Title: DHPR α1S subunit controls skeletal muscle mass and morphogenesis
Abstract: Article24 December 2009free access DHPR α1S subunit controls skeletal muscle mass and morphogenesis France Piétri-Rouxel Corresponding Author France Piétri-Rouxel UMR 7215, CNRS, UMR S 974 Inserm, Institut de Myologie, Université Pierre et Marie Curie, Paris, France Search for more papers by this author Christel Gentil Christel Gentil UMR 7215, CNRS, UMR S 974 Inserm, Institut de Myologie, Université Pierre et Marie Curie, Paris, France Search for more papers by this author Stéphane Vassilopoulos Stéphane Vassilopoulos Department of Bioengineering and Therapeutic Sciences, School of Pharmacy, University of California, San Francisco, CA, USA Search for more papers by this author Dominique Baas Dominique Baas UMR 5239, CNRS, ENS, Université Lyon 1 46, allée d'Italie Lyon cedex 07, France Search for more papers by this author Etienne Mouisel Etienne Mouisel UMR 7215, CNRS, UMR S 974 Inserm, Institut de Myologie, Université Pierre et Marie Curie, Paris, France Search for more papers by this author Arnaud Ferry Arnaud Ferry UMR 7215, CNRS, UMR S 974 Inserm, Institut de Myologie, Université Pierre et Marie Curie, Paris, France Université Paris Descartes, Paris, France Search for more papers by this author Alban Vignaud Alban Vignaud UMR 7215, CNRS, UMR S 974 Inserm, Institut de Myologie, Université Pierre et Marie Curie, Paris, France Search for more papers by this author Christophe Hourdé Christophe Hourdé UMR 7215, CNRS, UMR S 974 Inserm, Institut de Myologie, Université Pierre et Marie Curie, Paris, France Search for more papers by this author Isabelle Marty Isabelle Marty Inserm U836, Grenoble Institut des Neurosciences, Université Joseph Fourier, Grenoble, France Search for more papers by this author Laurent Schaeffer Laurent Schaeffer UMR 5239, CNRS, ENS, Université Lyon 1 46, allée d'Italie Lyon cedex 07, France Search for more papers by this author Thomas Voit Thomas Voit UMR 7215, CNRS, UMR S 974 Inserm, Institut de Myologie, Université Pierre et Marie Curie, Paris, France Search for more papers by this author Luis Garcia Luis Garcia UMR 7215, CNRS, UMR S 974 Inserm, Institut de Myologie, Université Pierre et Marie Curie, Paris, France Search for more papers by this author France Piétri-Rouxel Corresponding Author France Piétri-Rouxel UMR 7215, CNRS, UMR S 974 Inserm, Institut de Myologie, Université Pierre et Marie Curie, Paris, France Search for more papers by this author Christel Gentil Christel Gentil UMR 7215, CNRS, UMR S 974 Inserm, Institut de Myologie, Université Pierre et Marie Curie, Paris, France Search for more papers by this author Stéphane Vassilopoulos Stéphane Vassilopoulos Department of Bioengineering and Therapeutic Sciences, School of Pharmacy, University of California, San Francisco, CA, USA Search for more papers by this author Dominique Baas Dominique Baas UMR 5239, CNRS, ENS, Université Lyon 1 46, allée d'Italie Lyon cedex 07, France Search for more papers by this author Etienne Mouisel Etienne Mouisel UMR 7215, CNRS, UMR S 974 Inserm, Institut de Myologie, Université Pierre et Marie Curie, Paris, France Search for more papers by this author Arnaud Ferry Arnaud Ferry UMR 7215, CNRS, UMR S 974 Inserm, Institut de Myologie, Université Pierre et Marie Curie, Paris, France Université Paris Descartes, Paris, France Search for more papers by this author Alban Vignaud Alban Vignaud UMR 7215, CNRS, UMR S 974 Inserm, Institut de Myologie, Université Pierre et Marie Curie, Paris, France Search for more papers by this author Christophe Hourdé Christophe Hourdé UMR 7215, CNRS, UMR S 974 Inserm, Institut de Myologie, Université Pierre et Marie Curie, Paris, France Search for more papers by this author Isabelle Marty Isabelle Marty Inserm U836, Grenoble Institut des Neurosciences, Université Joseph Fourier, Grenoble, France Search for more papers by this author Laurent Schaeffer Laurent Schaeffer UMR 5239, CNRS, ENS, Université Lyon 1 46, allée d'Italie Lyon cedex 07, France Search for more papers by this author Thomas Voit Thomas Voit UMR 7215, CNRS, UMR S 974 Inserm, Institut de Myologie, Université Pierre et Marie Curie, Paris, France Search for more papers by this author Luis Garcia Luis Garcia UMR 7215, CNRS, UMR S 974 Inserm, Institut de Myologie, Université Pierre et Marie Curie, Paris, France Search for more papers by this author Author Information France Piétri-Rouxel 1, Christel Gentil1, Stéphane Vassilopoulos2, Dominique Baas3, Etienne Mouisel1, Arnaud Ferry1,4, Alban Vignaud1, Christophe Hourdé1, Isabelle Marty5, Laurent Schaeffer3, Thomas Voit1 and Luis Garcia1 1UMR 7215, CNRS, UMR S 974 Inserm, Institut de Myologie, Université Pierre et Marie Curie, Paris, France 2Department of Bioengineering and Therapeutic Sciences, School of Pharmacy, University of California, San Francisco, CA, USA 3UMR 5239, CNRS, ENS, Université Lyon 1 46, allée d'Italie Lyon cedex 07, France 4Université Paris Descartes, Paris, France 5Inserm U836, Grenoble Institut des Neurosciences, Université Joseph Fourier, Grenoble, France *Corresponding author. UMR 7215, CNRS, UMR S 974 Inserm, Institut de Myologie, Université Pierre et marie Curie, 105 boulevard de l'Hôpital, Paris 75013, France. Tel.: +33 01 4077 9635; Fax: +33 01 5360 0802; E-mail: [email protected] The EMBO Journal (2010)29:643-654https://doi.org/10.1038/emboj.2009.366 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info The α1S subunit has a dual function in skeletal muscle: it forms the L-type Ca2+ channel in T-tubules and is the voltage sensor of excitation–contraction coupling at the level of triads. It has been proposed that L-type Ca2+ channels might also be voltage-gated sensors linked to transcriptional activity controlling differentiation. By using the U7-exon skipping strategy, we have achieved long-lasting downregulation of α1S in adult skeletal muscle. Treated muscles underwent massive atrophy while still displaying significant amounts of α1S in the tubular system and being not paralysed. This atrophy implicated the autophagy pathway, which was triggered by neuronal nitric oxide synthase redistribution, activation of FoxO3A, upregulation of autophagy-related genes and autophagosome formation. Subcellular investigations showed that this atrophy was correlated with the disappearance of a minor fraction of α1S located throughout the sarcolemma. Our results reveal for the first time that this sarcolemmal fraction could have a role in a signalling pathway determining muscle anabolic or catabolic state and might act as a molecular sensor of muscle activity. Introduction Skeletal muscle growth is dependent on innervation and exercise. Prolonged periods of muscle inactivity because of bed rest, hindlimb unloading, immobilization, microgravity or denervation result in a significant muscle atrophy (Borisov et al, 2001; Adams et al, 2003; Dedkov et al, 2003; De-Doncker et al, 2005). Direct electrical stimulations can, to a large extent, substitute for innervation and preserve or even restore almost normal properties in denervated muscle fibres (Kern et al, 2004; Squecco et al, 2008). Nevertheless, mechanisms implicated in muscle homeostasis under such circumstances are poorly understood. These molecular mechanisms are likely triggered by action potentials or prompted by the contractile function itself. In neurons, L-type Ca2+ channels have been shown to act as voltage-gated sensors linked to transcriptional activity controlling differentiation (Wheeler et al, 2008). In skeletal muscle, L-type Ca2+ channels are made of five subunits (α1S, α2, β, γ, δ). The α1S, known as the dihydropyridine receptor (DHPR), is a four-repeat transmembrane protein, which contains the basic functional elements of the channel, including the Ca2+ selective pore and the S4 'voltage-sensing' transmembrane segment (Ahern et al, 2001). It is predominantly located in T-tubules, although a minor fraction of DHPR has been found on the sarcolemma (Jorgensen et al, 1989). α1S is also located in triads where it interacts with the type 1 ryanodine sensitive Ca2+ release channel (RyR1) and forms the voltage sensor of the excitation–contraction (EC) coupling. We hypothesize that subtle modification of α1S contents in adult muscle could allow to decipher unexpected functions for this multi-functional protein. Indeed, complete knockout of the α1S subunit during development is lethal as observed in the mdg/mdg dysgenic mouse model (Pai, 1965; Banker, 1977; Chaudhari, 1992). To create a viable model, which would overcome this problem, we have chosen a strategy for long-lasting RNA interference based on the use of U7 snRNA chimaeras. The U7 system has been successfully used in several paradigms for rescuing mutated mRNAs by either prompting exon skipping or exon inclusion to restore protein synthesis (Goyenvalle et al, 2004; Marquis et al, 2007). Here, we used this system to decline amounts of α1S subunit by forced skipping of an exon affecting its mRNA reading frame. To guarantee long-lasting effects, the U7 snRNA chimaeras were delivered intramuscularly by using adenovirus-associated viral vectors (AAV). After 6 months, treated muscles showed a significant atrophy even though muscle fibres were not EC uncoupled and were still capable to contract when stimulated. We also show that decreasing α1S induced redistribution of the neuronal nitric oxide synthase (nNOS), activation of FoxO3A, increased expression of key autophagy-related genes and initiated autophagosome formation. Finally, confocal investigations allowed to identify which subcellular compartment displayed the decrease of α1S after exon skipping treatment. We found that atrophy and ultrastructural disorganization of muscle fibres correlated with α1S disappearance from the sarcolemma while α1S remained present in T-tubules and triads. All together, these results suggest that the minor sarcolemmal fraction of α1S has a role in a signalling pathway controlling muscle maintenance. At that level, α1S might act as a sensor of muscle activity in charge of controlling muscle mass and morphogenesis. Results Knockdown of the α1S subunit The α1S subunit is encoded by the Cacna1s gene, made of 46 exons and located, in mouse, on chromosome 1 (Drouet et al, 1993). Exon 16 encodes an amino acid sequence forming a cytoplasmic loop (II–III loop) connecting trans-membrane repeats II and III of the α1S subunit, which interacts directly with RyR1 and is crucial for skeletal muscle EC coupling (Tanabe et al, 1990; Casarotto et al, 2006; Cui et al, 2009). To knock down the α1S subunit we have designed U7 snRNA chimaeras interfering specifically with its pre-mRNA splicing. We have chosen to target exon splicing enhancers (ESE) and the acceptor splice site (SA) of exon 16 (U7-ESE, U7-SA). A U7-Ctrl, not affecting α1S splicing, was used as mock control (Figure 1A). Engineered U7 genes were cloned into AAV2 vector backbones, which were type 1 pseudotyped for intramuscular delivery: AAV2/1 (U7-ESE), AAV2/1 (U7-SA), AAV2/1 (U7-Ctrl). To improve skipping of exon 16, both AAV2/1 (U7-ESE) and AAV2/1 (U7-SA) vectors were co-injected into tibialis anterior muscles (TAΔDHPR). Contralateral muscles were used as controls (TACtrl) and received mock AAV2/1 (U7-Ctrl). To show that these genomes were still active and produced the interfering U7 transcripts at 6 months post injection, total RNA from TAΔDHPR muscles was extracted, reverse transcribed into DNA and PCR amplified using a forward primer in the antisense sequence (ESE) and a reverse primer in the U7 stem loop. Results confirmed that vector expression was persistent up to 6 months post injection (Supplementary Figure S1). Figure 1.Generation of a local knockdown of α1S subunit. (A) Schema of exonic chaining of pre-mRNA, exons 15, 16 and 17 are represented by blue boxes. Sequences of exon acceptor site SA (in blue) and exon splicing enhancer ESE (in black) of exon 16 were targeted by antisense sequences introduced in U7smOPT cassettes. (B) qRT–PCR analysis using primers overlapping exons 15 and 16 permitted to quantify the non-skipped α1S form: 18±6% (**P⩽0.001, n=19) and 10±7% (**P⩽0.001 n=19) (black bars) of the total α1S mRNA (grey bars) at 2 and 6 months post injection in TA of AAV1-(U7-ESE) and AAV1-(U7-SA):ΔDHPR or AAV1-(U7-Ctrl): c as a control, respectively. (C) Six months post injection, lysates from TAΔDHPR (ΔDHPR) and TACtrl (c) were immuno-blotted for α1S or α-actin for four mice. Graph depicts mean±s.e.m. of relative expression of α1S subunit determined by densitrometry and nomalized to the α-actin expression for each muscle. Results were expressed in protein levels of α1S subunit in TAΔDHPR normalized to TACtrl for each mice, **P⩽0.001, n=4. (D) Longitudinal cryo-sections from TAΔDHPR (ΔDHPR) and TACtrl (c) were stained with anti-α1S subunit (red) and anti-laminin (green) antibodies, nuclei were visualized by Dapi (blue) and imaged by confocal microscopy. Bars represent 20 μm. Download figure Download PowerPoint Skipping exon 16 would allow abortion of α1S synthesis by provoking a shift of its mRNA reading frame creating a premature stop codon in position 33 of exon 17 (Figure 1A). Skipping efficiency was assessed by real-time quantitative RT–PCR (qRT–PCR) on total mRNA (Supplementary Table S1). Such analysis showed that levels of the unskipped α1S represented 18±6% (P⩽0.001, n=19) and 10±7% (P⩽0.001 n=19) of the total α1S mRNAs at 2 and 6 months post injection, respectively (Figure 1B). Western blotting revealed a significant decrease of α1S. Six months post injection, α1S levels in TAΔDHPR were about 56±13% (P⩽0.001, n=5) compared with the contralateral TACtrl (Figure 1C). It is unlikely that skipped mRNAs could still be translated into a truncated protein. Indeed, such a truncated protein should display a theoretical molecular weight of about 80 kDa. Western blots using polyclonal antibodies made against the N-terminal protein never detected such a band (Supplementary Figure S2). Levels of α1S were also assessed by immuno-fluorescence staining on TAΔDHPR muscle sections. At 6 months, the majority of fibres showed a decrease in staining intensity (Figure 1D). Atrophy, fibrosis and reduced maximal force after α1S knockdown A significant atrophy was observed in injected TAΔDHPR. Six months after injection, TAΔDHPR showed a loss of 41±3% of muscle mass compared to contralateral TACtrl (P⩽0.001, n=8) (Figure 2A; Table 1). Morphometric analysis showed that there was no change in the number of fibres per muscle. Atrophy was due to a decrease of the size of muscle fibres (Table 1) as shown by the distribution of cross-section diameters, which was shifted towards lower values (Figure 2B). Average fibre diameter was 24±2 μm in injected TAΔDHPR compared to 43±4 μm in contralateral TACtrl (Table 1; Figure 2C). This atrophy was not accompanied by a modification of muscle fibre phenotype. Typically, the TA was almost entirely composed of fast-twitch fibres, expressing type 2b and 2x myosin heavy chains (MHC) isoforms and a very small number of muscle fibres expressing the MHC-2a isoform (Raffaello et al, 2006). No significant modifications were observed in the number of fibres expressing MHC-2x and -2b showing that the major muscle phenotype was maintained in treated muscle compared to contralateral (Table 1). However, there was a slight decrease in the number of fibres expressing MHC-2a in TAΔDHPR compared to TACtrl (Table 1). Still, TAΔDHPR muscles showed striking morphological abnormalities without evidence of necrosis or core formation. There was no sign of infiltration by scavenger cells, which is a hallmark of necrosis, nor positive immuno-labelling for neonatal and embryonic MHCs (data not shown), a more sensitive marker of regeneration given that this isoform is expressed during muscle development and particularly the formation of new fibres from satellite cells (Ecob-Prince et al, 1986; Mouly et al, 1993). In treated muscles (TAΔDHPR), subsequent atrophy was associated with fibrosis (Figure 2C) as shown by morphometric analysis using Sirius red for staining collagen deposition (Figure 2D). Injected TAΔDHPR muscles displayed a four-fold increase in fibrosis compared with the contralateral muscles (Figure 2E). Muscle function was assessed by the measurement of in situ muscle force in response to nerve stimulation, as described earlier (Vignaud et al, 2005a). Compared with values measured in the contralateral limb, absolute and specific maximal isometric tetanic force measured in injected TAΔDHPR were shown to be reduced by 62% (40±11 g compared to 106±20 g, P⩽0.001, n=8) and 34% (1.1±0.3 g/mg compared to 1.6±0.2 g/mg, P⩽0.05, n=8), respectively (Table 1). Figure 2.Consequence of α1S subunit knockdown. (A) Six months post injection, TAs from eight mice were dissected and weighed. (B) The internal diameters (shortest diameter) from all fibres throughout the total muscle section were recorded and analysed. Muscles from five different animals were examined. The bar graph presents mean±s.e.m. of the number of myofibres by fibre diameter class for TAΔDHPR (black) and TACtrl (grey). (C) Transversal sections of TAΔDHPR (ΔDHPR) and TACtrl (c) were stained with haematoxylin and eosin, bars represent 100 μm. (D) To quantify fibrosis, transversal sections of total muscle were stained with Red Sirius and quantified using Histolab Software (marked in blue), data were normalized with total surface of each muscle (orange line surrounding the sections), bars represent 500 μm. (E) Quantification is presented in bar graph and showed 4.1±0.1 fold increase of fibrosis in TAΔDHPR (ΔDHPR) compared to TACtrl (c) (**P<0.001, n=4). Download figure Download PowerPoint Table 1. Morphometric and functional characteristic of TAΔDHPR TACtrl TAΔDHPR Mass (mg) 65±8 39±3** Total number of fibres 3405±161 3249±443 Mean of CSA (mm) 43±4 25±2** MHC composition (%) MHC-1 0.09±0.03 0.03±0.03 MHC-1/2a 0.00 0.02±0.01 MHC-2a 1.5±0.7 0.3±0.06* MHC-2a/2x 2.7±0.4 1.08±0.50 MHC-2x 38.90±1.60 33.50±3.50 MHC-2x/2b 16.40±3.90 16.50±1.30 MHC-2b 40.40±3.50 47.80±4.80 Absolute MTI force (g) 106±20 40±11** Specific MTI force (g/mg) 1.6±0.2 1.1±0.3* Investigations 6 months post injection of AAV1 (U7-ESE) and AAV1 (U7-SA) into the TA (TAΔDHPR), or AAV1 (U7-Ctrl) (TACtrl) were carried out; n=8 for mass, total number of fibres, mean of cross-section area (CSA), absolute and maximal tetanic isometric (MTI) force investigations; and n=3 for MHC composition analyses. Values are mean±s.e.m. and were significatively different for TAΔDHPR from TACtrl, **P⩽0.001; *P⩽0.05. Altogether, these data show that α1S knockdown in normal muscle resulted in important morphological changes involving atrophy and fibrosis. These alterations were not caused by muscle paralysis although fibre dismantling ultimately affected muscle function. α1S knockdown induces nNOS redistribution, FoxO3A nuclear translocation and upregulation of autophagy-related gene expression It has been proposed that nNOS, a molecular partner of the dystrophin–glycoprotein complex, has a role in the mechanism of atrophy in denervation-induced and hindlimb unloading models (Suzuki et al, 2007). In TAΔDHPR muscles, levels of nNOS mRNA were found to be reduced by 41±1% (P⩽0.001, n=9) (Figure 3A), a significant decrease that was confirmed at the protein level by western blot analysis (Supplementary Figure S3A). Immuno-histochemical analysis revealed that nNOS staining shifted from the sarcolemma to the cytoplasm (Figure 3B), a redistribution that was not due to the disruption of the dystrophin–glycoprotein complex, which was not affected in TAΔDHPR muscles (see stainings for dystrophin and associated α, β and γ sarcoglycans in Supplementary Figure S4). We evaluated NOS activity by using NADPH-d histochemical analysis (Chen et al, 2008). As expected, NOS activity was redistributed towards the cytoplasm in TAΔDHPR atrophic fibres (Supplementary Figure S3B). Figure 3.nNOS membrane dissociation, FoxO3A accumulation in myonuclei and upregulation of autophagy-related genes as consequences of α1S subunit loss. (A) mRNA from TAΔDHPR and TACtrl tissues were extracted and nNOS expression was quantified by qRT–PCR. Results are expressed as mean±s.e.m., **P<0.001, n=4. (B) Transversal cryo-sections of TAΔDHPR (ΔDHPR) and TACtrl (c) were stained with anti-nNOS (red), anti-laminin (green) antibodies, nuclei with Dapi (blue) and imaged by confocal microscopy. Bars represent 20 μm. (C) The tissues from extracts were analysed by western blot with FoxO3a-P antibody and normalized with α-actin antibody. Graph depicts mean±s.e.m. of relative expression of FoxO3a-P determined by densitometry and nomalized to the α-actin expression for each muscle, *P⩽0.005, n=3. (D) Longitudinal cryo-sections of TAΔDHPR (ΔDHPR) and TACtrl (c) were stained with anti-FoxO3a (red), anti-laminin (green) antibodies, nuclei with Dapi (blue) and imaged by confocal microscopy. Bars represent 20 μm. (E) mRNA from TAΔDHPR and TACtrl tissues were extracted and regulation of autophagy genes expression was followed by qRT–PCR. Bnip, CathepsinL, LC3 and PI3KIII expression (noted in red) were significantly increased in TAΔDHPR compared with the controlateral TACtrl, **P<0.001, n=4. Download figure Download PowerPoint It has been shown in tail suspension experimental models that nNOS redistribution induced muscle atrophy through regulation of FoxO3A (Suzuki et al, 2007), a pivotal transcription factor involved in muscle mass regulation (Mammucari et al, 2007). Phosphorylated forms of FoxO3A are found in the cytoplasmic compartment, whereas dephosphorylated forms translocate to the nucleus to activate transcription of target genes (Cao et al, 2005). Western blot analysis of TAΔDHPR muscles using an antibody recognizing the phosphorylated form showed that FoxO3A was mainly dephosphorylated (Figure 3C). Concurrently, immuno-staining of TAΔDHPR muscle sections with an antibody recognizing the non-phosphorylated form of FoxO3A showed its accumulation in myonuclei (Figure 3D), thus confirming its translocation after activation. It is known that the ubiquitin-proteasomal pathway is markedly increased in atrophying muscle because of transcriptional activation of ubiquitin, several proteasomal subunit genes and the two muscle-specific ubiquitin ligases atrogin-1/MAFbx and MuRF1 (Cao et al, 2005). In TAΔDHPR muscle extracts, qRT–PCR revealed that atrogin-1/MAFbx and MuRF1 were not over-expressed at 6 month post injection (Figure 3E). Moreover, atrogin-1/MAFbx and MuRF1 were not even transiently upregulated before this time point (expression of these genes was measured 1, 2, and 4 months post injection) suggesting that the ubiquitin-proteasome pathway was not involved in TAΔDHPR induced atrophy. We then inquired whether the induction of atrophy was accompanied by an upregulation of autophagy-related genes, LC3, Gabarapl1, Atg4b and Atg12; the genes involved in regulating autophagy Vps34 (a class III PI3K), Bnip and finally the lysosomal proteinase cathepsinL gene (Kelekar, 2008). By using qRT–PCR, we showed that Bnip3, LC3, PI3KIII and CathepsinL expression was significantly increased in TAΔDHPR compared to contralateral TACtrl (Figure 3E). These results indicate that knockdown of α1S levels induced atrophy through the autophagy pathway that was most probably triggered by nNOS redistribution, FoxO3A translocation to myonuclei and activation of autophagy-related gene expression. Fall of α1S induces autophagosome formation and alters morphology of the sarcoplasmic reticulum Autophagic activity and presence of autophagic vacuoles were revealed at the ultrastructural level by using electron microscopy (EM). Characteristic double-membrane-limited structures corresponding to autophagosomes were observed in atrophic TAΔDHPR muscle fibres (Figure 4A). Autolysosome structures, characterized by highly electron dense signals and corresponding to the fusion of autophagosomes and lysosomes, were frequently observed. The occurrence of autophagosomes was also assessed on tissue sections and isolated fibres from TAΔDHPR muscles by using LC3 immuno-staining, a specific marker for autophagosome formation (Mizushima et al, 2004; Mizushima and Kuma, 2008). We found that LC3 staining on TAΔDHPR transverse sections labelled small vacuolar structures surrounded by dystrophin (Figure 4B). We then carried out double staining on isolated fibres by using antibodies against LC3 and the polyubiquitin-binding protein p62 (also known as sequestosome SQSTM1), which serves as an adaptor molecule mediating the degradation of polyubiquitinated proteins by the autophagic pathway (Raben et al, 2008). Confocal analysis showed widespread rising of colocalized LC3 and p62 staining all along ΔDHPR atrophied fibres, whereas control fibres did not displayed such a phenomenon (Figure 4C). Figure 4.Formation of autophagosomes in TAΔDHPR. (A) Ultrathin sections were imaged by electron microscopy, M, mitochondria; C, collagen fibres; T, T-tubule. Arrows show double-membrane vesicules called auphagosomes. (B) Transversal cryo-sections were stained with anti-LC3b (red), anti-dystrophin (green) antibodies, nuclei with Dapi (blue) and imaged by confocal microscopy. Upper panel: TACtrl (Ctrl) and lower panel: TAΔDHPR (ΔDHPR). Bars represent 20 μm. (C) Myofibres isolated from control (Ctrl) or 6 months post-injected FDB (ΔDHPR) muscles were processed for immuno-fluorescent labelling for P62 (green) and LC3b (red) and imaged by confocal microscopy. Scale bars, 10 μm. Download figure Download PowerPoint EM analysis of TAΔDHPR muscles also revealed that sarcomeres were smaller and less organized than controls, although Z and M lines were still properly aligned. Triads were correctly located and displayed electron dense structures enclosing the EC coupling machinery, but most of the terminal SR cisternae were unusually dilated and the longitudinal SR displayed unusual circumvolutions and invaginations (Figure 5A). Confocal analysis by using antibodies against SERCA, a marker of longitudinal SR, and RyR1, a marker of triads, confirmed this SR disorganization at the level of whole isolated fibres (Figure 5B). Western blot analysis showed that levels of RyR1 were only slightly reduced in TAΔDHPR muscles (Figure 6A). Triadin, a protein associated to the EC complex, was also not significantly affected (data not shown). Confocal double stainings showed that α1S and RyR1 were still co-localized in atrophic TAΔDHPR fibres (Figure 6B), a result in agreement with EM observations and the fact that atrophic TAΔDHPR fibres were still EC coupled. Figure 5.Impact of the α1S subunit knockdown in maintenance of muscle ultrastructure. (A) Ultrathin sections of TAΔDHPR (ΔDHPR) and TACtrl (c) were imaged by electron microscopy. Cis, terminal cisternae; T, tubule; SR, sarcoplasmic reticulum. Bars: 500 nm. (B) Myofibres isolated from Ctrl or ΔDHPR FDB muscles 6 months post injection were processed for immuno-fluorescent labelling for RyR1 (green), SERCA (red), Dapi (blue) and imaged by confocal microscopy. Scale bars, 10 μm. Download figure Download PowerPoint Figure 6.α1S subunit knockdown and RyR1 expression. (A) Lysates from TAΔDHPR (ΔDHPR) and TACtrl (c) were immuno-blotted for RyR1 or α-actin. (n=4). Graph depicts mean±s.e.m. of relative expression of RyR1 determined by densitometry and nomalized to the α-actin expression for each muscle. Results were expressed in protein levels of RyR1 in TAΔDHPR normalized to TACtrl for each mice, *P⩽0.005, n=4. (B) Longitudinal cryo-sections of TAΔDHPR (ΔDHPR) and TACtrl (c) were stained with anti-α1S subunit (red), anti-RyR1 (green) antibodies and imaged by confocal microscopy. Bars=20 μm. Download figure Download PowerPoint Atrophy in ΔDHPR muscle fibres correlates with the disappearance of the sarcolemmal fraction of α1S As some α1S expression remained preserved in exon skipping-treated muscles undergoing atrophy, we wondered in which compartment α1S was really decreased. To address this point, we have investigated more deeply α1S localization in treated muscles, particularly when atrophy was clearly established. This study was carried out by 3D confocal analysis on isolated flexor digitorum brevis (FDB) muscle fibres (Figure 7; Supplementary Movies S5A and S5B). In control fibres, α1S was chiefly found in the inner compartment encompassing T-tubules and triads. However, 3D-projection analysis revealed that a minor fraction of α1S was present on the sarcolemma as already shown by using EM techniques and suggested by subcellular fractionation (Jorgensen et al, 1989; Munoz et al, 1995; Vassilopoulos et al, 2009). In exon skipping-treated fibres undergoing atrophy, α1S was still present in the inner compartments and displayed a quite preserved striated distribution. The major effect concerned the sarcolemmal α1S fraction, which was missing in treated fibres. Figure 7.Localization of α 1S subunit. Whole skeletal muscle fibres enzymatically isolated from Ctrl (A–D) or ΔDHPR (E–H) FDB muscles 6 months post injection were processed for immuno-fluorescent labelling for α1 S subunit (red) and laminin (green). Scale bars, 10 μm; Arrows indicate α1 S subunit expression on sarcolemma; antibody labelling was visualized by serial confocal microscopy and represented as movies of the optical sections (Supplementary Movies S5A and S5B). (D, H) Present projections of confocal Z-series (step between each frame is 1 μm) along XZ and YZ planes