-innervation and re-growth to normal size [27, 33, 34]. Such “hypertrophy on recovery” (HOR) was significantly less in muscle fibers expressing shRNA to rptor and significantly higher in fibers expressing shRNA to Tsc2 (Figure 1C, D). To test whether shRNA-targeting acted on the initial atrophy or on re-growth, we also examined electroporated muscle fibers in a pure denervation-induced atrophy paradigm. No difference between non-electroporated and electroporated fibers was detected in Cd4 controls (Figure 1E, F). In contrast, muscle fibers expressing shRNA to Tsc2 were much bigger than non-electroporated fibers and, like in innervated muscle, the effect of TSC2 knockdown was abrogated by rapamycin (Figure 1E, F). Similar results were obtained by electroporating tibialis anterior (TA) muscle (Additional file 1, Figure S1E, F). These results thus show that acute alteration of mTORC1 activity affects the response of both, the slow oxidative soleus and fast glycolytic TA muscles to growth-stimulating and atrophy-inducing conditions.
Constitutive deletion of Tsc1 in skeletal muscle fibers affects muscles differentially
Quantification of Western blot analysis
Number of replicates
19 ± 2***
10 ± 1
31 ± 6
28 ± 5
23 ± 5***
12 ± 3
34 ± 6
39 ± 6
53 ± 1***
13 ± 6
48 ± 11
42 ± 17
6 ± 2***
24 ± 1
P-PKB/ Akt S473
4 ± 2***
22 ± 3
28 ± 7
22 ± 6
23 ± 5
18 ± 3
16 ± 7
17 ± 5
P-FoxO1 T24 /3a T23
2 ± 7**/ 8 ± 3*
11 ± 2/ 20 ± 5
P-PKB/ AktS 473 (Den. TA)
P-S6 S235/S236 (Den. TA)
47 ± 12***
5 ± 2
P-PKB/ Akt S473 (Den. Sol)
P-S6 S235/S236 (Den. Sol)
41 ± 10**
3 ± 5
number of replicates
P-PKB/ Akt S473 (Den. TA)
12 ± 2
2 ± 1
P-S6 S235/S236 (Den. TA)
P-PKB/ Akt S473 (Den. Sol)
19 ± 3
3 ± 2
P-S6 S235/S236 (Den. Sol)
Consistent with the activation of the mTORC1 targets and the role of mTORC1 in the control of protein translation, protein synthesis in EDL muscle of TSCmKO was increased (Additional file 1: Figure S2A). However, TSCmKO mice gained less weight than their control littermates. Starting from the age of five weeks, male TSCmKO mice were significantly lighter (Additional file 1: Figure S2B), whereas the weight difference in females did not reach significance (Additional file 1: Figure S2C). At least part of this weight difference was due to alteration in muscle mass as all but soleus muscles were significantly lighter than in control mice (Figure 2B). Thus, despite increased protein synthesis, all but soleus muscles are lighter in TSCmKO mice than in control mice.
To investigate the reason for these muscle-specific differences in weight, we focused on soleus and TA muscles in three-month-old mice. Hematoxylin & eosin (H&E) staining did not reveal any major alterations in either of the muscles (Figure 2C). The difference in the muscle weight was matched by changes in the muscle fiber size in soleus and TA muscle (Figure 2D, E). Detailed analysis of fiber types showed that both type I and type IIa fibers were larger in soleus muscle (Additional file 1: Figure S3; Additional file 1: Table S1). In TA muscle, the glycolytic type IIb fibers were significantly smaller whereas the oxidative type IIa/x fibers were not affected (Additional file 1: Figure S3, Additional file 1: Table S1). In summary, these data show that the response to the activation of mTORC1 differs between muscles and fiber types.
Sustained activation of mTORC1 increases the oxidative capacity in all muscles
Additional factors that are regulated by mTORC1 [16, 22, 38] and have been implicated in the control of muscle size are the transcriptional coactivators PGC1α and PGC1β [39, 40]. Moreover, PGC1α and PGC1β are major regulators of mitochondrial biogenesis . To test whether deletion of Tsc1 would also affect the PGC1 pathway and the oxidative capacity of skeletal muscle, we next compared expression of Pgc1α and Pgc1β in TA and soleus muscles of TSCmKO mice with littermate controls. Contrary to the expectation, transcript levels of Pgc1α were decreased in mutant muscles compared to controls (Figure 3D, E). The down-regulation of Pgc1α was more pronounced in soleus muscle, which expresses the highest level of PGC1α in wild-type mice . In contrast, mRNA levels of Pgc1β were increased to about 150% in all examined muscles of TSCmKO mice (Figure 3D, E). In support of a direct regulation of Pgc1β transcripts by mTORC1, Pgc1β expression was diminished in RAmKO mice (soleus muscle in RAmKO mice: 73 ± 4.6%; control mice: 100 ± 10.3%; mean ± SEM; N ≥5; P <0.05). Hence, unlike expression of the E3 ubiquitin ligases atrogin-1/MAFbx and MuRF1, expression of Pgc1α and Pgc1β did not differ between TA and soleus muscles in TSCmKO mice. Overexpression and knockdown experiments of PGC1β in C2C12 myotubes indicate that expression of Pgc1α is tightly regulated by PGC1β (Figure 3F). Such counter-regulation between PGC1α and PGC1β has also been reported in other tissues . Thus, the increased levels of Pgc1β transcripts in the TSCmKO mice likely suppress expression of Pgc1α. Interestingly, TSCmKO mice showed an increase in their capacity for oxidative phosphorylation in TA and soleus muscles as shown by stainings for NADH-TR (Figure 3G), succinate dehydrogenase (SDH; Additional file 1: Figure S4A, B) and cytochrome oxidase (COX; Additional file 1: Figure S4A, B). This increase was accompanied by a slight, although not significant, increase in the number of mitochondria as determined by qPCR of mitochondrial DNA (Additional file 1: Figure S4C). Taken together, these data suggest that PGC1β is responsible for the increased oxidative properties of skeletal muscle of TSCmKO mice.
mTORC1 is required for muscle fiber hypertrophy
There is evidence that FO also causes some damage and muscle regeneration and that satellite and other cells outside the muscle’s basal lamina contribute to the weight increase [47, 48]. As HSA-Cre is not expressed in non-muscle cells and satellite cells , we treated control mice with rapamycin during FO to eliminate mTORC1 function in all cells. This treatment abolished both the increase in weight and the shift in fiber size distribution (Additional file 1: Figure S5C), suggesting that mTORC1 expressed in non-muscle cells or in satellite cells might contribute to the increased weight of plantaris muscles in RAmKO mice after FO.
As FO induces a relative increase in the number of oxidative fibers , we also stained the overloaded plantaris from control and mutant mice by NADH-TR. As shown in Figure 4H, plantaris muscles remained largely non-oxidative in RAmKO mice, whereas in the overloaded plantaris of TSCmKO mice even the large myofibers remained highly oxidative (Figure 4I).
Soleus and TA muscles of TSCmKO mice respond differently to denervation-induced atrophy
Feedback control of PKB/Akt is active during muscle atrophy
The expression of the mTORC1 target PGC1α is also controlled by denervation . In innervated soleus muscle of RAmKO mice, Pgc1α mRNA levels are less than 40%  and Pgc1β mRNA levels are approximately 70% of control muscle. In denervated TA and soleus muscles of control mice, expression of Pgc1α and Pgc1β was lower than in innervated muscle (Figure 6G, H). Similarly, denervation lowered the levels of both transcriptional co-activators in RAmKO mice although the significant difference to control mice was lost (Figure 6G, H). In contrast, expression of Pgc1α and Pgc1β was very different in TSCmKO mice. While Pgc1α mRNA levels were decreased upon denervation both in TA and soleus muscles, Pgc1β was significantly increased in both muscles (Figure 6I, J). Taken together, our results show that atrophy is accelerated in RAmKO mice despite low levels of atrogin-1/MAFbx and MuRF1. Conversely, the sparing of soleus muscles from denervation-induced atrophy in TSCmKO mice could be based on the low levels of the two E3 ubiquitin ligases in this particular muscle. In contrast, the relative levels of Pgc1α and Pgc1β did not differ between TA and soleus muscles upon denervation and are thus unlikely contributors to the differential response.
Genetic inactivation of mTORC1 reverses the phenotype of TSCmKO mice
Here we describe the phenotype of mice in which mTORC1 is constitutively active in skeletal muscle (TSCmKO) and compare it to mice with inactivated mTORC1 signaling (RAmKO). While the oxidative changes in TSCmKO mice were largely the opposite of those observed in RAmKO mice and affected all examined muscles, the effect of mTORC1 activation on muscle size was unexpected as all muscles except soleus muscles were slightly but significantly smaller. Thus, our work highlights the existence of several feed-forward or auto-inhibitory loops that allow fine-tuning of the signaling networks involved in the control of muscle mass (Figure 7E).
Based on the current concepts, mTORC1 activation should result in an increase in muscle mass and muscle fiber size. This view is based on the findings that activation of the mTORC1 upstream components PKB/Akt or IGF-1 receptor causes an increase in muscle mass [5, 10, 11, 52, 53] and that this increase is rapamycin-sensitive [11, 53]. Moreover, overexpression of Rheb in single muscle fibers by electroporation leads to hypertrophy of the transfected fibers  and whole body knockout of the mTORC1 target S6K1 results in smaller muscle fibers . Consistent with these experiments, acute knockdown of TSC1/2 by shRNA resulted in slightly bigger muscle fibers in soleus or TA muscles, confirming that transient activation of the mTORC1 pathway is sufficient to induce muscle fiber growth. However, under conditions of prolonged activation of mTORC1 in TSCmKO mice, all muscles examined, with the exception of soleus, were smaller than in control mice. As mTORC1 targets are activated and protein synthesis in EDL muscle of TSCmKO mice is increased, the atrophy induced by chronic mTORC1 activation is likely related to the feedback inhibition of activated S6K onto IRS1, which in turn, decreases activation of PKB/Akt. This tight feedback control of S6K on IRS1-PKB/Akt was also observed in mice deficient for raptor or mTOR in some tissues including skeletal and heart muscle [16, 54–56] but not in others . Similarly, deletion of TSC1 strongly decreases activation of PKB/Akt in cultured mouse embryonic fibroblasts , whereas it does not at all affect PKB/Akt phosphorylation in some tissues [58, 59]. These data indicate that the feedback control of S6K depends on the cellular context and our data now show that this feedback is particularly strong in skeletal muscle.
Consistent with decreased inhibition of FoxO transcription factors by PKB/Akt, TA muscle from TSCmKO mice express high levels of MuRF1 and atrogin-1/MAFbx, involved in protein degradation through the proteasome [7, 8]. Hence, the atrophy observed in muscles of the TSCmKO mice is likely caused by the prevalence of the FoxO pathway over mTORC1 activation. This differs from the muscle hypertrophy observed using the transient, partial activation of mTORC1 with shRNA electroporation. Thus, the atrophy response caused by the sustained, saturated mTORC1 activation by genetic Tsc1 deletion may unveil a long-term adaptation of the FoxO pathway. Consistently, transient overexpression of Rheb does not seem to affect PKB/Akt phosphorylation , further supporting the idea that muscle atrophy in TSCmKO mice is related to the indirect PKB/Akt-dependent activation of FoxO pathways.
Importantly, contrasting with the atrophic phenotype of most muscles, sustained activation of mTORC1 leads to increased mass of soleus muscle in TSCmKO mice. Although PKB/Akt was similarly inhibited in soleus and TA muscles, expression of MuRF1 and atrogin-1/MAFbx was not increased in soleus muscle, indicating that an additional regulatory mechanism suppresses their expression, thereby overruling the regulation by PKB/Akt. This differential regulation of MuRF1 and atrogin-1/MAFbx expression did not seem to be mediated by PGC1α, previously identified as a negative regulator of FoxO , because there was no significant difference in PGC1α/β expression between soleus and TA muscles from TSCmKO mice.
With different atrophy and hypertrophy paradigms, we also demonstrate that mTORC1 plays a critical and complex role in muscle plasticity. Using shRNA electroporation, we show that transient activation of mTORC1 is sufficient to limit denervation-induced atrophy and to enhance fiber hypertrophy upon re-innervation. Similarly, TSCmKO mice display atrophy resistance to denervation in soleus muscle, which shows only moderate expression of the E3 ubiquitin ligases MuRF1 and atrogin-1/MAFbx. By contrast, long-term activation of mTORC1 did not protect TA muscle from atrophy and did not exacerbate the hypertrophy response to overloading of plantaris muscle. These results indicate that the increased protein synthesis by mTORC1 hyperactivation is not sufficient to maintain muscle mass in cases where the FoxO-MuRF1-atrogin-1/MAFbx axis is active due to the absence of PKB/Akt signaling. Importantly, both transient and long-term inactivation of mTORC1 increased denervation-induced atrophy and prevented muscle growth associated with re-innervation or overloading, indicating that increased protein synthesis is required even when the catabolic proteasomal activity is reduced. Thus, our results provide genetic evidence that muscle growth requires mTORC1.
In our previous work, we demonstrated that raptor-deficient skeletal muscles show a strongly decreased oxidative capacity due to changes in mitochondrial function . This loss of oxidative capacity correlated with a substantial decrease in the transcript levels of Pgc1α, consistent with the direct regulation of Pgc1α expression by mTOR , and could be restored by transgenic expression of PGC1α . Contrary to the expectations and the effect of mTORC1 activation in embryonic fibroblasts , all examined muscles of TSCmKO mice showed a decreased expression of Pgc1α but increased levels of Pgc1β. Thus, the increase in the oxidative capacity in TSCmKO mice may be mediated by PGC1β. Indeed, PGC1β has also been shown to be sufficient to increase oxidative capacity in skeletal muscle despite the concomitant reduction in PGC1α expression . Moreover, depletion of both PGC1α and PGC1β results in much more severe loss of oxidative capacity than depletion of either protein alone . The reason for the unexpected down-regulation of Pgc1α transcripts in TSCmKO mice might be the counter-regulation of PGC1α and PGC1β. We show here that overexpression of PGC1β in C2C12 myotubes results in a strong suppression of the endogenous Pgc1α expression and, conversely, Pgc1β knockdown leads to increased expression of Pgc1α transcripts. These data indicate that the total amount of both PGC1 co-activators is tightly controlled in skeletal muscle.
Our study provides new functional insights into the molecular mechanism of muscle atrophy and hypertrophy. The data demonstrate that mTORC1 modulation downstream of PKB/Akt is subject to biological robustness. A fine-tuned feedback loop controlled by the anabolic mTORC1 pathway mediates crosstalk to E3 ubiquitin ligase system that increases protein degradation and thus compensates for imbalance. However, this feedback system fails to fully re-establish muscle homeostasis, leading to prevalence of either an anabolic or a catabolic net response. Our observations emphasize that muscle growth requires both activated PKB/Akt and mTORC1 in parallel, and they provide a new rationale for the development of pharmacologic agents that target this system.
We thank Drs. Xian Chu Kong and Céline Costa for their help with the shRNA constructs. We thank Dr. Philippe Pierre (CIML Parc Scientifique de Luminy, Marseille, France) for providing us with the anti-puromycin antibody. This work was supported by the Cantons of Basel-Stadt and Baselland, grants from the Swiss National Science Foundation, the Swiss Foundation for Research on Muscle Disease, Swiss Life and the Association Française contres les Myopathies (AFM).
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