ABSTRACT
Skeletal muscle atrophy is a debilitating process that is associated with a wide variety of conditions including inactivity, disease and aging. Here, we demonstrate that the actin, myocardin-related transcription factors and serum response factor (actin–Mrtf–Srf) pathway is specifically downregulated in the muscle atrophy that is induced through disuse in mice. We show in vivo that the abolition of mechanical signals leads to the rapid accumulation of G-actin in myonuclei and the export of the Srf coactivator Mrtf-A, resulting in a decrease of Mrtf–Srf-dependent transcription that contributes to atrophy. We demonstrate that inhibition of the actin–Mrtf–Srf axis through overexpression of nuclear non-polymerizable actin, through pharmacological inhibition of Mrtf–Srf and through muscle-specific Srf deletion worsens denervation-induced atrophy. Conversely, maintenance of high levels of activity of Srf or Mrtfs in denervated muscle, through overexpression of constitutively active derivatives, counteracts atrophy. Altogether, our data provide new mechanistic insights into the control of muscle mass upon disuse atrophy by the actin–Mrtf–Srf pathway, highlighting Srf as a key mediator of mechanotransduction in muscle.
INTRODUCTION
Skeletal muscle wasting represents a major health problem because it is the consequence of a wide variety of pathological conditions, including inactivity (bed rest or nerve injury), starvation, chronic diseases and neuromuscular disorders.
Muscle atrophy always entails loss of organelles and cytoplasm, and involves protein-breakdown pathways (Bonaldo and Sandri, 2013). In particular, the ubiquitin-proteasome system induces the degradation of sarcomeric proteins, such as α-actin, through the muscle E3 ubiquitin ligase MuRF1 (Polge et al., 2011). In addition, disuse atrophy is associated with a decrease in the expression of genes encoding contractile proteins (Giger et al., 2009). This suggests that the control of muscle mass also stems from transcriptional and pre-translational regulatory processes. To date, it is unknown whether specific signaling pathways are involved in disuse muscle atrophy.
Serum response factor (Srf) is a member of the MADS-box transcription factor family. One major class of Srf targets is muscle-specific and comprises several genes encoding sarcomeric proteins (α-actins, myosins) (Pipes et al., 2006). The ability of Srf to regulate downstream target genes depends on its association with coactivators – including Ets-domain proteins, which are activated by MAPK, and myocardin-related transcription factors (Mrtfs), the nuclear accumulation of which is controlled by Rho GTPases and actin dynamics (Posern and Treisman, 2006). Using a mouse model of adult myofiber Srf deletion, we have previously reported that Srf is crucial for adult skeletal muscle hypertrophy, whereas it is not needed for myofiber maintenance (Guerci et al., 2012).
Here, we investigate the role of the actin–Mrtf–Srf pathway in skeletal muscle atrophy of different origins. We show that Srf activity is specifically downregulated by the abolition of mechanical cues and is governed by a rapid G-actin accumulation in myonuclei, which results in a decrease in intranuclear Mrtfs. Moreover, we demonstrate in vivo that denervation-induced muscle atrophy is aggravated by inhibiting the actin–Mrtf–Srf pathway and is counteracted by maintaining a high level of activity of either Srf or Mrtfs. Altogether, our results reveal the actin–Mrtf–Srf axis as a key pathway controlling disuse atrophy because its alteration directly impacts the extent of atrophy.
RESULTS AND DISCUSSION
Srf activity decreases upon disuse atrophy
To examine the contribution of Srf to skeletal muscle atrophy, we electroporated a Srf luciferase reporter construct into the tibialis anterior muscle of control mice, which showed a significant decrease in Srf transcription activity following denervation (Fig. 1A). As a consequence, the expression of Srf target genes, such as α-skeletal actin (Acta1) and Srf, which is its own target, was reduced in denervated gastrocnemius and plantaris, and tibialis anterior muscles (Fig. 1B; supplementary material Fig. S1A). Tallying with the diminished expression of Srf transcripts, a decreased Srf protein content was detected (Fig. 1C; supplementary material Fig. S1B). We then examined Srf subcellular localization within myofibers, as its extranuclear localization has been reported in atrophied rat muscles after 14 days of an ‘intensive care unit model’ (Ochala et al., 2011) and in beating-arrested neonatal rat cardiomyocytes (Lange et al., 2005); Srf extranuclear localization has also been associated with nuclear accumulation of MuRF2. However, up to 7 days post denervation, Srf protein was located inside myonuclei (supplementary material Fig. S1C), suggesting that, in this system and at this timing, the observed decrease in Srf activity is not a consequence of the relocalization of Srf protein through the activity of MuRF2.
Srf activity decreases upon disuse atrophy. (A) Responsiveness of an Srf reporter construct in tibialis anterior muscle from the contralateral leg or that of the denervated leg 7 days post denervation of control mice (n = 3). (B) Normalized expression of Srf and Acta1 in gastrocnemius and plantaris muscles from the contralateral leg or at 7 days post denervation (n = 4). (C) Representative western blot analysis for Srf with Ponceau staining. Normalized fold change in Srf protein level in gastrocnemius and plantaris muscles (GP) 7 days post denervation (n = 4). (D) Normalized expression of Srf and Acta1 in the gastrocnemius and plantaris muscles of mice submitted to 7 days of calorie restriction or ad libitum feeding (n = 4). (E) Representative western blot analyses for Srf with Ponceau staining. Normalized fold change in Srf protein level in gastrocnemius and plantaris muscles (GP) after 7 days of calorie restriction (n = 3). Quantitative data are means±s.e.m. *P<0.05, **P<0.001.
We next asked whether Srf activity is modulated in muscle atrophy that has been triggered by different cues. Denervation-induced atrophy is due to sciatic nerve severing, leading to the suppression of nerve impulses, muscle contraction and stretching. Atrophy induced by gastrocnemius tenotomy entails the reduction of mechanical signals and the prevention of stretching whilst maintaining muscle innervation. In line with the results obtained upon denervation, the expression of Acta1 and Srf was reduced in tenotomized gastrocnemius muscle (supplementary material Fig. S1D), suggesting that in vivo Srf activity is controlled through muscle stretching rather than through a trophic contribution of the nerve and excitation–contraction coupling.
Calorie-restriction-induced atrophy relies on nutrient shortage and not on the lack of mechanical cues. In contrast to observations following denervation and tenotomy, the expression of Srf targets was not downregulated after calorie restriction (Fig. 1D). Srf expression was even upregulated, but did not lead to increased Srf protein (Fig. 1E). Taken together, these data indicate that disuse, but not calorie-restriction-induced, muscle atrophy specifically leads to a sustained decrease in the overall activity of Srf.
Srf activity level regulates denervation-induced atrophy
To determine whether reduced Srf activity plays a functional role in denervation-induced atrophy, the extent of atrophy in denervated muscles from tamoxifen-injected HSA-Cre-ERT2∶Srfflox/flox mutant mice that lacked Srf in myofibers was analyzed (Fig. 2A; supplementary material Fig. S2A) (Guerci et al., 2012). After denervation, muscle weight and myofiber cross-sectional area (CSA) measurements revealed that mutant mice displayed a significantly higher atrophy of tibialis anterior, and gastrocnemius and plantaris muscles than those of control mice (Fig. 2B-D). This was not associated with higher expression levels of the ubiquitin ligases MAFbx, MuRF1 and Fbxo30 in mutant muscles, suggesting that their greater atrophy is not due to differences in the activation of the ubiquitin-proteasome pathway (supplementary material Fig. S2B). When subjected to calorie restriction, mutant and control mice presented similar extents of muscle atrophy (supplementary material Fig. S3), confirming the specific implication of Srf in disuse atrophy.
The level of Srf activity regulates denervation-induced atrophy. (A) Control and Srf-mutant mice were injected with tamoxifen (TMX) 14 days before denervation. Muscles were isolated 7 days post denervation. (B) Ratio of the weight of the tibialis anterior (TA), and gastrocnemius and plantaris (GP) muscles 7 days post denervation to those of the contralateral leg of control and mutant mice (n = 10). (C) Immunostaining of dystrophin in the contralateral and denervated leg tibialis anterior muscle of control and mutant mice 7 days post denervation. (D) Myofiber CSA to body weight ratio for tibialis anterior muscle in the denervated and contralateral legs in control and mutant mice 7 days post denervation (n = 4). AU, arbitrary units. (E) Tibialis anterior muscles (TA) from control mice were injected (or not) with AAV-SRFVP16 14 days before denervation. Muscles were isolated 7 days post denervation. (F) Ratio of the weight of the tibialis anterior muscle 7 days post denervation to that of the contralateral leg of mice that had been injected (or not) with AAV-SRFVP16 (n = 6). (G) Immunostaining of dystrophin in tibialis anterior muscle from the contralateral or denervated leg at 7 days post denervation that had been injected (or not) with AAV-SRFVP16. (H) Myofiber CSA to body weight ratio in tibialis anterior muscle from the contralateral or denervated leg at 7 days post denervation, which had been injected (or not) with AAV-SRFVP16 (n = 4). Quantitative data are means±s.e.m. *P<0.05, **P<0.001. Scale bars: 100 µm.
We then hypothesized that disuse atrophy could be counteracted by maintaining Srf activity through injection of control tibialis anterior muscles with adeno-associated viruses (AAVs) driving the expression of a constitutively active Srf derivative (AAV-SRFVP16) (Fig. 2E). Following denervation, overexpression of SRFVP16 induced a significant resistance to atrophy (Fig. 2F-H) and led to the persistence of endogenous Srf and Acta1 transcript levels (supplementary material Fig. S2C). Collectively, these results demonstrate in vivo that the level of Srf activity modulates the extent of denervation-induced muscle atrophy. It is worth noting that, in non-denervated muscles, the activation or blunting of Srf activity does not change myofiber size (Fig. 2D,H), despite its effect on the expression of target genes such as Acta1. This suggests that, during denervation, a decrease in the activity of Srf leads to an amplification of the transcriptional atrophic program, acting in synergy with other pathways that are involved in protein catabolism.
Disuse leads to nuclear G-actin accumulation and subsequent exclusion of Mrtfs from myonuclei
We next asked which signaling pathway could result in a decrease of Srf activity upon disuse. Because biophysical stimuli can alter actin polymerization and because Srf activity can be regulated by actin dynamics, we hypothesized that disuse could lead to alterations in monomeric G-actin quantity and/or localization. Strikingly, staining of G-actin, revealed by using fluorescent DNase I, showed that denervation induced a rapid and sustained accumulation of monomeric actin in myonuclei (Fig. 3A). By contrast, there was no difference in staining of G-actin between calorie-restricted and control muscles (Fig. 3A). Because Srf has been shown to control the expression of genes regulating actin treadmilling (Olson and Nordheim, 2010), the accumulation of nuclear G-actin could stem from a decrease of Srf activity upon disuse. Non-denervated Srf-deleted and AAV-SRFVP16-injected muscles showed the same G-actin staining as that in control muscles (supplementary material Fig. S4A), indicating that nuclear G-actin accumulation following denervation is a consequence of disuse and not of a reduction in Srf activity. Moreover, our data establish a link between the increase in nuclear G-actin levels and a physiopathologically relevant condition, muscle disuse atrophy.
Disuse leads to nuclear G-actin accumulation and subsequent exclusion of Mrtfs from myonuclei. (A) G-actin (DNase I), dystrophin (green) and DAPI staining in the tibialis anterior muscle of the contralateral leg, or that 14 h, 3 days (3D) or 7 days (7D) post denervation, or after 7 days of calorie restriction. (B) Intranuclear or shifted localization of overexpressed Mrtf-A–GFP in tibialis anterior muscle upon immunostaining of GFP. Nuclei are visualized through expression of H2B–mCherry and DAPI staining, and sarcolemma is identified through staining of dystrophin (gray). (C) Proportion of nuclei displaying intranuclear or shifted Mrtf-A–GFP localization in the tibialis anterior muscle of the contralateral leg, or at 14 h or 3 days (3D) post denervation, or after 3 days calorie restriction (n = 3–4). (D) Intranuclear or shifted localization of endogenous Mrtf-A (red), dystrophin (green) and DAPI in tibialis anterior muscle. (E) Proportion of nuclei displaying intranuclear or shifted endogenous Mrtf-A localization in the contralateral or 3 days post denervation tibialis anterior muscle (n = 4). (F) Proportion of nuclei displaying intranuclear or shifted Mrtf-A–GFP localization in tibialis anterior muscle that had been electroporated (or not) with R62DNLS-actin (n = 3). (G) Proportion of nuclei displaying intranuclear or shifted Mrtf-A–GFP localization in tibialis anterior muscle that had been electroporated (or not) with R62D-actin (n = 3). Quantitative data are means±s.e.m. *P<0.05 vs intranuclear Mrtf-A–GFP (–), §P<0.05 vs shifted Mrtf-A–GFP (–). Scale bars: 50 µm (A); 5 µm (B,D).
Actin dynamics control Srf activity by regulating the subcellular localization of Mrtfs – G-actin binds to Mrtfs, inhibiting their entry into (Pawlowski et al., 2010) and stimulating their export from nuclei (Vartiainen et al., 2007). To determine whether the increased levels of nuclear G-actin caused by denervation are correlated with the altered subcellular localization of Mrtfs, tibialis anterior muscles were electroporated with a plasmid encoding Mrtf-A tagged with green fluorescent protein (Mrtf-A–GFP), together with a Histone2B-mCherry plasmid to identify electroporated myofiber nuclei. Immunostaining for GFP revealed two different subcellular localizations of Mrtf-A – intranuclear, characterized by an exact colocalization of Mrtf-A–GFP with mCherry-tagged Histone 2B (H2B–mCherry), and shifted, in which some Mrtf-A–GFP did not colocalize with chromatin (Fig. 3B; supplementary material S4B,C and illustrated by 3D reconstructions in supplementary material Fig. S4D). In addition, immunostaining for Emerin suggested that at least some Mrtf-A–GFP is perinuclear and located outside of the nuclear envelope (supplementary material Fig. S4E,F). Importantly, denervation led to a rapid decrease in intranuclear Mrtf-A–GFP (Fig. 3C). In line with this, immunostaining for endogenous Mrtf-A confirmed that denervation diminished Mrtf-A localization in myonuclei (Fig. 3D,E). By contrast, there was no difference in the distribution of Mrtf-A–GFP in muscle from mice that had been fed in a normal or restricted manner (Fig. 3C). Taken together, these data show that Mrtf-A nuclear exclusion is a specific response to changes in mechanical load.
As post-denervation kinetics of nuclear G-actin accumulation coincide with that of decreasing intranuclear Mrtf-A, we speculated that an increase in nuclear G-actin could induce Mrtf-A shuttling out of myonuclei. To test this hypothesis, we artificially elevated the amount of nuclear G-actin by electroporating tibialis anterior muscle with a R62DNLS-actin plasmid, which encodes a constitutively nuclear non-polymerizable actin derivative (supplementary material Fig. S4G), together with the Mrtf-A–GFP plasmid. Overexpression of R62DNLS-actin in non-denervated muscle was sufficient to exclude Mrtf-A–GFP from myonuclei to a similar extent to that observed post denervation (Fig. 3F). In fibroblasts, it has been shown that the interaction between G-actin and Mrtfs can occur in both the nucleus and cytoplasm (Vartiainen et al., 2007). Interestingly, overexpression of R62D-actin, leading to an accumulation of G-actin predominantly in the cytoplasm (supplementary material Fig. S4H), did not change Mrtf-A–GFP localization, suggesting that, in muscle tissue, shuttling of Mrtfs, and subsequent activation of Srf–Mrtf, is principally modulated by nuclear G-actin and does not involve the regulation of cytosolic actin pathways (Fig. 3G). As G-actin shuttles actively between cytoplasm and nuclei through specific transporters, G-actin transport could be affected by loss of mechanical cues, resulting in its nuclear accumulation. Furthermore, actin polymerization and depolymerization processes also take place in nuclei and could modulate nuclear G-actin content (Grosse and Vartiainen, 2013). In particular, the nuclear actin regulatory protein Mical-2 has been shown to lower the amount of nuclear G-actin (Lundquist et al., 2014), and denervated muscles exhibited a reduction of Mical-2 expression (supplementary material Fig. S4I). We can thus speculate that abolition of mechanical signals increases nuclear G-actin by disrupting nuclear actin dynamics through the deregulation of actin regulatory proteins, such as Mical-2.
The actin–Mrtf–Srf pathway controls disuse atrophy
We next sought to ascertain the role of the actin–Mrtf–Srf pathway in regulating disuse atrophy. Because denervation-induced atrophy is aggravated in muscles that lack Srf, we hypothesized that atrophy should also be worsened through reducing the activity of Srf–Mrtf by using R62DNLS-actin. In line with the increased Mrtf-A–GFP nuclear exclusion that is induced through R62DNLS-actin (Fig. 3F), R62DNLS-actin-electroporated tibialis anterior muscle showed a higher denervation-induced atrophy than control muscles, mimicking the Srf-knockout phenotype (Fig. 4A). Mrtfs-dependent Srf activity can also be inhibited by using the compound CCG-1423 (Minami et al., 2012). Tibialis anterior, and gastrocnemius and plantaris muscles of CCG-1423-injected mice were more atrophied after denervation than the muscles of vehicle-injected mice, whereas calorie-restriction-induced atrophy was unaffected (Fig. 4B,C). These data show in vivo that specific downregulation of the activity of Mrtfs aggravates disuse muscle atrophy.
The actin–Mrtfs–Srf pathway controls disuse atrophy. (A) Myofiber CSA to body weight ratio for tibialis anterior muscle from the contralateral leg or that 7 days post denervation, electroporated (or not) with R62DNLS-actin (n = 3–17). (B) Ratio of the weight of the tibialis anterior (TA), and gastrocnemius and plantaris (GP) muscles 7 days post denervation to those of the contralateral leg of control mice that had been injected with CCG-1423 or vehicle, (n = 5). (C) Ratio of the weight of the tibialis anterior (TA), and gastrocnemius and plantaris (GP) muscles after 7 days of calorie restriction to those of control mice that had been injected with CCG-1423 or vehicle (n = 5). (D) Myofiber CSA to body weight ratio for tibialis anterior muscle 7 days post denervation or from the contralateral leg, electroporated (or not) with Mrtf-A-XXX or Mrtf-A–GFP (n = 4–17). (E) The ratio of the Srf-deleted myofiber CSA to body weight for tibialis anterior muscle 7 days post denervation or from the contralateral leg, electroporated (or not) with Mrtf-A-XXX (n = 3–4). Quantitative data are means±s.e.m. *P<0.05, **P<0.001. AU, arbitrary units.
To determine whether muscle atrophy could be rescued by maintaining high activity levels of Mrtfs, we overexpressed in myofibers a constitutively active Mrtf-A derivative (Mrtf-A-XXX), which is defective for actin binding and is located inside myonuclei exclusively (supplementary material Fig. S4J). Fibers that expressed Mrtf-A-XXX became completely resistant to denervation-induced atrophy (Fig. 4D), suggesting that, among Srf cofactors, Mrtfs are the main modulators of Srf activity in this system. Interestingly, Mrtf-A–GFP, which has wild-type actin-binding sites, did not rescue atrophy (Fig. 4D), strongly supporting a crucial role for the binding of G-actin to Mrtfs in the muscle atrophic response to disuse.
Mrtfs have been shown to interact with other transcription regulators, such as Smads (Charbonney et al., 2011; Wang et al., 2012). In order to investigate whether the atrophy resistance induced by active Mrtf-A requires Srf, Mrtf-A-XXX was electroporated into muscles that lacked Srf. Denervated mutant muscles displayed the same level of atrophy, regardless of Mrtf-A-XXX overexpression (Fig. 4E), demonstrating that the resistance to denervation-induced atrophy conferred by Mrtf-A-XXX depends on Srf.
In conclusion, our study shows that the alteration of any components of the actin–Mrtf–Srf pathway is sufficient to affect the extent of disuse muscle atrophy and identifies nuclear G-actin and Mrtfs as the main upstream regulators that are involved. Our data provide novel genetic and mechanistic insights into how muscle mass is regulated through a lack of mechanical stimulation and unravels the actin–Mrtf–Srf axis as a key specific regulator of disuse atrophy, establishing a link with human pathophysiological atrophy due to inactivity, nerve injury or motor neuron diseases. Therapeutic targeting of the actin–Mrtf–Srf pathway might improve the condition and quality of life of an individual with disuse muscle atrophy because maintaining the activity of Mrtfs fully counteracts the phenotype.
MATERIALS AND METHODS
Mouse protocols
Srfflox/flox and HSA-Cre-ERT2∶Srfflox/flox mice have been described previously (Lahoute et al., 2008). Two-month-old HSA-Cre-ERT2∶Srfflox/flox and Srfflox/flox males were given four tamoxifen injections (1 mg/day; Sigma) and are referred as mutant and control mice, respectively.
Denervation was performed by sectioning the sciatic nerve of one leg. The intact contralateral leg was used as an internal control. For calorie restriction, mice were placed in individual cages and fed with 50% of the daily food amount or fed ad libitum. Where indicated, mice were injected intraperitoneally and daily with CCG-1423 (0.15 mg/kg; Cayman Chemical) or vehicle. Treatment with CCG-1423 started 2 days prior to and then during the timeframe of the experiment.
Gene electrotransfer was performed as described previously (Guerci et al., 2012). Tibialis anterior muscles were injected with the indicated plasmids: 1 µg Mrtf-A–GFP+3 µg H2B–mCherry+5 µg empty vector; 10 µg Mrtf-A-XXX or R62D-actin or R62DNLS-actin+3 µg H2B–mCherry; 5 µg 3DA-luciferase+2 µg TK-Renilla+5 µg empty vector. Mice were denervated or calorie restricted 7 days after electroporation. Reporter assays were performed using the Dual-Luciferase kit (Promega).
AAV-SRFVP16 particles were produced as described previously (Guerci et al., 2012). AAV transduction was performed by injecting tibialis anterior muscles with 1.87×1010 viral genomes 14 days before denervation.
All experiments were conducted in accordance with the European guidelines for the care and use of laboratory animals and were approved by the institutional ethic committee (number 00315.01).
Plasmids
The Srf-reporter construct (3DA-luciferase: 3 CArGs upstream Luciferase gene) has been described previously (Geneste et al., 2002). The TK-Renilla plasmid (Promega) was used as an electroporation efficiency control. Plasmid H2B–mCherry encodes the Histone-2B–mCherry fusion protein (Abgene). Plasmids encoding hemagglutinin (HA)-tagged-Mrtf-A-XXX, Mrtf-A–GFP, HA-tagged-R62D-actin and Flag-tagged-R62DNLS-actin have been described previously (Guettler et al., 2008; Posern et al., 2002; Stern et al., 2009; Vartiainen et al., 2007).
RNA extraction and qRT-PCR
RNA extraction and quantitative real-time (qRT)-PCR analysis were performed as described previously (Guerci et al., 2012). Values were normalized using Hydroxymethylbilane synthase (Hmbs). The following primers were used: Acta1-F, 5′-CTGAGCGCAAGTACTCAGTGTGGA-3′; Acta1-R, 5′-TTCCAAAAACAGGCGCCGGCTGCA-3′; Srf-F, 5′-CACCTACCAGGTGTCGGAAT-3′; Srf-R, 5′-GCTGTGTGGATTGTGGAGGT-3′; Hmbs-F, 5′-TGCACGATCCTGAAACTCTG-3′; Hmbs-R, 5′-TGCATGCTATCTGAGCCATC-3′.
Immunostaining
Muscle cryosections (8 µm) were fixed in 4% paraformaldehyde for 8 min, blocked in 5% goat serum, 4% BSA, 0.2% Triton X-100 in PBS, incubated overnight with primary antibodies, washed with PBS, incubated with appropriate secondary antibodies and counterstained with DAPI. Tibialis anterior muscles overexpressing Mrtf-A–GFP were prefixed in 4% paraformaldehyde for 2 h and incubated overnight in 40% sucrose before freezing. The primary antibodies used were against Srf and Mrtf-A (Santa Cruz), dystrophin (Novocastra), laminin-α2 and GFP (Abcam), HA (Roche), Flag (Sigma) and emerin (Novus Biologicals). For G-actin staining, Texas-Red-conjugated DNase-I was used following the manufacturer’s protocol (Molecular Probes).
Image acquisition
Digital images were acquired using a Zeiss Axiovert 200M microscope with 10×, 20× or 40× magnification, or using a Spinning Disk Leica confocal microscope with a 100× oil-immersion objective, cooled CCD CoolSNAP-HQ2 camera (Photometrics) and Metamorph 7.7.5 (Molecular Devices). Images were composed and edited in Adobe Photoshop and ImageJ. Background was reduced using brightness and contrast adjustments applied to the whole image. Three-dimensional reconstitutions from confocal microscopy images were performed using Imaris 7.2 (Bitplane).
Morphometric analyses
Myofiber CSA was analyzed by using immunostaining of dystrophin, marking myofiber sarcolemma, and then using Metamorph. Between 1000 and 1500 myofibers were analyzed per non-electroporated or AAV-injected muscle. For electroporated tibialis anterior muscle, 40–200 myofibers (positive for HA-Mrtf-A-XXX, Flag-R62DNLS-actin or Mrtf-A–GFP) were analyzed.
Quantification of Mrtf-A–GFP subcellular distribution was performed on Mrtf-A–GFP-positive and H2B–mCherry-positive myonuclei (30–250 myonuclei/muscle). For endogenous Mrtf-A, 40–120 myonuclei/muscle were analyzed.
Western blotting analysis
Western blotting was performed as described previously (Lahoute et al., 2008). Immunoblots were hybridized with anti-Srf antibody and normalized by using Ponceau staining. Protein bands were quantified by using Bio-Rad Quantity One (version 4.6.6).
Statistical analysis
Statistics were performed by using SigmaPlot 12.5 (Systat). When two groups were compared, the significance of differences between the means was assessed with a Student's t-test after normality and equal variance tests were passed. For multiple comparison, a two-way ANOVA test was used after normality and equal variance tests were passed and was followed by using a Bonferroni t-test for pairwise multiple comparison. P-values of <0.05 were considered statistically significant.
Acknowledgments
We thank Richard Treisman (Cancer Research UK, London, UK) and Guido Posern (Institute for Physiological Chemistry, Halle, Germany) for providing materials. We are grateful to Pascal Maire (Cochin Institute, Paris, France), Luisa Dandolo (Cochin Institute, Paris, France), Anne Lombès (Cochin Institute, Paris, France) and Dominique Daegelen (Cochin Institute, Paris, France) for critical reading of the manuscript.
Footnotes
Competing interests
The authors declare no competing interests.
Author contributions
L.C. designed and carried out experiments, analyzed results and wrote the manuscript. G.H. and A.P. conducted experiments and analyzed results. V.R-H. and A.G. provided expertise in physiological analysis. A.S. designed and carried out experiments, analyzed results, wrote the manuscript and provided financial support.
Funding
This work was supported by The French Muscular Dystrophy Association (AFM); Agence Nationale de la Recherche (ANR) [grant JC08-327703; 13-BSV1-0005-01]; and Labex WhoAmI.
Supplementary material available online at http://jcs.biologists.org/lookup/suppl/doi:10.1242/jcs.155911/-/DC1
- Received May 2, 2014.
- Accepted October 15, 2014.
- © 2014. Published by The Company of Biologists Ltd
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