Myostatin reduces Akt/TORC1/p70S6K signaling, inhibiting myoblast ...

Am J Physiol Cell Physiol 296: C1258 –C1270, 2009. First published April 8, 2009; doi:10.1152/ajpcell.00105.2009.

Myostatin reduces Akt/TORC1/p70S6K signaling, inhibiting myoblast differentiation and myotube size Anne Ulrike Trendelenburg,1 Angelika Meyer,1 Daisy Rohner,1 Joseph Boyle,1 Shinji Hatakeyama,1 and David J. Glass2 1

Novartis Institutes for Biomedical Research, Forum 1, Novartis Campus, Basel, Switzerland; and 2Novartis Institutes for Biomedical Research, Cambridge, Massachusetts

Submitted 6 March 2009; accepted in final form 23 March 2009

Trendelenburg AU, Meyer A, Rohner D, Boyle J, Hatakeyama S, Glass DJ. Myostatin reduces Akt/TORC1/p70S6K signaling, inhibiting myoblast differentiation and myotube size. Am J Physiol Cell Physiol 296: C1258 –C1270, 2009. First published April 8, 2009; doi:10.1152/ajpcell.00105.2009.—Myostatin is a negative regulator of skeletal muscle size, previously shown to inhibit muscle cell differentiation. Myostatin requires both Smad2 and Smad3 downstream of the activin receptor II (ActRII)/activin receptor-like kinase (ALK) receptor complex. Other transforming growth factor-␤ (TGF␤)-like molecules can also block differentiation, including TGF-␤1, growth differentiation factor 11 (GDF-11), activins, bone morphogenetic protein 2 (BMP-2) and BMP-7. Myostatin inhibits activation of the Akt/mammalian target of rapamycin (mTOR)/p70S6 protein synthesis pathway, which mediates both differentiation in myoblasts and hypertrophy in myotubes. Blockade of the Akt/mTOR pathway, using small interfering RNA to regulatory-associated protein of mTOR (RAPTOR), a component of TOR signaling complex 1 (TORC1), increases myostatin-induced phosphorylation of Smad2, establishing a myostatin signaling-amplification role for blockade of Akt. Blockade of RAPTOR also facilitates myostatin’s inhibition of muscle differentiation. Inhibition of TORC2, via rapamycin-insensitive companion of mTOR (RICTOR), is sufficient to inhibit differentiation on its own. Furthermore, myostatin decreases the diameter of postdifferentiated myotubes. However, rather than causing upregulation of the E3 ubiquitin ligases muscle RING-finger 1 (MuRF1) and muscle atrophy F-box (MAFbx), previously shown to mediate skeletal muscle atrophy, myostatin decreases expression of these atrophy markers in differentiated myotubes, as well as other genes normally upregulated during differentiation. These findings demonstrate that myostatin signaling acts by blocking genes induced during differentiation, even in a myotube, as opposed to activating the distinct “atrophy program.” In vivo, inhibition of myostatin increases muscle creatine kinase activity, coincident with an increase in muscle size, demonstrating that this in vitro differentiation measure is also upregulated in vivo. human skeletal muscle cells; transforming growth factor-␤-like molecules; Smad signaling; mammalian target of rapamycin complex signaling; transducer of regulated Ca2⫹-responsive element-binding protein activity; S6 kinase; MuRF1; MAFbx; atrogin; MuRF-1 SKELETAL MUSCLE ATROPHY—a

loss of muscle mass and fiber size—is a hallmark of cachexia, a severe comorbidity seen in congestive heart failure, chronic obstructive pulmonary disease, closed head trauma, and severe burns (12). Recently, two markers of skeletal muscle atrophy have been characterized, the muscle-specific E3 ubiquitin ligases muscle RING-finger 1 (MuRF1) and muscle atrophy F-box (MAFbx; also called atrogin-1). MuRF1 and MAFbx have been demonstrated to be

Address for reprint requests and other correspondence: D. J. Glass, Novartis Institutes for Biomedical Research, 100 Technology Square, Rm. 4210, Cambridge, MA 02139 (e-mail: [email protected]). C1258

upregulated in at least 13 distinct models of skeletal muscle atrophy (3, 6 – 8, 11, 13, 17, 19, 22, 24, 28, 40, 41, 47, 50), in both rodents and humans (34, 42). For example, it was recently demonstrated that in the acute atrophy that occurs in diaphragm muscle, in patients on mechanical ventilation, MAFbx expression increased more than twofold, and MuRF1 was increased by greater than fivefold (27). The expression of these E3 ligases MuRF1 and MAFbx can be opposed by activating the insulin-like growth factor 1 (IGF-1)/Akt pathway (24). Akt phosphorylates and thus inhibits the family of FoxO transcription factors, which are required for MuRF1 and MAFbx upregulation (43, 48). The IGF-1/Akt pathway also induces skeletal muscle hypertrophy by increasing protein synthesis pathways (21, 33, 38). IGF-1 modulation of protein synthesis operates via Akt/mammalian target of rapamycin (mTOR) signaling. mTOR can be activated both by growth factor signaling, via Akt, and by the presence of branched chain amino acids. There are two distinct mTOR complexes, called TORC1 and TORC2 (2). TORC1 includes a protein called RAPTOR (regulatory-associated protein of mTOR); this complex can be inhibited by rapamycin (45). In a second, rapamycinindependent, complex, TORC2, mTOR is complexed with a protein called RICTOR (rapamycin-insensitive companion of mTOR), which has been characterized as being able to phosphorylate Akt on serine 473, in a positive feedback mechanism (44). The roles of RICTOR and RAPTOR in myostatin or transforming growth factor-␤ (TGF-␤) inhibition of muscle differentiation have not been established. In addition to IGF-1, other secreted proteins have been demonstrated to perturb skeletal muscle size. Myostatin, also called growth differentiation factor 8 (GDF-8), is a TGF-␤ family member and a negative regulator of muscle mass (26). Myostatin’s effect was demonstrated in studies with mice that were made null for the myostatin gene (30), and also by correlating increases in muscle mass that were observed in strains of cattle with a loss of myostatin (14, 18, 31); the loss of myostatin resulted in a more than doubling in muscle mass. It has been suggested that other TGF-␤ superfamily molecules, distinct from myostatin, play a role in modulating skeletal muscle size, since myostatin⫺/⫺ mice that are mated with mice that are transgenic for follistatin (TGfollistatin), which is capable of inhibiting not only myostatin, but also its close relative GDF-11, and other TGF-␤ molecules such as the activins, resulted in an even greater increase in muscle size (33). In vitro studies with myostatin have been performed on rodent cells. In these studies, it has been shown that myostatin can block the differentiation of myoblasts into myotubes (23, 29, 37, 51). Experiments both in vitro and in vivo have demonstrated that myostatin signals by first binding the type II

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MYOSTATIN INHIBITS TORC1

activin receptor, IIb, which then allows for interaction with type I receptors ALK4 or ALK5 (49). The binding of myostatin to these receptor complexes results in the phosphorylation and activation of the transcription factors Smad2 and Smad3, which translocate to the nucleus upon phosphorylation (35). In the current study, the myostatin signaling pathways downstream of the Smads are further elucidated. EXPERIMENTAL PROCEDURES

Cell culture and treatment. Human skeletal muscle cells (HuSkMC; Lonza) were cultured in growth medium (GM) consisting of skeletal muscle basal medium (skBM; Lonza) supplemented with 20% FCS (Amimed). Differentiation was initiated 24 to 48 h after seeding by changing to serum-free differentiation medium consisting of skBM. For small interfering RNA (siRNA) experiments, cells were transfected 24 h after seeding in GM and differentiation was initiated after another 24 h. For reporter gene experiments, HuSkMC were transfected 24 h after seeding with pGL3-CAGA12-luc in serum-free skBM for 4 h and then returned to GM for another 36 h, before compound addition. If effects on HuSkMC differentiation were assessed, compounds were added at the onset of differentiation and cells were differentiated into myotubes for up to 72 h. If effects on mature HuSkMC myotubes were assessed, compounds were added on 3-day differentiated myotubes for additional up to 48 h. For human embryonic kidney (HEK)-293T cells stably transfected with pGL3-CAGA12-luc, cells were seeded in serum-reduced medium (2% FCS) for 24 h, medium was removed, and cells were stimulated with a 1:1 mixture of mouse serum and serum-reduced medium (2% FCS) in the presence of 500 ng/ml Fc-TGF-␤ R (to measure myostatin/GDF-11 activity) for another 24 h. Biochemicals and antibodies. Myostatin, TGF-␤1, activin A, activin AB, activin B, growth differentiation factor 11 (GDF-11), bone morphogenetic factor 2 (BMP-2), bone morphogenetic factor 7 (BMP7), human TGF-␤ RIIb/Fc chimera (Fc-TGF-␤ R), and follistatin were from R&D Systems; long-R3-insulin-like growth factor I (IGF-I) and SB-431542 (SB) were from Sigma-Aldrich. Myostatin propeptide D76A was kindly provided by the Novartis Biologic Units (M. Zurini). Stock solutions were prepared either in phospho-buffered saline (PBS) supplemented with 0.1% bovine serum albumin (BSA; follistatin, Fc-TGF-␤ R, activin A, activin AB, and activin B), in 10 mM HCl (IGF-I), in 4 mM HCl supplemented with 0.1% BSA (myostatin, TGF-␤1, BMP-2, BMP-7, and GDF-11), or in dimethyl sulfoxide (SB). For immunostaining, primary antibody against myosin heavy chain (MyHC; clone A4.1025) was from Upstate, and secondary antibody [Alexa Fluor 488 F (AB’)] was from Invitrogen. For Western blot analysis, antibodies against phospho-Smad2 (Ser465/ 467) (pSmad2) and MyoD were from Upstate; antibodies against ␣-tubulin were from Sigma-Aldrich; antibodies against myogenin were from BD Transduction Laboratories; and antibodies against Smad2/3 [Smad2/3, phospho-Akt (Ser473)] (pAkt, catalog no. 9271), Akt, phospho-p70 S6 kinase (Thr389) (pp70), and phospho-FoxO1 (Thr24)/FoxO3 (Thr32) (pFoxO) were from Cell Signaling. As secondary antibodies, goat anti-rabbit IgG horseradish peroxidase (HRP) and goat anti-mouse IgG HRP from SuperSignal West Femto Maximum Sensitivity Substrate (Pierce) were used. For ELISA, as capture antibodies, pSmad2 (Upstate) or Smad2/3 (Cell Signaling) were used. As detection antibody, Smad2/3 (BD Transduction Laboratories) were used, and as secondary antibody, anti-mouse IgG HRP was used (Cell Signaling). Western blot analysis. For HuSkMC, lysis buffer consisting of phosphosafe extraction reagent (Novagen) supplemented with 1% protease inhibitor cocktail (Sigma-Aldrich) was added. Homogenates were centrifuged for 10 min at 4°C (14,000 rpm). Supernatants were collected, and protein contents were measured using bicinchoninic acid (BCA) kit for protein determination (Sigma-Aldrich). Samples AJP-Cell Physiol • VOL

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were diluted in SDS-PAGE sample buffer and denatured for 5 min at 95°C. Equal amounts of protein were loaded per lane of 10% Bis-Tris NuPAGE gels, electrophoresed, and then transferred onto nitrocellulose membranes. Membranes were blocked in TBS with 5% (wt/vol) nonfat milk powder. Primary antibodies were incubated in Trisbuffered saline (TBS) with 0.1% Tween 20 and either 3% BSA (pAKT, AKT, MyoD, myogenin, and tubulin), 5% (Smad2/3, pp70S6, and pFoxO), or 5% nonfat milk (pSmad2), and secondary antibodies were incubated in TBS with 0.1% Tween 20, 0.05% SDS, and 5% nonfat milk. Immunoreactivity was detected by SuperSignal West Femto Maximum Sensitivity Substrate and exposed to film. RNA analysis. RNA was isolated using a RNEasy Mini Kit (Qiagen), per the manufacturer’s protocol. cDNA synthesis was performed using a QuantiTect Kit (Qiagen). Quantitative real-time PCR was performed using an Applied Biosystems 7500 FAST real-time PCR machine. The Taqman probe sets MuRF-1 (Hs00261590_m1), MAFbx (Hs00369709_m1), MyoD (Hs00159528_m1), myogenin (Hs00231167_m1), Smad2 (Hs00998186_m1), and Smad3 (Hs00232222_m1) were purchased from Applied Biosystems. siRNA. The siRNA pools designed against Smad2, Smad3, RICTOR, RAPTOR, and a control (NTC) targeting a not known mammalian sequence together with DharmaFECT 1 from Dharmacon were used. Final siRNA concentrations were 100 nM. Immunostaining. After washing with cytoskeleton stabilizing buffer (CSB), consisting of 80 mM PIPES, 5 mM EGTA, 1 mM MgCl2, 40 g/l PEG3500 in distilled water (pH 7.4), cells were fixed with 4% paraformaldehyde in CSB. Cells were then permeabilized with 0.2% Triton in CSB, and nonspecific binding was blocked with normal goat serum (Zymed) followed by incubation with MyHC diluted in PBS and subsequently with Alexa Fluor 488 F (AB’) diluted in PBS. Cells were mounted with ProLong Gold antifade reagent with DAPI (Invitrogen). Reporter gene assays. The reporter plasmid pGL3-CAGA12-luc, containing the TGF-␤-responsive reporter gene construct CAGA12luc cloned into the reporter construct pGL3 (Promega), and HEK293T cells stably transfected with this reporter plasmid were kindly provided by C. Lu (Developmental and Molecular Pathways, Novartis Pharma). Transfection in HuSkMC was performed using 50 ng DNA (pGL3-CAGA12-luc), PLUS reagent (Invitrogen), and Lipofectamine reagent (Invitrogen). Reporter gene activity was measured using Britelite Plus (Perkin Elmer); chemiluminescence was read using a Spectramax M5 (Molecular Devices). Creatine kinase activity assay. Cells were washed three times with PBS and then lysed with reporter lysis buffer (Promega) and stored until measurement at ⫺80°C. Muscle lysates were prepared in lysis buffer consisting of phosphosafe extraction reagent supplemented with 1% protease inhibitor cocktail. Creatine kinase (CK) activity was measured using the CK (IFCC) reagent (Thermo Electron). The CK reagent was prepared according to the manufacturers instructions. Lysates were adjusted to room temperature, CK reagent was added, and absorbance was immediately read at 340 nm for 20 min; reading interval was 1 min. CK standard curves were freshly prepared using CK from rabbit muscle (Roche Diagnostics). Protein content was determined using BCA kit. ELISA. For Smad2, ELISA plates were coated with capture antibodies (pSmad2 or Smad2/3) overnight at 4°C. Nonspecific binding was blocked with PBS containing 5% nonfat milk for 2 h. Cell lysates diluted 1:10 in PBS with 5% nonfat milk containing 0.05% Tween 20 were added and incubated overnight at 4°C. Detection antibody (Smad2/3) was added for 1.5 h at room temperature followed by incubation with secondary antibody (anti-mouse IgG) 1 h at room temperature. After addition of SuperSignal ELISA Femto Maximum Sensitivity substrate (catalog no. 37074; Pierce), chemiluminescence was read using a Spectramax M5 (Molecular Devices). For Akt, Pathscan pAkt and Akt ELISA kits from Cell signaling were used per manufacturer’s protocol.

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Diameter measurement. To analyze diameters, four pictures were taken per well and from each picture the 30 biggest myotubes were measured (using Cell imaging software from Olympus); myotube diameters were determined as average from three independent measurements per myotube. Afterward, the 10 biggest myotubes from each picture were selected, and the means ⫾ SE were calculated for each well (i.e., mean of 40 biggest myotubes on the four pictures). In vivo experiments. Female CB17/ICR-Prkdcscid/Crl mice (Scid mice; Charles River, Research Models and Services, Sulzfeld, Germany) aged 8 –9 wk were used. Animal experimentation was carried out according to the regulations effective in the Canton of Basel-City, Switzerland, and all protocols were approved by the Animal Welfare Committee of Novartis Institutes for Biomedical Research Basel. Myostatin propeptide D76A mutant was administered at a dose of 7.25 mg/kg ip on days 0, 3, 7, 14, and 21 at a volume of 5 ml/kg. Body weights were determined at the start of the experiment (day 0) and two times per week thereafter. Twenty-eight days after the start of the experiment (day 0), mice were euthanized under CO2 anesthesia. Serum and muscle samples were taken for analysis of muscle weight, CK activity, and serum myostatin activity. Statistical analysis. Differences between groups were analyzed using one-way ANOVA for all experiments. Values were considered statistically significant at P ⬍ 0.05. Concentration-response data were evaluated by sigmoid curve fitting (Using XLfit software) yielding EC50 values (concentrations causing half-maximal effects) and Emax (maximal effects). Body weight changes were determined against initial body weight taken at day 0 (first day of treatment). RESULTS

Establishment of human skeletal myoblast (HuSkMC) system (by demonstrating inhibition of differentiation by myostatin, several other TGF-␤ family members, and by using an inhibitor of ALK4, 5, and 7 receptors). An effort was made to find an in vitro muscle cell culture system that responded to myostatin at physiological levels and that would serve as a model for how myostatin might act in the human subject. Therefore, primary human myoblasts (HuSkMC) were used for this study. The effects on myogenic differentiation were assessed by attempting to differentiate HuSkMC in either the absence or presence of recombinant myostatin (1.0 ng/ml to 100 ng/ml), for 3 days after switching confluent myoblasts to differentiation media. The cultures were then fixed and stained with DAPI, to visualize nuclei, and were treated with antibodies to MyHC (Fig. 1A). Myostatin treatment resulted in a decrease in both the size and the number of HuSkMC myotubes, and these decreases occurred in a concentration-dependent manner (Fig. 1, C and D). Subsequent analysis of the fusion index and the diameters of the myotubes demonstrated that myostatin treatment inhibited the fusion index by up to 68% (Fig. 1C) and inhibited the myotube diameter by up to 37% (Fig. 1D). Moreover, CK activity, which increases during differentiation, is also reduced by myostatin, with an IC50 of 0.59 ng/ml (Fig. 1B) and a maximal inhibition of 84% (Fig. 1E). We next analyzed the sensitivity of HuSkMC differentiation to other TGF-␤ family members: TGF-␤1, GDF-11, activins, BMP-2, and BMP-7 (Fig. 1B and data not shown). All of these proteins inhibited HuSkMC differentiation (Fig. 1B). The rank order of potency was TGF-␤1 (IC50 ⫽ 0.20 ng/ml) ⱖ GDF-11 (0.23 ng/ml) ⬎ myostatin (0.59 ng/ml) ⬎ activin B (1.3 ng/ml, not shown) ⬎ activin AB (2.4 ng/ml; not shown) ⬎ activin A (5.0 ng/ml) ⱖ BMP-7 (5.6 ng/ml; not shown) ⬎ BMP-2 (29 AJP-Cell Physiol • VOL

ng/ml) (Fig. 1B). These data demonstrate that multiple TGF-␤ family members can inhibit skeletal muscle differentiation, including some that have been previously shown to signal via Smads that are distinct from those activated by myostatin and GDF-11. Finally, myogenic cells were shown to produce and release active myostatin and other TGF-␤ proteins, which can then act in an autocrine manner (36). To determine the endogenous tone of myostatin-like activity in HuSkMC, these cells were switched into differentiation media and then were either left unperturbed, as control, or were treated with follistatin (1 ␮g/ml), an inhibitor of myostatin, GDF-11, and activin, or with a pharmacologic inhibitor of the type I ALK receptors, SB431542 (SB; 1 ␮M). After 3 days, the degree of differentiation was assessed. Both follistatin and SB markedly increased both the number and size of HuSkMC myotubes, indicating facilitation of HuSkMC differentiation, and thus implying that the muscle cells autocrinely produce either myostatin or related molecules (Fig. 1, C and D). The fusion index was increased by 50% to 75%, and myotube diameters were enlarged by 39% to 61% (Fig. 1C). Also, CK activity was increased by 74% to 130% (Fig. 1E). Moreover, both follistatin and SB significantly reduced inhibition of HuSkMC differentiation by myostatin (10 and 100 ng/ml) as indicated by increasing the fusion index, myotube diameter, and CK activity, above the levels seen in untreated differentiating myotubes (Fig. 1E). These data demonstrate that HuSkMC differentiation is opposed by the autocrine production of myostatin and related molecules, and that myostatin can be blocked using a pharmacologic agent that inhibits ALK 4/5/7. Inhibition of HuSkMC differentiation by TGF-␤ family members requires Smad2 and Smad3 signaling. Next, we sought to determine the relative contribution of Smad2 and Smad3 activation to the effects of myostatin and other TGF-␤ family members on HuSkMC differentiation. A phosphorylation-specific antibody demonstrated activation of Smad2/3 after an hour of treatment with myostatin, by both Western blot analysis (Fig. 2A, left) and ELISA analysis, used to quantify Smad levels (Fig. 2A, right). Curve fitting of the phospho-Smad2 ELISA data yielded an IC50 of 8.9 ng/ml, and a maximal increase in Smad phosphorylation of approximately eightfold (Fig. 2A, right). Similar induction of Smad2 phosphorylation was observed with other TGF-␤ family members (data not shown). To discriminate between the requirement for Smad2 and Smad3, HuSkMC were treated with siRNA directed against either gene, 24 h before differentiation, and then differentiated in the absence or presence of TGF-␤ members or SB (Fig. 2, B and C). Smad2 mRNA and protein levels were selectively reduced by siSmad2 treatment, as were Smad3 levels by siSmad3 treatment; furthermore, the siRNAs were demonstrated to be specific (Fig. 2B). Knockdown of either Smad2 or Smad3 alone tended to reduce inhibition of HuSkMC differentiation by myostatin and TGF-␤1, but the concomitant knockdown of both Smad2 and Smad3 had an additive effect, as indicated by the significant rescue of fusion index, myotube diameter, and CK activity (Fig. 2C). Similarly, inhibition of HuSkMC differentiation by GDF-11, activin A, or by BMP-7 was reduced after Smad2 and Smad3 knockdown. As a specificity control, a proinflammatory cocktail, consisting of IL-1␣, TNF-␣, and INF-␥, was used. The effects of this cocktail were not perturbed by the Smad siRNAs (data not shown).

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Fig. 1. Inhibition of human skeletal muscle cells (HuSkMC) differentiation by myostatin and other transforming growth factor-␤ (TGF-␤) family members. HuSkMC were differentiated into myotubes for 3 days in the presence of myostatin or other TGF-␤ family members. The myostatin inhibitor follistatin and the activin receptor-like kinase (ALK) 4, 5, and 7 inhibitor SB-431542 (SB) were tested for blockade of myostatin activity and to determine the endogenous tone of myostatin-like signaling in HuSkMC (cells treated with follistatin or SB alone). A: myotubes differentiated for 3 days, in the absence [control (Con)] or presence of myostatin (1–100 ng/ml; Myo), were stained with anti-myosin heavy chain (MyHC) and DAPI. Shown are representative pictures. B: creatine kinase (CK) activity was analyzed from HuSkMC myotubes treated with either myostatin or the indicated TGF-␤ family member differentiated for 3 days. Data are expressed as percentage of control (no TGF-␤ family members). IC50 values were obtained by logistic curve fitting. Shown are means ⫾ SE from 3 to 7 independent experiments. GDF-11, growth differentiation factor 11; BMP-2, bone morphogenetic protein 2. C–E: analysis of fusion index (C), myotube diameter (from pictures in A) (D), and CK activity (E) from HuSkMC myotubes differentiated for 3 days. Data are expressed as percentage of control (no myostatin and no myostatin inhibitor; first bar). Shown are means ⫾ SE from 3 to 8 independent experiments. Differences from control (no myostatin and no myostatin inhibitor; first bar): *P ⬍ 0.05; differences from myostatin alone: #P ⬍ 0.05. AJP-Cell Physiol • VOL

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Myostatin inhibits the Akt pathway and can be opposed by IGF-1 treatment. IGF-1 has been characterized as a positive regulator of muscle differentiation, acting via the phosphatidylinositol 3-kinase/Akt pathway (10, 39). We therefore were

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interested in determining whether myostatin might act by perturbing Akt signaling, as had been previously demonstrated in murine cardiac cells (32), and, if so, whether IGF-1 treatment could oppose this effect. While the ALK inhibitor SB

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Fig. 3. Myostatin inhibits, whereas ALK4/5/7 inhibition facilitates the Akt pathway in differentiating HuSkMC; IGF-I can block myostatin’s antidifferentiation effects. A: ELISA of pSmad2 and total Smad2/3 protein (using an antibody that blots both Smad2 and Smad3), pAkt, and Akt. Shown are an ELISA of pSmad2 and Smad2/3 proteins (left) and an ELISA of pAkt and Akt proteins (right) from samples of HuSkMC treated with either nothing, myostatin, SB, IGF-1, or combinations, as indicated, for 24 h, starting at the onset of differentiation. Data are expressed as percentage of control (no treatment; first lanes). Shown are representative ELISAs. B: immunoblotting of pAkt and Akt, pFoxO, pp70S6, and ␣-tubulin from samples of HuSkMC treated with either nothing, myostatin, SB, IGF-1, or combinations, as indicated. Left: immunoblots of samples treated with compounds for 24 h, starting at the onset of differentiation. Right: blots of samples treated with siRNA against Smad2, Smad3, and NTC (the scrambled siRNA control) for 24 h before onset of differentiation. Myostatin was added for 24 h at onset of differentiation. Shown are representative immunoblots. C: analysis of CAGA-luc reporter gene assay (RGA). Shown is reporter gene activity from samples of HuSkMC transiently transfected with pGL3-CAGA12-luc, and treated with either myostatin alone, or in combination with IGF-1 as indicated. RGA data are expressed as chemiluminescence units. Shown are means ⫾ SE from 2 to 3 independent experiments. D: analysis of CK activity from HuSkMC myotubes differentiated for 3 days, and treated with nothing, myostatin alone, or myostatin in combination with IGF-I, as indicated. Data are expressed as percentage of control (no IGF-I, no myostatin). Shown are means ⫾ SE from 3 to 6 independent experiments. Differences from myostatin alone: #P ⬍ 0.05.

used as positive control significantly inhibited both basal levels of Smad2/3 phosphorylation, and those induced by myostatin (by up to 80%; Fig. 3A, left), 24 h of IGF-1 treatment had no effect on myostatin-induced Smad2/3 phosphorylation (Fig. 3A, left; compare bars 3 and 8). To analyze potential effects of IGF-I on Smad2/3 function, we assessed Smad reporter activity. IGF-1 treatment had no effect on myostatin-induced Smad reporter activity (Fig. 3C), indicating that IGF-I could not oppose myostatin by these mechanisms.

Treatment of human myoblasts with myostatin for 24h decreased phosphorylation of Akt by up to 50% (Fig. 3A, right, and Fig. 3B, left, top row). In addition to inhibiting Akt, myostatin decreased phosphorylation of p70S6 kinase and the pro-atrophy transcription factor FoxO1, both of which are normally phosphorylated by Akt (Fig. 3B, left). However, Akt, p70S6 kinase, and FoxO1 phosphorylation were restored by treatment with IGF-1, indicating that IGF-1/Akt signaling is dominant over myostatin/Smad/Akt inhibition (Fig. 3B). The

Fig. 2. Inhibition of HuSkMC differentiation by myostatin and TGF-␤ requires both Smad2 and Smad3. A: immunoblotting and ELISA of pSmad2 and Smad2/3. Left: immunoblots. Right: ELISA data of samples from HuSkMC treated with myostatin for 1 h starting at the onset of differentiation. ELISA data are expressed as chemiluminescence units. Shown are representative immunoblots or means ⫾ SE from 3 to 4 independent experiments (ELISA). B: RT-PCR and Western analysis of Smad2 and Smad3, in the absence or presence of small interfering RNAs (siRNAs). Shown are RT-PCR analysis of Smad2 and Smad3 mRNA (left) and an immunoblot of Smad2/3 (right) from samples of HuSkMC that were treated with siRNA to either control (NTC, a scrambled sequence), Smad2 (siSmad2), Smad3 (siSmad3), or both, as indicated. HuSkMC were then differentiated into myotubes for 3 days. RT-PCR data are expressed as relative quantification (RQ) against NTC. Representative experiments are shown. C: analysis of fusion index, myotube diameter, and CK activity from HuSkMC myotubes differentiated for 3 days, treated with siRNA against Smad2 (siSmad2), Smad3 (siSmad3), or NTC. Twenty-four hours after siRNA treatment, HuSkMC were differentiated into myotubes for 3 days in the absence or presence of compounds. Data are expressed as percentage (fusion index), micrometer (myotube diameter), or mUnits/mg protein (CK activity). Shown are means ⫾ SE from 3 to 7 independent experiments. Differences from control (no compound in NTC group; first bar): *P ⬍ 0.05; differences from NTC: #P ⬍ 0.05. AJP-Cell Physiol • VOL

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inhibition of Akt by myostatin could be blunted using siRNA to either Smad2 or Smad3 (not shown). Treatment with both siRNAs effectively restored Akt (Fig. 3B, right) and pFoxO signaling (data not shown). These data demonstrate both that myostatin blocks Akt signaling, and that IGF-1 can rescue Akt signaling without perturbing Smad2/3 activation by myostatin. In addition to restoring Akt phosphorylation, IGF-1 also partially rescues the differentiation of myostatin-treated myoblasts, as determined by measuring fusion index, diameters (data not shown), and CK activity (Fig. 3D). These data therefore suggest that myostatin’s inhibition of differentiation might in part be due to its inhibition of Akt signaling, since restoration of Akt phosphorylation is coincident with the partial rescue of differentiation (Figs. 3D and 8). Role of Akt/TORC pathways in myostatin’s inhibition of HuSkMC differentiation. Since myostatin signaling negatively regulates Akt, we sought to determine the contribution of the pathways downstream of Akt on myostatin signaling. To study the effects of the Akt/TORC pathways, we used siRNA against RICTOR to inhibit TORC2, and siRNA against RAPTOR to inhibit TORC1 (Figs. 4 and 8). Treatment with RICTOR siRNA led to an inhibition of HuSkMC differentiation by itself, as indicated by a reduction in CK activity, by 28% (Fig. 4A); this blockade of differentiation was accompanied by an almost total inhibition of pAKT (Fig. 4B, fourth row, compare lane 1 to lane 7), as had been recently demonstrated in the setting of a muscle-specific deletion of RICTOR (20). Inhibition of RICTOR caused an additive effect with myostatin, further inhibiting differentiation of HuSkMC myoblasts (Fig. 4A), and a further reduction of pAkt, (Fig. 4B, fourth row, compare lanes 1, 7, and 8). Inhibition of RICTOR tended to also increase TGF-␤1 inhibition of both differentiation (Fig. 4A), but did not reach statistical significance. In contrast to siRNA to RICTOR, and as expected, siRNA inhibition of RAPTOR was not sufficient to decrease pAkt473, and if anything may somewhat increase pAkt (Fig. 4B, fourth row, compare lanes 1 and 10, and Fig. 8), as recently reported (1). While inhibition of RAPTOR by itself also did not block differentiation, it significantly facilitated myostatin’s inhibitory action on differentiation (Fig. 4A). Interestingly, in the presence of siRNA to RAPTOR there was a greater increase in pSmad2 than with myostatin alone (Fig. 4B, second row, compare lanes 2 and 11), indicating that blockade of Akt/ TORC1 positively feeds back to the myostatin pathway, increasing myostatin-induced activation of Smad2. In contrast, action of TGF-␤1 on either differentiation (Fig. 4A) or pSmad2 signaling (Fig. 4B) was not significantly perturbed by siRNA to RAPTOR, indicating subtle differences in the mode of action between myostatin and TGF-␤1. Myostatin reduces diameters of mature HuSkMC myotubes but does not induce the E3 ubiquitin ligases MuRF1 and MAFbx. It had previously been reported that when active myostatin was delivered in adult mice, muscle atrophy was observed, as determined by a decrease in muscle mass and fiber diameter (52). We therefore sought to determine the effect of myostatin on differentiated HuSkMC myotubes, and the mechanism by which this effect occurred. As a control for the ability to modify myotube diameter, IGF-1 was also used to treat HuSkMC myotube cultures (data not shown, but see Fig. 6A). Myotubes were allowed to form for 3 days before treatments were initiated. As previously reported for both mouse (38) and AJP-Cell Physiol • VOL

Fig. 4. RAPTOR inhibition facilitates inhibitory effects of myostatin, but not TGF-␤1, on HuSkMC differentiation and Smad signaling. A: analysis of CK activity from HuSkMC myotubes differentiated for 3 days, treated with siRNA against RICTOR (siRIC), RAPTOR (siRAP), or NTC (scrambled siRNA, as a control). Twenty-four hours after siRNA treatment, HuSkMC were differentiated into myotubes for 3 days in the absence or presence of compounds. Data are expressed as mUnits/mg protein (CK activity). Shown are means ⫾ SE from 3 to 4 independent experiments. Differences from control (no compound in NTC group; first bar): *P ⬍ 0.05; differences from NTC: #P ⬍ 0.05. B: immunoblotting of pFoxO, pSmad2, and Smad2/3 (using an antibody that cross-reacts between total Smad2 and Smad3 proteins), pAkt, and Akt, from samples of HuSkMC treated with either scrambled siRNA (NTC, as a control), or with siRIC, siRAP, or siSmad2/3, starting 24 h before onset of differentiation.

human (16) myotubes, IGF-1 was able to induce significant myotube hypertrophy (data not shown, but see Fig. 6A). In contrast, myostatin exposure resulted in a significant decrease in myotube diameter by up to 37% (Fig. 5A, photographs; quantified in Fig. 5A, top bar graph). The E3 ubiquitin ligases MuRF1 and MAFbx have been shown to be upregulated in multiple physiological settings of atrophy (12). Therefore we next asked whether myostatin increases the expression of these genes, as had previously been reported (29). Surprisingly, rather than an increase in expression, we observed a decrease in both MuRF1 and MAFbx levels, by up to 66% (Fig. 5B); only the highest concentrations of myostatin, 100 ng/ml, began to push expression back up toward that seen in unperturbed myotubes (Fig. 5B). Since both MuRF1 and MAFbx are induced from very low levels, to their basal levels, upon muscle differentiation (data not shown), and since myostatin decreases differentiation when applied to myoblasts, we next asked whether the decrease in MuRF1 and MAFbx expression was a

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Fig. 5. Myostatin reduces diameters of mature HuSkMC myotubes but does not induce atrophy markers, at physiological concentrations. HuSkMC were differentiated into myotubes for 3 days and treated with myostatin for an additional 48 h. A: 3-day differentiated HuSkMC myotubes treated with myostatin for 48 h were stained with MyHC and DAPI. Shown are representative pictures in lower (top) and higher (bottom) magnification. Arrows (top) highlight identical myotubes at bottom, shown at higher magnification. In bar graphs at right, myotube diameter (top) and CK activity (bottom) of HuSkMC that were treated with increasing concentrations of myostatin, as indicated, were analyzed. For myotube diameters, data are expressed as micrometers. Shown are means ⫾ SE from 3 to 5 independent experiments. Differences from control (no myostatin; first bar): *P ⬍ 0.05. For CK activity, data are expressed as percentage of control (no myostatin; first bar). Shown are means ⫾ SE from 4 independent experiments. Differences from control: *P ⬍ 0.05. B: analysis of muscle RING-finger 1 (MuRF1; left) and muscle atrophy F-box (MAFbx; right) mRNA (by RT-PCR) from samples of 3-day differentiated HuSkMC myotubes treated for 48 h with myostatin. Data are expressed as RQ (relative quantification against control; first bars). Shown are means ⫾ SE from 2 to 3 independent experiments. C: analysis of MyoD and myogenin mRNA (by RT-PCR) from samples of 3-day differentiated HuSkMC myotubes treated for 48 h with myostatin. Data are expressed as RQ (relative quantification against control; first bars). Shown are means ⫾ SE from 2 to 3 independent experiments.

sign of a more general blockade in differentiation-regulated genes. When applied to human myotubes, myostatin decreased CK activity up to 50% (Fig. 5A, bottom right), and also the levels of myogenin and MyoD, by up to 82% (Fig. 5C). These AJP-Cell Physiol • VOL

data are in line with a previous report that myostatin blocks muscle differentiation-associated gene expression in myotubes (9), and extends those findings even to genes such as the pro-atrophy program. Therefore, instead of myostatin inducing

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Fig. 6. ALK4/5/7 inhibition and IGF-I reduce myostatin action in mature HuSkMC myotubes. HuSkMC were differentiated into myotubes for 3 days and treated with nothing (⫺), myostatin, SB, follistatin, IGF-1, or combinations, as indicated, for an additional 24 to 48 h. A: analysis of myotube diameters from HuSkMC myotubes treated for 48 h with myostatin alone or in combination with SB or IGF-I. Data are expressed as percentage of control (no treatment; first bar). Shown are means ⫾ SE from 7 to 8 independent experiments. Differences from control: *P ⬍ 0.05; differences from myostatin alone: #P ⬍ 0.05. B: immunoblotting of MyoD vs. tubulin (as a loading control), and myogenin as compared with tubulin, from HuSkMC myotubes treated for 48 h with myostatin alone or in combination with SB, follistatin, or IGF-I. Shown are representative immunoblots. C: immunoblotting of pSmad2 and Smad2/3 (left) and pAkt and Akt (right) from samples of HuSkMC myotubes treated for 24 h with myostatin alone or in combination with SB and IGF-I. Shown are representative immunoblots. Smad2/3 signaling is increased, and Akt signaling is reduced, by myostatin, in a concentration-dependent fashion.

the reported “atrophy pathway,” it apparently causes a decrease in muscle mass by inhibiting the expression of genes required for muscle maintenance (Fig. 5C). ALK receptor inhibition or IGF-I treatment reduces myostatin’s effects on differentiated HuSkMC myotubes. To determine whether the myostatin-induced reduction in size of HuSkMC myotubes was caused by the ActRII/ALK receptor complex, we tested the ALK receptor inhibitor SB (Fig. 6A). SB was able to block the decrease in myotube diameter caused by myostatin (Fig. 6A, compare bars 2 and 4). Treatment of myotubes with SB alone caused an increase in myotube size by 21%. Myotubes treated with myostatin demonstrated a decrease in size by over 25%, which was opposed by simultaneous treatment with IGF-1 (Fig. 6A). IGF-1 also blocked the decrease in expression of muscle-specific proteins, MyoD and myogenin, caused by myostatin as did the myostatin inhibitor, follistatin and SB (Fig. 6B). In myotubes, IGF-1 blockade of myostatin’s efficacy was again not at the level of Smad2/3, but rather by maintaining Akt signaling (Fig. 6C). As was observed in myoblasts, myostatin was able to decrease Akt phosphorylation in a dose-responsive fashion in myotubes (Fig. 6C, right). These data indicate that myostatin and IGF-1 have similar effects on differentiation markers and Akt signaling in both myoblasts and myotubes, resulting in distinct phenotypes (blockade of differentiation in myoblasts, vs. a decrease in myotube size in myotubes). AJP-Cell Physiol • VOL

Increased muscle weight and CK activity by myostatin inhibition in vivo. We next performed in vivo studies to see whether it was possible to correlate the perturbation of CK activity with changes in muscle size, as had been observed in vitro. Scid mice were treated with the mutant myostatin D76A propeptide for 4 wk (Fig. 7A). In the myostatin propeptide-treated group, body weight as well as the muscle weight of soleus (16%) , tibialis (9%), and quadriceps (9%) muscles were significantly increased compared with PBS-treated controls (Fig. 7A). In addition, the weight of gastrocnemius (by 4%) and pectoralis (by 7%) muscles also tended to increase (Fig. 7A) without reaching statistical significance. Moreover, CK activity in gastrocnemius muscle lysates was significantly increased after application of myostatin D76A propeptide, suggesting increased differentiation of skeletal muscle in vivo (Fig. 7A). Since myostatin has been shown to circulate in the bloodstream, we were interested in measuring its activity from serum samples as biomarker using a myostatin-responsive reporter (the gene plasmid pGL3-CAGA12-luc, transfected into HEK293T cells). Scid mice were treated with a single dose of myostatin D76A propeptide, and serum samples were taken 24 h after treatment. To isolate the myostatin/GDF-11 component reporter, gene activity was measured in the presence of Fc-TGF-␤ RIIb (data not shown). In the myostatin propeptidetreated group, serum activity was reduced by 72% compared with PBS-treated controls (Fig. 7B).

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Fig. 7. Increased muscle mass, CK activity, and reduced serum activity in mice treated with myostatin propeptide. A: analysis of body and muscle weight as well as CK activity from Scid mice treated for 4 wk with myostatin propeptide D76A. Data are expressed as changes in body weight (g), percent increase of muscle weight (vs. control PBS group), or units/mg protein (CK activity). Shown are means ⫾ SE from 12 animals. Differences from control (PBS group): *P ⬍ 0.05. B: analysis of serum myostatin/GDF-11 activity using CAGA-luc reporter gene assay (RGA) of Scid mice 24 h after treatment with myostatin propeptide. Shown is reporter gene activity from samples of HEK-293T cells stably transfected with pGL3-CAGA12-luc. RGA data are expressed as chemiluminescence units. Shown are means ⫾ SE from 4 serum samples. Differences from control (PBS group): *P ⬍ 0.05.

DISCUSSION

In this study we report on the effects of myostatin and other TGF-␤ molecules on human skeletal myoblasts (HuSkMC) and myotubes. These HuSkMCs respond to myostatin at physiological concentrations, 0.1–300 ng/ml, resulting in a decrease in fusion index, myotube diameter, CK activity, and expression of MyoD and myogenin. It was previously demonstrated that follistatin, a more general inhibitor of TGF-␤ molecules, could induce an additive increase in muscle mass when combined with myostatin (25). In this study, a range of other TGF-␤ molecules are shown to be able to block muscle differentiation, including the more distantly related activins, and BMP-2. This is an important demonstration, since many therapeutic approaches either in or heading toward the clinic are focused on myostatin alone. Myostatin’s effects in the HuSkMC system were confirmed to be via ALK receptors, as had been previously shown in other systems, since SB, an inhibitor that targets ALKs 4, 5, and 7, can restore myostatin-inhibited differentiation. Treatment with siRNAs that inhibit Smad2, Smad3, or both Smad2 and 3 simultaneously demonstrated that myostatin’s antidifferentiation effects require both of these Smads; while inhibition of either Smad2 or Smad3 could blunt myostatin’s effects, simultaneous blockade of both Smads had an additive AJP-Cell Physiol • VOL

effect. These results are in agreement with results from the Sandri group, published in a companion article (44a). We have previously shown that myotubes could be induced to hypertrophy by the IGF-1/Akt pathway, which induces protein synthesis required for muscle hypertrophy, and which blocks protein degradation ubiquitination pathways that are active during skeletal muscle atrophy (12). Akt induces protein synthesis via the Akt/mTOR/p70S6K pathway (5, 38) (Fig. 8). Furthermore, Akt blocks the upregulation of MuRF1 and MAFbx, by phosphorylating and inhibiting the FoxO family of transcription factors (43, 48) (Fig. 8). MuRF1 and MAFbx encode E3 ubiquitin ligases, which are required for muscle atrophy (4). In the current study, it was demonstrated that myostatin inhibits activation of Akt, in both myoblasts and myotubes. Akt signals via two mTOR signaling complexes, TORC1 and TORC2 (2). It was recently reported that muscle-specific ablation of RAPTOR, but not RICTOR, results in a dystrophic phenotype (1); a separate group reported that a muscle-specific deletion of RICTOR impaired insulin-stimulated activation of the Akt pathway (20). However, these signaling pathways had not been analyzed for their potential roles in myostatin signaling. In this study it was shown that inhibition of RICTOR, which is a component of TORC2, has a significant inhibitory

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Fig. 8. Signaling diagram illustrating myostatin, TGF-␤, and IGF-1 pathway interactions. Myostatin’s effects require both Smad2 and Smad3, which block muscle differentiation, including the E3 ligases MuRF1 and MAFbx, which are normally upregulated during muscle atrophy. This distinguishes the effects of myostatin with “typical atrophy.” Smad2 and 3 activation are both required for myostatin’s inhibitory effects on Akt. Inhibition of RAPTOR, and thus mammalian target of rapamycin (mTOR) complex 1 (TORC1), is additive with myostatin signaling. Smad2, 3 phosphorylation increases when RAPTOR is downregulated. RICTOR is itself sufficient to block differentiation. TGF-␤ family members that signal through the TGF-␤ type II receptor can also activate Smad2 and 3. Treatment with IGF-1 can counteract myostatin’s antidifferentiation effects, indicating the IGF-1/Akt pathway is dominant over the myostatin pathway. p70S6K, p70 S6 kinase; ActRII, activin receptor II; PI3K, phosphatidylinositol 3-kinase; PIP3, phosphatidylinositol-3, 4, 5-trisphosphate.

effect on muscle differentiation by itself. Relevant to this finding, mice that are homozygous nulls for RICTOR die during embryogenesis, at day 10.5 of development, and these embryos were smaller than wild-type controls (15, 46). The antidifferentiation effect of RICTOR that was observed is additive to that induced by myostatin, indicating that they may act independently of each other. In contrast, inhibition of RAPTOR, and thus TORC1, does not by itself block muscle differentiation, but does contribute to myostatin’s inhibitory effects; this suggests a cooperation between the two on Akt, at the serine 473 phosphorylation site. Furthermore, inhibition of RAPTOR potentiates myostatin-induced Smad2 phosphorylation, establishing a feed-forward mechanism: myostatin activates Smad2, which is required to block Akt, inhibiting TORC1, which in turn potentiates myostatin’s activation of Smad2. These findings are outlined in a signaling diagram (Fig. 8). Akt acts to inhibit the upregulation of the E3 ligases MuRF1 and MAFbx, which are required for skeletal muscle atrophy. When MuRF1 and MAFbx expression was assessed upon myostatin treatment of skeletal myotubes, we were surprised to see that these genes were in fact downregulated, along with myogenin, and MyoD. It is important to note that along with MyoD and myogenin, MuRF1 and MAFbx are induced upon muscle differentiation. Therefore, it appears that myostatin can block the induction of muscle’s differentiation program, even when applied to already formed differentiated muscle cells. Combined with the decrease in Akt-induced protein synthesis signaling, myostatin’s decrease in genetic differentiation programs would help to explain how it causes a decrease in skeletal muscle mass when applied to adult muscle. AJP-Cell Physiol • VOL

Addition of IGF-1 was shown to block the effects of myostatin, when applied to either myoblasts or myotubes. This finding is consistent with prior data indicating that IGF-1 is both a positive modulator of differentiation, hypertrophy, and Akt signaling; however, it is of interest that IGF-1/Akt signaling is dominant over myostatin-induced inhibition of Akt. The precise intersection between the two pathways may be multifold, but the current study demonstrates that Akt is a particular nexus, and that IGF-1 can rescue the activation of Akt that is blunted by myostatin. In vivo studies with an inhibitor of myostatin, the myostatin propeptide, demonstrated that CK activity could be increased by blocking myostatin, as occurred in vitro, and that this perturbation of differentiation correlates with myostatin inhibition’s established increase in muscle mass in the murine model. The rather modest increases in muscle mass of ⬃15% were consistent with a prior study where myostatin was inhibited in adult animals. This confirms that much of the increase in mass observed in myostatin-null animals, where muscle is approximately twofold greater than in wild-type mice, is a function of developmental effects of the protein. In summary, these data demonstrate that myostatin and its relatives are active on both myoblasts and myotubes, via a similar antidifferentiation mechanism, and that these phenotypes are mediated at least in part by perturbation of Akt/ TORC1 signaling. Furthermore, the demonstration that IGF-1 can dominantly overcome myostatin inhibition adds to the rationale for IGF-1-based treatment regimens in clinical settings where myostatin is active. ACKNOWLEDGMENTS We thank the Muscle Diseases group at the Novartis Institutes for Biomedical Research (NIBR) for enthusiastic support, along with the rest of the NIBR community, in particular, Mark Fishman, Brian Richardson, Andrew Mackenzie, and Estelle Trifilieff for support, and to Helene Rivet and Valerie Lefevre for assistance. Finally, thanks to Chris Lu for the TGF-␤ reporter constructs and cell lines as well as Mauro Zurini for the supply of myostatin propeptide D76A. REFERENCES 1. Bentzinger C, Romanino K, Cloe¨tta D, Lin S, Mascarenhas J, Oliveri F, Xia J, Casanova E, Costa C, Brink M, Zorzato F, Hall M, Ru¨egg M. Skeletal muscle-specific ablation of raptor, but not of rictor, causes metabolic changes and results in muscle dystrophy. Cell Metab 8: 411– 424, 2008. 2. Bhaskar PT, Hay N. The two TORCs and Akt. Dev Cell 12: 487–502, 2007. 3. Bingwen Jin YPL. Curcumin prevents lipopolysaccharide-induced atrogin1/MAFbx upregulation and muscle mass loss. J Cell Biochem 100: 960 –969, 2007. 4. Bodine SC, Latres E, Baumhueter S, Lai VK, Nunez L, Clarke BA, Poueymirou WT, Panaro FJ, Na E, Dharmarajan K, Pan ZQ, Valenzuela DM, DeChiara TM, Stitt TN, Yancopoulos GD, Glass DJ. Identification of ubiquitin ligases required for skeletal muscle atrophy. Science 294: 1704 –1708, 2001. 5. Bodine SC, Stitt TN, Gonzalez M, Kline WO, Stover GL, Bauerlein R, Zlotchenko E, Scrimgeour A, Lawrence JC, Glass DJ, Yancopoulos GD. Akt/mTOR pathway is a crucial regulator of skeletal muscle hypertrophy and can prevent muscle atrophy in vivo. Nat Cell Biol 3: 1014 – 1019, 2001. 6. Crepaldi T, Bersani F, Scuoppo C, Accornero P, Prunotto C, Taulli R, Forni PE, Leo C, Chiarle R, Griffiths J, Glass DJ, Ponzetto C. Conditional activation of MET in differentiated skeletal muscle induces atrophy. J Biol Chem 282: 6812– 6822, 2007. 7. Deruisseau KC, Kavazis AN, Deering MA, Falk DJ, Van Gammeren D, Yimlamai T, Ordway GA, Powers SK. Mechanical Ventilation induces alterations of the ubiquitin-proteasome pathway in the diaphragm. J Appl Physiol 98: 1314 –1321, 2004.

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MYOSTATIN INHIBITS TORC1 8. Dogra C, Changotra H, Wedhas N, Qin X, Wergedal JE, Kumar A. TNF-related weak inducer of apoptosis (TWEAK) is a potent skeletal muscle-wasting cytokine. FASEB J 21: 1857–1869, 2007. 9. Durieux AC, Amirouche A, Banzet S, Koulmann N, Bonnefoy R, Pasdeloup M, Mouret C, Bigard X, Peinnequin A, Freyssenet D. Ectopic expression of myostatin induces atrophy of adult skeletal muscle by decreasing muscle gene expression. Endocrinology 148: 3140 –3147, 2007. 10. Florini JR, Ewton DZ, Coolican SA. Growth hormone and the insulinlike growth factor system in myogenesis. Endocr Rev 17: 481–517, 1996. 11. Glass DJ. Molecular mechanism modulating muscle mass. Trends Mol Med 9: 344 –350, 2003. 12. Glass DJ. Skeletal muscle hypertrophy and atrophy signaling pathways. Int J Biochem Cell Biol 37: 1974 –1984, 2005. 13. Gomes MD, Lecker SH, Jagoe RT, Navon A, Goldberg AL. Atrogin-1, a muscle-specific F-box protein highly expressed during muscle atrophy. Proc Natl Acad Sci USA 98: 14440 –14445, 2001. 14. Grobet L, Martin LJ, Poncelet D, Pirottin D, Brouwers B, Riquet J, Schoeberlein A, Dunner S, Menissier F, Massabanda J, Fries R, Hanset R, Georges M. A deletion in the bovine myostatin gene causes the double-muscled phenotype in cattle. Nat Genet 17: 71–74, 1997. 15. Guertin DA, Stevens DM, Thoreen CC, Burds AA, Kalaany NY, Moffat J, Brown M, Fitzgerald KJ, Sabatini DM. Ablation in mice of the mTORC components raptor, rictor, or mLST8 reveals that mTORC2 is required for signaling to Akt-FOXO and PKCalpha, but not S6K1. Dev Cell 11: 859 – 871, 2006. 16. Jacquemin V, Furling D, Bigot A, Butler-Browne GS, Mouly V. IGF-1 induces human myotube hypertrophy by increasing cell recruitment. Exp Cell Res 299: 148 –158, 2004. 17. Jagoe RT, Lecker SH, Gomes M, Goldberg AL. Patterns of gene expression in atrophying skeletal muscles: response to food deprivation. FASEB J 16: 1697–1712, 2002. 18. Kambadur R, Sharma M, Smith TP, Bass JJ. Mutations in myostatin (GDF8) in double-muscled Belgian Blue and Piedmontese cattle. Genome Res 7: 910 –916, 1997. 19. Krawiec BJ, Frost RA, Vary TC, Jefferson LS, Lang CH. Hindlimb casting decreases muscle mass in part by proteasome-dependent proteolysis but independent of protein synthesis. Am J Physiol Endocrinol Metab 289: E969 –E980, 2005. 20. Kumar A, Harris TE, Keller SR, Choi KM, Magnuson MA, Lawrence JC Jr. Muscle-specific deletion of rictor impairs insulin-stimulated glucose transport and enhances basal glycogen synthase activity. Mol Cell Biol 28: 61–70, 2008. 21. Lai KMV, Gonzalez M, Poueymirou WT, Kline WO, Na E, Zlotchenko E, Stitt TN, Economides AN, Yancopoulos GD, Glass DJ. Conditional activation of Akt in adult skeletal muscle induces rapid hypertrophy. Mol Cell Biol 24: 9295–9304, 2004. 22. Lang CH, Huber D, Frost RA. Burn-induced increase in atrogin-1 and MuRF-1 in skeletal muscle is glucocorticoid independent but downregulated by IGF-I. Am J Physiol Regul Integr Comp Physiol 292: R328 – R336, 2007. 23. Langley B, Thomas M, Bishop A, Sharma M, Gilmour S, Kambadur R. Myostatin inhibits myoblast differentiation by down-regulating MyoD expression. J Biol Chem 277: 49831– 49840, 2002. 24. Latres E, Amini AR, Amini AA, Griffiths J, Martin FJ, Wei Y, Lin HC, Yancopoulos GD, Glass DJ. Insulin-like growth factor-1 (IGF-1) inversely regulates atrophy-induced genes via the phosphatidylinositol 3-kinase/Akt/mammalian target of rapamycin (PI3K/Akt/mTOR) pathway. J Biol Chem 280: 2737–2744, 2005. 25. Lee SJ. Quadrupling muscle mass in mice by targeting TGFbeta signaling pathways. PLoS ONE 2: e789, 2007. 26. Lee SW, Dai G, Hu Z, Wang X, Du J, Mitch WE. Regulation of muscle protein degradation: coordinated control of apoptotic and ubiquitin-proteasome systems by phosphatidylinositol 3 kinase. J Am Soc Nephrol 15: 1537–1545, 2004. 27. Levine S, Nguyen T, Taylor N, Friscia ME, Budak MT, Rothenberg P, Zhu J, Sachdeva R, Sonnad S, Kaiser LR, Rubinstein NA, Powers SK, Shrager JB. Rapid disuse atrophy of diaphragm fibers in mechanically ventilated humans. N Engl J Med 358: 1327–1335, 2008. 28. Li YP, Chen Y, Li AS, Reid MB. Hydrogen peroxide stimulates ubiquitin conjugating activity and expression of genes for specific E2 and E3 proteins in skeletal muscle myotubes. Am J Physiol Cell Physiol 285: C806 –C812, 2003. AJP-Cell Physiol • VOL

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