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Endocrinology Vol. 148, No. 7 3140-3147
Copyright © 2007 by The Endocrine Society

Ectopic Expression of Myostatin Induces Atrophy of Adult Skeletal Muscle by Decreasing Muscle Gene Expression

Anne-Cécile Durieux, Adel Amirouche, Sébastien Banzet, Nathalie Koulmann, Régis Bonnefoy, Marielle Pasdeloup, Catherine Mouret, Xavier Bigard, André Peinnequin and Damien Freyssenet

Unité Physiologie et Physiopathologie de l’Exercice et Handicap (A.-C.D., A.A., R.B., D.F.), EA3062, Université Jean Monnet, 42023 Saint-Etienne, France; Institute of Anatomy (A.-C.D.), University of Berne, CH-3000 Bern, Switzerland; and Département des Facteurs Humains (S.B., N.K., M.P., X.B.) and Département de Radiobiologie et Radiopathologie (C.M., A.P.), Centre de Recherche du Service de Santé des Armées, 38702 La Tronche, France

Address all correspondence and requests for reprints to: Damien Freyssenet, Unité Physiologie et Physiopathologie de l’Exercice et Handicap, Faculté de Médecine, 15 rue Ambroise Paré, 42023 Saint-Etienne cedex 2, France. E-mail: damien.freyssenet{at}univ-st-etienne.fr.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Myostatin is a master regulator of myogenesis and early postnatal skeletal muscle growth. However, myostatin has been also involved in several forms of muscle wasting in adulthood, suggesting a functional role for myostatin in the regulation of skeletal muscle mass in adult. In the present study, localized ectopic expression of myostatin was achieved by gene electrotransfer of a myostatin expression vector into the tibialis anterior muscle of adult Sprague Dawley male rats. The corresponding empty vector was electrotransfected in contralateral muscle. Ectopic myostatin mRNA was abundantly present in muscles electrotransfected with myostatin expression vector, whereas it was undetectable in contralateral muscles. Overexpression of myostatin elicited a significant decrease in muscle mass (10 and 20% reduction 7 and 14 d after gene electrotransfer, respectively), muscle fiber cross-sectional area (15 and 30% reduction 7 and 14 d after gene electrotransfer, respectively), and muscle protein content (20% reduction). No decrease in fiber number was observed. Overexpression of myostatin markedly decreased the expression of muscle structural genes (myosin heavy chain IIb, troponin I, and desmin) and the expression of myogenic transcription factors (MyoD and myogenin). Incidentally, mRNA level of caveolin-3 and peroxisome proliferator activated receptor {gamma} coactivator-1{alpha} was also significantly decreased 14 d after myostatin gene electrotransfer. To conclude, our study demonstrates that myostatin-induced muscle atrophy elicits the down-regulation of muscle-specific gene expression. Our observations support an important role for myostatin in muscle atrophy in physiological and physiopathological situations where myostatin expression is induced.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
A DECREASE IN skeletal muscle mass can be the consequence of ongoing physiological (ageing) and pathological processes (cancer, myopathies, peripheral neuropathies, respiratory insufficiency, sepsis). Whatever the inciting event, skeletal muscle atrophy is characterized by a decrease in protein content, fiber diameter, force production, and fatigue resistance, all factors contributing to a loss of autonomy and an increased institutionalization of the affected people.

Various hormones, growth factors, and cytokines have emerged as important regulators of skeletal muscle mass (reviewed in Refs. 1 and 2). Myostatin, a TGF-ß family member also called growth differentiation factor-8, has been identified as a master negative regulator of skeletal muscle mass during embryogenesis and early postnatal muscle growth (Refs. 3, 4, 5 , recently reviewed in Ref. 6). The most striking biological features of myostatin were observed in myostatin-null mice. These mice show a two- to three-times increase in skeletal muscle mass, mainly due to increases in both myofiber size and myofiber number (5). In vitro studies have then clearly established that the molecular basis of the inhibition of muscle development by myostatin involves the negative regulation of myoblast proliferation and differentiation (7, 8, 9, 10, 11, 12). Therefore, muscle fiber hypertrophy and hyperplasia encountered in myostatin-null mice were attributed to the negative role played by myostatin on myoblasts during embryogenesis and early postnatal growth (13).

Given its remarkable effects on skeletal muscle development, it has been suggested that myostatin may also contribute to the regulation of skeletal muscle mass in adulthood. Muscle disuse consecutive to chronic osteoarthritis or ageing, is associated with increased myostatin mRNA level (14) and protein level (15). Similarly, muscle wasting in HIV-infected men is also associated with elevated myostatin serum concentration (16). Injection of cells overexpressing myostatin in adult mice is accompanied by severe muscle loss (17). Conversely, an antibody strategy aimed at inhibiting myostatin activity was shown to increase skeletal muscle mass and strength in adult mice (18). Clearly, the rate and extent of muscle mass changes in these studies cannot be explained solely by an effect of myostatin on satellite cells, myoblast proliferation, and myoblast differentiation, and raises the possibility that this effect may directly result from the action of myostatin on skeletal muscle fibers.

Therefore, the aim of this study was to determine whether myostatin overexpression could induce skeletal muscle atrophy in adulthood. The effect of myostatin overexpression on muscle gene expression was also studied. Using gene electrotransfer of a myostatin expression vector into the tibialis anterior (TA) muscles of young adult rats, we report that myostatin overexpression rapidly lowers muscle mass by decreasing muscle fiber cross-sectional area without change in muscle fiber number. We also show that the expression of muscle-specific genes was markedly decreased by myostatin overexpression. Altogether, these data identify myostatin as a major regulator of skeletal muscle mass in adulthood.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Animals
Experiments were carried out on Sprague Dawley male rats (307.8 ± 2.8 g, n = 59) in the animal facilities of the Faculté de Médecine (Saint Etienne) according to the newest guiding principle for research (19).

Plasmid DNA
pcDNA-Myostatin, full-length murine myostatin cDNA into pcDNA3.1 Zeo expression vector (9). pMyoD-Luc, 2.7-kb fragment of the human MyoD gene (proximal promoter and the transcription start site) linked to the 4-kb far upstream enhancer region into pGL2 Basic luciferase reporter vector (20). pTnI-Luc, 2.3-kb promoter fragment of the quail troponin I gene into the pGL2 Basic luciferase reporter vector (21). pCMV-ß-galactosidase was from Clontech (Palo Alto, CA). Plasmids were amplified in JM109 bacteria, purified (EndoFree Plasmid Mega Kit; Qiagen, Valencia, CA), and dissolved in sterile endotoxin-free 0.9% saline solution.

Electrotransfer of plasmid DNA
Animals were anesthetized with an ip injection of sodium pentobarbital (60 mg/kg body weight). In a first experiment, 500 µg of pcDNA-myostatin (0.625 µg/µl) was injected into the entire TA muscle by using a 301/2-gauge needle. Contralateral TA muscle was similarly injected with the corresponding empty vector (pcDNA3.1 Zeo). Thirty seconds after injection, a total of 24 trains of 500 electric pulses (100 µs, 50 mA) were delivered (22, 23). In a second experiment designed to evaluate whether myostatin transactivates MyoD and troponin I promoters, the middle belly portion of TA muscle was injected with 50 µg of pcDNA-myostatin with either 50 µg of pMyoD-Luc or 50 µg of pTnI-Luc. pCMV-ß-galactosidase (50 µg) was coinjected to correct for variations in transfection efficiency. Contralateral TA muscle was similarly injected but pcDNA3.1 Zeo was used instead of pcDNA-myostatin. Thirty seconds after injection, eight trains of 125 electric pulses (100 µs, 50 mA) were applied.

Tissue collection
Seven (experiments 1 and 2) and 14 d (experiment 1) after gene electrotransfer, animals were anesthetized as described above. TA muscles were removed, weighed, and stored at –80 C for subsequent analyses. Animals were then killed by an overdose of sodium pentobarbital.

Histomorphometric analyses
Portions of TA muscles embedded in cryopreservative (Cryomount; Histolab, Göteborg, Sweden) were cut (12 µm) and stained with hemalun-eosin-safran. Eight to 11 photographs covering the entire muscle section were used to determine the whole muscle cross-sectional area. Five fields consistently positioned across muscle sections were chosen and the cross-sectional area of 60 fibers per field was counted. Fiber density was determined from the counts of the number of muscle fibers in five nonoverlapping 400-µm2 areas. Total muscle fiber number was calculated by multiplying mean fiber density times overall area of the muscle. Analyses were performed with a light microscope connected to a computerized image analysis system (NIH image 1.61).

Myosin heavy chain (MHC) distribution
MHC extraction and SDS-PAGE were performed according to Ref. 24 . Silver-stained gels were then scanned and analyzed for MHC distribution (GS-700; Bio-Rad, Hercules, CA).

Protein isolation, firefly luciferase, and ß-galactosidase assays
Proteins isolation and measurement of firefly luciferase activity were performed as described previously (23). To correct for interindividual variations in transfection efficiency, luciferase activity was normalized to ß-galactosidase activity (23).

mRNA isolation and reverse transcription reaction
Muscle samples in RNALater (Qiagen) were disrupted in lysis buffer (Qiagen) (5 µl/mg of muscle). mRNA was isolated from 10 mg of lysed tissue in 50 µl using MagNA Pure LC mRNA isolation kit II in a MagNA Pure LC instrument (Roche Applied Science, Basel, Switzerland). RT was carried out with 3 µl of mRNA solution in 10 µl using the Reverse Transcriptase Core Kit (Eurogentec, Seraing, Belgium) with 50 µM oligo(dT)15 primer and RNase inhibitor (2 IU).

Primer design
Primer design, optimization, and specificity checking were performed as described previously (25). Acidic ribosomal phosphoprotein P0 (ARBP), forward, 5'-CCTGCACACTCGCTTCCTAGAG-3', reverse, 5'-CAACAGTCGGGTAGCCAATCTG-3' (GenBank NM_022402); ß-actin (ACTB), forward, 5'-TCAGGTCATCACTATCGGCAATG-3', reverse, 5'-TTTCATGGATGCCACAGGATTC-3' (GenBank NM_031144); Caveolin-3, forward, 5'-TGGTGAACAGAGACCCCAAGAAC-3', reverse, 5'-CACGCCATCGAAGCTGTAAGTG-3' (GenBank NM_013124); cyclophylin A (CycA), forward, 5'-TATCTGCACTGCCAAGACTGAGTG-3', reverse, 5'-CTTCTTGCTGGTCTTGCCATTCC-3' (GenBank NM_017101); glyceraldehyde-3-phosphate dehydrogenase (GAPDH), forward, 5'-CCAATGTATCCGTTGTGGATCTGAC-3', reverse, 5'-GCTTCACCACCTTCTTGATGTCATC-3' (GenBank NM_017008); hypoxanthine guanine phosphoribosyl transferase (HPRT), forward, 5'-CTCATGGACTGATTATGGACAGGAC-3', reverse, 5'-GCAGGTCAGCAAAGAACTTATAGCC-3' (GenBank NM_012583); MHC IIb and MHC IIx primers were as described previously (26); myostatin, forward, 5'-GGGCATGATCTTGCTGTAACCTTC-3', reverse, 5'-CGTGGAGTGTTCATCACAGTCAAG-3' (GenBank NM_019151); recombinant myostatin, forward, 5'-CGCTGTGGGTGCTCATGAGA-3' (GenBank NM_010834), reverse, 5'-CAACAGATGGCTGGCAACTAGAA-3' (pcDNA3.1-Zeo polyadenylation signal); peroxisome proliferator activated receptor {gamma} coactivator-1{alpha} (PGC-1{alpha}), forward, 5'-ACGCAGGTCGAATGAAACTGAC-3', reverse, 5'-TGGTGGAAGCAGGGTCAAAATC-3' (GenBank NM_031347); Tata box binding protein (TBP), forward, 5'-GCCACGAACAACTGCGTTGAT-3', reverse, 5'-AGCCCAGCTTCTGCACAACTCTA-3' (GenBank XM_217785); troponin I fast isoform, forward, 5'-GGACCTGAGAGCCAACCTGAA-3', reverse, 5'-TCCTCCAGTCACCCACGTCA-3' (GenBank NM_017185).

Real-time quantitative PCR
PCR was carried with LC Fast Start DNA Master SYBR Green kit (Roche Applied Science) from 0.5 µl of cDNA in a 20 µl final volume [4 mM MgCl2, 0.4 µM of each primer (except ARBP, ACTB, GAPDH, and myostatin 0.5 µM)]. PCR was performed using a Lightcycler (Roche Applied Science) for 45 cycles at 95 C for 20 sec; 50 C (MHC IIb), 54 C (ACTB), 55 C (PGC-1{alpha}, troponin I fast isoform), 57 C (ARBP, recombinant myostatin, caveolin-3, GAPDH), 58 C (CycA, TBP), 59 C (HPRT), 60 C (MHC IIx), and 61 C (myostatin) for 5 sec; and a final step of 10 sec at 72 C. Crossing point values were calculated from Lightcycler Software version 3.5 (Roche Applied Science) using the second derivative maximum method. Quantification was achieved using a pool of all cDNA samples as calibrator according to the comparative threshold cycle method (27), using the geometric average of three internal control genes (CycA, ARBP, TBP). Expression stability of six potential reference genes (ACTB, ARBP, CycA, GAPDH, HPRT, and TBP) was initially assessed using GENORM software (28). Gene-stability ranking from less stable genes to most stable genes was ACTB, GAPDH, HPRT, ARBP, and CycA/TBP (Fig. 1AGo). Vn/n + 1 is the pairwise variation analysis between the GENORM normalization factors from the n and n + 1 most stable genes (28). Pairwise variations of ARBP, CycA, and TBP were less than 0.15 (Fig. 1BGo), the threshold below which the inclusion of an additional control gene is not required. Therefore, ARBP, CycA, and TBP were used for normalization.


Figure 1
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  		FIG. 1. Determination of the optimal number of control genes for normalization. A, Gene-stability ranking using gene stability measure M. M is defined as the average pairwise variation of a particular gene with all other control genes. Genes with the lowest M values have the most stable expression. Stability of six reference genes, ARBP, ACTB, CycA, GAPDH, HPRT, and TBP, was analyzed. B, Determination of the number of control genes required for normalization. Vn/n + 1 is the pairwise variation analysis between the normalization factors from the n and n + 1 most stable genes. Normalization values were obtained using GENORM Software (28 ). A V value of 0.15 is the threshold, below which the inclusion of an additional control gene is not required (28 ). A V2/3 value of 0.141 was obtained for CycA + TBP + ARBP, which therefore were used for normalization.

 
Protein isolation and immunoblot analyses
MyoD and myogenin immunoblottings.
Muscle samples were homogenized in 15 vol of buffer consisting of 1% NP-40, 0.5% sodium deoxycholate, 0.1% SDS, 1 mM phenylmethylsulfonyl fluoride, 5 µg/ml aprotinin, 5 µg/ml leupeptin, 5 µg/ml pepstatin, and 20 mM Tris-HCl (pH = 6.8), or 4% SDS, 0.1 mM phenylmethylsulfonyl fluoride, and 1 µl/ml protease inhibitor cocktail set III (Calbiochem, San Diego, CA), respectively. Homogenates were centrifuged at 12,500 x g for 15 min at 4 C. Proteins (50 and 75 µg for MyoD and myogenin, respectively) were separated on SDS-PAGE and transferred to nitrocellulose membranes. Membranes were incubated overnight with a rabbit monoclonal antibody to MyoD (1/500 vol/vol; BD PharMingen, San Diego, CA) or a mouse monoclonal antibody to myogenin (1/500 vol/vol; Santa Cruz Biotechnology, Santa Cruz, CA). Incubation with horseradish peroxidase-conjugated goat antirabbit antibody (1/2000 vol/vol; Santa Cruz Biotechnology) and goat antimouse antibody (1/1200 vol/vol; Santa Cruz Biotechnology) was then performed for chemiluminescence detection (ECL; Amersham, Piscataway, NJ). Signal quantification was determined by densitometry (GS 700; Bio-Rad).

Desmin immunoblotting.
Proteins were extracted as previously described (29). Proteins (15 µg/lane) were separated on SDS-PAGE and transferred to nitrocellulose membranes. Membranes were incubated overnight with a mouse monoclonal antibody overnight at 4 C (1/500 vol/vol; Dako, Carpinteria, CA). Rabbit antimouse IgG (1/2000 vol/vol; Dako) conjugated to horseradish peroxidase was used for chemiluminescent detection of proteins (ECL; Amersham). The films were scanned and quantified using NIH image 1.61.

Statistics
Data are presented as means ± SE (n = 6–8 per group). Statistical comparisons were performed using paired t test and one-way and two-way ANOVA as applicable using Super ANOVA 1.11 software (Abacus Concepts, Berkeley, CA). The 0.05 level of confidence was accepted for statistical significance.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
In vivo model of myostatin-induced skeletal muscle atrophy
An in vivo model of myostatin-induced muscle mass atrophy was first generated. Whole TA muscles were electrotransfected by an im injection of a plasmid DNA encoding the murine myostatin cDNA under the control of cytomegalovirus promoter. Contralateral TA muscles were injected with the corresponding empty vector. Myostatin mRNA level was strongly increased in response to myostatin gene electrotransfer (Fig. 2AGo). When primers were designed to specifically amplify myostatin mRNA encoded by the vector, RT-PCR analysis showed that ectopic myostatin mRNA was not detected in the TA muscles electrotransfected with the empty vector, whereas ectopic myostatin mRNA was abundantly present in muscles electrotransfected with the myostatin expression plasmid (Fig. 2BGo). Importantly, ectopic myostatin mRNA level was decreased by about 4-fold 14 d after gene electrotransfer when compared with d-7 values.


Figure 2
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  		FIG. 2. Expression of myostatin is induced in response to myostatin gene electrotransfer. TA muscles electrotransfected with pcDNA-myostatin or the corresponding empty vector were removed 7 and 14 d after gene electrotransfer. A, Myostatin mRNA level. RT-PCR analysis was performed with primers that both recognize the endogenous and exogenous myostatin. B, Ectopic myostatin mRNA level. RT-PCR analysis was performed with primers specifically designed to amplify myostatin mRNA encoded by the expression vector. Note that it is not possible to subtract exogenous myostatin from total myostatin to obtain endogenous myostatin, as this calculation would assume equal RNA extraction efficiencies and RT efficiencies for both endogenous and exogenous myostatin. Results are expressed as means ± SE (n = 8). n.d., Nondetectable.

 
Morphometric and histomorphometric characteristics
Seven and 14 d after myostatin gene electrotransfer, TA muscle weight was decreased by about 10 (P < 0.01) and 20% (P < 0.01), respectively, when compared with control TA muscles (Table 1Go). Accordingly, whole TA muscle section was also significantly reduced in myostatin-electrotransfected TA muscles (Fig. 3AGo and Table 1Go). This effect appeared to be limited to the TA muscle, as extensor digitorum longus muscle weight remained unchanged (data not shown). Decrease in muscle mass was the result of a decrease in fiber cross-sectional area (Fig. 3Go, B and C), but did not result from a decrease in myofiber number (Table 1Go). Importantly, the reduction in muscle fiber cross-sectional area was consistently observed throughout the muscle cross-section and did not depend on muscle fiber localization. Analysis of fiber size distribution showed that the proportion of smaller fibers was greater in myostatin-electrotransfected muscles than in contralateral muscles 7 d after myostatin gene electrotransfer (P < 0.01) (Fig. 3CGo). This change in fiber cross-sectional area was even more dramatic at 14 d. At this time, the mean cross-sectional area of 1800 fibers from six animals was reduced by 28% (P < 0.001). In agreement with these data, total muscle protein content was significantly decreased in response to myostatin gene electrotransfer (Table 1Go).


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  		TABLE 1. Muscle mass, muscle protein content, and histomorphometric characteristics

 

Figure 3
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  		FIG. 3. Histomorphometric analyses. Muscles electrotransfected with pcDNA-myostatin or the corresponding empty vector were removed 7 and 14 d after gene electrotransfer. A, Hemalun-eosin-safran staining of whole muscle cross-sectional area of control (left)- and myostatin-electrotransfected (right) muscles 14 d after electrotransfer (bars equal 1 mm). B, Magnified view of representative muscle sections 14 d after gene electrotransfer (bars equal 120 µm). C, Frequency histograms of fiber cross-sectional area 7 (upper) and 14 (lower) d after gene electrotransfer. Results are expressed as means ± SE (n = 8). **, P < 0.01, and ***, P < 0.001 relative to corresponding contralateral muscles. {dagger}, P < 0.05, and {dagger}{dagger}, P < 0.01 relative to corresponding d-7 values.

 
Myostatin gene electrotransfer down-regulates the expression of muscle-specific genes
The significant decrease in muscle protein content suggests that myostatin exerts a negative control on muscle-specific protein expression. Therefore, we analyzed the expression of several muscle-specific genes encoding structural proteins. mRNA level of MHC IIb, the main MHC isoform in TA muscle, was significantly decreased by about 2-fold 7 and 14 d after myostatin gene electrotransfer (Fig. 4AGo), whereas MHC IIx mRNA level was nonsignificantly decreased. Importantly, MHC isoform distribution remained unchanged (Fig. 4BGo), illustrating that the reduction in MHC IIb isoform expression by myostatin does not induce fiber type transition.


Figure 4
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  		FIG. 4. Ectopic expression of myostatin down-regulates the expression of muscle-specific protein. A, RT-PCR analysis of MHC IIb and MHC IIx mRNA levels. mRNA level is expressed as the percentage of contralateral muscle values. B, Relative distribution of MHC isoforms. SDS-PAGE profile (upper). The plantaris muscle is shown to indicate the respective position of all the different mature MHC isoforms. The regenerating soleus muscle is shown to indicate the position of embryonic (Emb) and neonatal (Neo) MHC isoforms. Note that these isoforms were absent in electrotransfected muscles. Percentage of MHC isoforms relative to all MHC isoforms (lower). C, Transactivation of troponin I promoter and troponin I mRNA level. A plasmid containing a 2.3-kb fragment of fast troponin I promoter (pTnI-Luc) linked to a luciferase reporter gene was electrotransfected together with an expression plasmid containing the myostatin cDNA or the corresponding empty vector. Luciferase activity was measured 7 d later and corrected for transfection efficiency. mRNA level is expressed as the percentage of contralateral muscle values. D, Desmin protein level. Also shown are representative immunoblots in the different conditions. Results are expressed as means ± SE (n = 6–8). *, P < 0.05, **, P < 0.01, and ***, P < 0.001 relative to corresponding contralateral muscles.

 
A reporter plasmid of fast troponin I promoter was cotransfected either with the myostatin expression vector or the corresponding empty vector. Fast troponin I promoter transactivation was significantly reduced by about 2-fold 7 d after myostatin gene electrotransfer (Fig. 4CGo). Surprisingly, a corresponding decrease in troponin I mRNA level was not observed (Fig. 4CGo), suggesting a possible increase in troponin I mRNA stability 7 d after myostatin electrotransfer. However, troponin I mRNA level was decreased 14 d after myostatin electrotransfer (Fig. 4CGo). Finally, desmin protein level was also significantly reduced by about 50% both 7 and 14 d after myostatin gene electrotransfer (Fig. 4DGo). Overall, these data indicate that myostatin-induced muscle mass loss elicits the down-regulation of muscle-specific gene expression (MHC IIb, troponin I, desmin) without affecting MHC isoform distribution.

Myostatin gene electrotransfer down-regulates the expression of myogenic genes (MyoD and myogenin), caveolin-3, and PGC-1{alpha}
To get further insights into the mechanisms involved in the down-regulation of muscle-specific protein expression, we next determined the expression of muscle transcription factors, MyoD and myogenin. A promoter reporter plasmid of the myogenic transcription factor MyoD was cotransfected either with the myostatin expression vector or the corresponding empty vector. Muscles were removed 7 d after gene electrotransfer. As shown in Fig. 5AGo, MyoD promoter transactivation was reduced significantly by about 30% in response to myostatin gene electrotransfer. Consistent with these data, MyoD protein level also was decreased significantly 7 d after myostatin gene electrotransfer (Fig. 5BGo). Similarly, myogenin protein level also was reduced significantly (Fig. 5CGo). However, both MyoD and myogenin protein level returned to control values 14 d after gene electrotransfer.


Figure 5
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  		FIG. 5. Ectopic expression of myostatin down-regulates the expression of MyoD and myogenin. A, Transactivation of MyoD promoter. A plasmid containing the proximal promoter and the transcription start site of MyoD gene (pMyoD-Luc) linked to a luciferase reporter gene was electrotransfected together with an expression plasmid containing the myostatin cDNA or the corresponding empty vector. Luciferase activity was measured 7 d later and corrected for transfection efficiency. B and C, Immunoblot analysis of MyoD and myogenin protein levels. Also shown are representative immunoblots in the different conditions. Muscles electrotransfected with pcDNA-myostatin or the corresponding empty vector were removed 7 and 14 d after gene electrotransfer. Results are expressed as means ± SE (n = 6–8). *, P < 0.05 relative to corresponding contralateral muscles.

 
To deepen the mechanisms involved in myostatin-induced muscle atrophy, mRNA level of caveolin-3 and PGC-1{alpha} was determined. It was recently reported that caveolin-3, the muscle-specific isoform of caveolins, inhibited the activation of myostatin type I receptor and subsequent downstream signaling events (30). In the present study, caveolin-3 mRNA level was significantly reduced 14 d after myostatin gene electrotransfer (Fig. 6Go). Furthermore, PGC-1{alpha}, which directly regulates caveolin-3 promoter transactivation (31) and prevents skeletal muscle atrophy (32), was also significantly reduced 14 d after myostatin gene electrotransfer (Fig. 6Go).


Figure 6
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  		FIG. 6. Ectopic expression of myostatin decreases caveolin-3 and PGC-1{alpha} mRNA level. mRNA level is expressed as the percentage of contralateral muscle values. Results are expressed as means ± SE (n = 6–8). *, P < 0.05, and **, P < 0.01 relative to corresponding contralateral muscles.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
All in vivo experimental models used to explore the role of myostatin have been obtained from cattle with naturally occurring mutations in myostatin gene (3, 5) and from transgenic mice displaying disruption of myostatin gene (4, 33, 34). Production of transgenic mice overexpressing myostatin (35) or the myostatin inhibitory propeptide (36) have been also developed. In these animal models, the effects of myostatin on muscle mass thus critically depend on the action of myostatin on myoblasts and satellite cells during myogenesis and early postnatal growth. In an attempt to identify proper effects of myostatin on skeletal muscle fibers, we took advantage of the gene electrotransfer technique, which allows acute and temporal expression of a gene of interest (22, 23, 37). With this technique, the number of transfected muscle fibers is a critical parameter that determines the efficiency of the experimental model. It was recently reported that gene electrotransfer of an IGF-1 expression plasmid in 35% of muscle fibers was sufficient to attenuate skeletal muscle atrophy induced by glucocorticoid (38). We previously showed that the average percentage of electrotransfected fibers with a ß-galactosidase expression plasmid reached 30% in TA muscle (22, 23), which, therefore, must be high enough to induce skeletal muscle adaptation. Accordingly, gene electrotransfer of a myostatin expression vector, which resulted in a dramatic and specific increase in ectopic myostatin mRNA, induced a marked muscle atrophy. Another important question is whether or not myostatin exerts its action on nontransfected muscle fibers. In contrast to the histological feature of transgene expression, which is limited to clusters of muscle fibers (22, 23), reduction in muscle fiber size was consistently observed throughout the muscle cross-section and did not depend on muscle fiber localization (data not shown). These observations, together with the fact that myostatin functions in an autocrine/paracrine manner (10, 39), suggest that myostatin also acts on nontransfected muscle fibers. Finally, another important issue of the experimental procedure is the observation that nonoptimized gene electrotransfer protocols have been reported previously to result in muscle damage and the subsequent activation of satellite cells for muscle regeneration (22, 40, 41, 42, 43, 44). This might thus skew data analysis and interpretation. In the present study, characteristics of electric pulses were optimized (22), and we were unable to detect the expression of embryonic and neonatal MHC isoforms (markers of ongoing regeneration process), indicating that the present protocol was safe for skeletal muscle.

Our data clearly identifies myostatin as a powerful regulator of skeletal muscle mass in adulthood. Myostatin overexpression decreased muscle mass, fiber cross-sectional area, and protein content both 7 and 14 d after gene electrotransfer. Importantly, muscle atrophy was even more pronounced 14 d after myostatin gene electrotransfer when ectopic myostatin mRNA was decreased by about 4-fold when compared with d-7 values. This suggests that myostatin expression was high enough to decrease muscle mass and/or that other mechanisms occur to prolong myostatin action when its ectopic expression has decreased. Overall, our data clearly show that the present protocol could be used as an in vivo model to investigate the mechanisms involved in myostatin-induced muscle atrophy.

Some hypotheses can be raised to explain at the molecular level the effects of myostatin on TA muscle, including an increase in proteolysis, a reduction in protein synthesis, as well as a reduction in muscle gene expression. In the present study, we particularly investigated this latter possibility. As the contractile apparatus occupies more than 80% of the fiber volume (45), any decrease in myofibrillar gene expression may have major incidence on the regulation of skeletal muscle mass. Our current analyses of promoter transactivation and immunoblots clearly indicate that myostatin gene electrotransfer down-regulates the expression of muscle structural proteins (MHC IIb, troponin I fast isoform, and desmin). Our data are in agreement with recent observations showing that myostatin decreases muscle gene expression during in vitro differentiation of C2C12 myoblasts (46). Together, these data show that myostatin has the capability to induce muscle mass loss in adulthood by down-regulating the expression of muscle-specific genes.

The effect of myostatin on MHC isoform mRNA level was mainly restricted to MHC IIb. It was previously shown that myostatin was preferentially expressed in type IIb muscle fibers (47). However, because myostatin was expressed from a cytomegalovirus promoter, one should expect that myostatin expression was irrespective of muscle fiber type. The differential effect of myostatin on MHC expression could be linked to the greater relative distribution of MHC IIb isoform in TA muscle. A more favorable molecular context for the action of myostatin in type IIb muscle fibers, as well as a greater myostatin receptor density at the surface of type IIb muscle fibers, could also explain the more prominent action of myostatin on MHC IIb isoform. Such suggestions have to be experimentally tested. Importantly, whereas MHC IIb mRNA level was significantly decreased, the relative distribution of MHC isoforms remained unchanged, indicating that myostatin overexpression does not alter muscle fiber type. Our molecular analyses clearly demonstrate that myostatin markedly down-regulates the expression of myogenic genes (MyoD and myogenin) 7 d after gene electrotransfer. This probably contributes to the decrease in the expression of MHC IIb, troponin I, and desmin, because expression of these genes has been shown to be responsive to MyoD and/or myogenin (48, 49, 50). Overall, our data are in agreement with previous reports showing that myostatin decreases myogenic gene expression during in vitro differentiation of myoblasts (7, 8, 10, 46). Thus, one recurrent feature of the present study appears to be that some of the mechanisms triggered by myostatin during in vitro myogenesis are transposable to adult mature muscle fibers.

Muscle gene expression was still repressed 14 d after gene electrotransfer when ectopic expression of myostatin was largely blunted (4-fold reduction when compared with d 7 value) and expression of muscle regulatory factors had returned to control level. It was recently reported that caveolin-3, the muscle-specific isoform of caveolins the gene mutations of which are involved in the pathogenesis of limb-girdle muscular dystrophy 1C (51), inhibits myostatin signaling by suppressing myostatin receptor activation and downstream signaling (30). In the present study, the down-regulation of caveolin-3 expression 14 d after gene electrotransfer may contribute to relieve the inhibition of myostatin signaling by caveolin-3 and prolong the effects of myostatin, despite the decrease in myostatin ectopic expression. This is further strengthened by the observation that expression of PGC-1{alpha}, which regulates caveolin-3 promoter transactivation (31), was also significantly reduced 14 d after myostatin gene electrotransfer. Importantly, PGC-1{alpha} has also been recently reported as a protecting factor against muscle mass by down-regulating the expression of atrogenes involved in muscle proteolysis (32). The possibility that an increase in proteolysis is involved in myostatin-induced muscle atrophy is currently under investigation.

In conclusion, we have demonstrated that in vivo myostatin overexpression by gene electrotransfer induces severe atrophy in adult skeletal muscle. Our results also show that the down-regulation of muscle-specific gene expression plays a critical role in muscle mass loss. Given that myostatin appears to be a critical factor regulating muscle mass in adulthood, our data may be transposable in most physiological and physiopathological situations in which myostatin expression is induced.


   Acknowledgments
 
We are grateful to Vincent Dumas for histomorphometric analysis. We thank R. Rios (Departamento de Fisologia, Universidad de Santiago, Santiago de Compostela, Spain), H. Patel (Department of Biochemistry, McMaster University, Ontario, Canada), and S. Konieczny (Department of Biological Sciences, Purdue University, West Lafayette, IN) for the kind donation of pcDNA-Myostatin, pMyoD-Luc, and pTnI-Luc, respectively.


   Footnotes
 
A.A. is supported by an Allocation Doctorale de Recherche de la Région Rhônes Alpes. This work was supported by the Association Française contre les Myopathies Grant 2005.0214 (to D.F.).

Disclosure Statement: The authors have nothing to disclose.

First Published Online March 29, 2007

Abbreviations: ACTB, ß-Actin; ARBP, acidic ribosomal phosphoprotein P0; CycA, cyclophylin A; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; HPRT, hypoxanthine guanine phosphoribosyl transferase; MHC, myosin heavy chain; PGC-1{alpha}, peroxisome proliferator activated receptor {gamma} coactivator-1{alpha}; TA, tibialis anterior; TBP, Tata box binding protein.

Received November 9, 2006.

Accepted for publication March 16, 2007.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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