Sarcolipin overexpression impairs myogenic differentiation in Duchenne muscular dystrophy

Abstract

Reduction in the expression of sarcolipin (SLN), an inhibitor of sarco(endo)plasmic reticulum (SR) Ca²⁺-ATPase (SERCA), ameliorates severe muscular dystrophy in mice. However, the mechanism by which SLN inhibition improves muscle structure remains unclear. Here, we describe the previously unknown function of SLN in muscle differentiation in Duchenne muscular dystrophy (DMD). Overexpression of SLN in C₂C₁₂ resulted in decreased SERCA pump activity, reduced SR Ca²⁺ load, and increased intracellular Ca²⁺ (${\text{Ca}}_{\text{i}}^{\text{2+}}$) concentration. In addition, SLN overexpression resulted in altered expression of myogenic markers and poor myogenic differentiation. In dystrophin-deficient dog myoblasts and myotubes, SLN expression was significantly high and associated with defective ${\text{Ca}}_{\text{i}}^{\text{2+}}$ cycling. The dystrophic dog myotubes were less branched and associated with decreased autophagy and increased expression of mitochondrial fusion and fission proteins. Reduction in SLN expression restored these changes and enhanced dystrophic dog myoblast fusion during differentiation. In summary, our data suggest that SLN upregulation is an intrinsic secondary change in dystrophin-deficient myoblasts and could account for the ${\text{Ca}}_{\text{i}}^{\text{2+}}$ mishandling, which subsequently contributes to poor myogenic differentiation. Accordingly, reducing SLN expression can improve the ${\text{Ca}}_{\text{i}}^{\text{2+}}$ cycling and differentiation of dystrophic myoblasts. These findings provide cellular-level supports for targeting SLN expression as a therapeutic strategy for DMD.

INTRODUCTION

Duchenne muscular dystrophy (DMD) is the most common form of muscular dystrophy caused by mutations in the DMD gene encoding dystrophin. Loss of dystrophin protein results in salient clinical features such as muscle wasting, respiratory failure, and cardiomyopathy (7, 15, 21, 23, 31, 40, 42). Studies have shown that muscle degeneration, improper muscle fiber regeneration, and fibrosis are major causes of muscle wasting in DMD (9, 46, 51). Emerging studies have suggested that intrinsic defects in mechanisms causing satellite cell differentiation in dystrophic muscles could impair the muscle regeneration in DMD (9). However, the molecular mechanism(s) that initiates these defects in dystrophic muscles is poorly understood.

It is well documented that intracellular Ca²⁺ (${\text{Ca}}_{\text{i}}^{\text{2+}}$) concentration is a critical regulator of muscle cell differentiation (10, 38). One of the major morphological events associated with muscle differentiation is the formation of sarco(endo)plasmic reticulum (SR), a highly specialized internal Ca²⁺ store (17, 41). Studies have shown that maintaining the SR Ca²⁺ content is an important modulator of ${\text{Ca}}_{\text{i}}^{\text{2+}}$ levels (36, 53) and muscle differentiation (17, 45, 47). In dystrophic muscles, abnormal Ca²⁺ cycling through sarcolemma and SR increases the ${\text{Ca}}_{\text{i}}^{\text{2+}}$ load, an important stimulus that triggers pathological events such as activation of Ca²⁺-dependent proteases, improper muscle regeneration, and necrosis (1, 2, 8, 22, 57). Among several mechanisms that are responsible for Ca²⁺ dysregulation, improving the SR Ca²⁺-ATPase (SERCA) function, which restores Ca²⁺ to the SR during excitation-contraction coupling, is found to be highly effective in normalizing the ${\text{Ca}}_{\text{i}}^{\text{2+}}$ levels and mitigating the muscle disease in DMD (18, 19, 32, 49). Recently, we have shown that sarcolipin (SLN), an inhibitor of the SERCA pump, is significantly elevated in cardiac and skeletal muscles of animal models and patients with DMD (43, 55). These findings suggested that SLN upregulation could be a major cause of SERCA dysfunction in DMD. Accordingly, reducing SLN expression improved SERCA function and ameliorated the severe muscular dystrophy in a phenotypic mouse model of DMD (55). These studies also indicated that reduction in SLN expression can improve the muscle regeneration in DMD (55).

On the basis of the above findings, we hypothesized that high levels of SLN in dystrophic myoblasts could cause SERCA inhibition and ${\text{Ca}}_{\text{i}}^{\text{2+}}$ overload, which in turn impairs myogenic differentiation in DMD. Here, we tested this hypothesis using C₂C₁₂, an immortalized mouse myoblast cell line, and primary myoblasts isolated from a dog model of DMD.

METHODS

Animal studies.

Studies using mdx and SLN knockout (sln^−/−) mice were approved by the Institutional Animal Care and Use Committee of New Jersey Medical School, Rutgers University, Newark, New Jersey. The experimental dogs were on a mixed genetic background consisting of golden retriever, Labrador retriever, Corgi, and beagle and generated in-house at the University of Missouri by artificial insemination. Affected dogs carry various mutations in the dystrophin gene that abort dystrophin expression (29). Tissues for the isolation of primary myoblasts were obtained from normal and DMD dogs following protocols approved by the Institutional Animal Care and Use Committee of University of Missouri, Columbia, Missouri.

Mice.

We used 3–4-mo-old male and female mice on C57BL/6 background for the experiments. The mdx:sln^+/− and mdx:sln^−/− mice were generated by crossing the mdx mice (43) with sln^−/− mice (4). Mice were kept under a 12-h light-dark cycle with a temperature of 22–24°C and 60–70% humidity and fed ad libitum with a normal chow diet. The genotypes of the mice were identified by PCR analysis as previously described. No samples, mice, or data points were excluded from the data analysis. Animals were not randomized except for the genotypes. Pectoral tissues were dissected out of wild-type, mdx, mdx:sln^+/−, and mdx:sln^−/− mice, rinsed in sterile PBS, and flash-frozen in liquid nitrogen.

Adenovirus.

Recombinant adenovirus expressing mouse SLN (Ad.SLN) was generated as described before (5). Briefly, mouse SLN cDNA was cloned into the pShuttle-CMV vector and transfected into HEK 293 cells, and the recombinant adenoviruses were harvested 7–10 days later. The viral titer was determined by plaque assay. Adenovirus harboring β-galactosidase (Ad.LacZ) was used as a control.

Adeno-associated virus.

The short hairpin RNA specific for canine SLN (cshSLN; 5′-CTGTTTCTCAACTTCACTATTGTTTCAAGAGAACAATAGTGAAGTTGAGAAACAG-3′) was cloned into the adeno-associated virus (AAV) vector as described before (55). Recombinant AAV was produced using the AAV-DJ/8 helper-free expression system (Cell Biolabs). Briefly, the recombinant AAV vector expressing cshSLN (pAAV.cshSLN) was cotransfected into the 293AAV cell line along with pAAV-DJ/8 and pHelper in a 1:1:1 ratio using polyethylenimine. After 72 h of transfection, cells were harvested and purified by the iodixanol gradient/ultracentrifugation method. The AAV fraction obtained from the gradient centrifugation was concentrated by a Vivaspin 20 concentrator (100-kDa cutoff; Sartorius, Germany). The virus titer was determined using the Cell Biolabs AAV quantitation kit.

C₂C₁₂ mouse myoblasts.

C₂C₁₂ myoblasts were purchased from the American Type Culture Collection (CRL-1772) and cultured in Dulbecco’s modified Eagle’s medium (DMEM) without sodium pyruvate supplemented with 10% fetal bovine serum (FBS) and antibiotics at 37°C in a humidified atmosphere of 5% CO₂ and 95% air. Cells were stored as aliquots after four passages and used for further experiments. Differentiation was induced when cells were fully confluent by replacing the growth medium with DMEM supplemented with 2% horse serum (Hyclone). The differentiation medium was replaced every 48 h. Adenoviral infection (500 multiplicity of infection) into C₂C₁₂ cells was performed while switching to the differentiation medium.

Primary dog myoblasts.

Primary myoblasts were isolated from the biceps femoris muscle of normal and DMD dogs as described before (6, 11). Briefly, a 1-cm³ tissue sample was collected from each dog, posteuthanasia, cleaned of connective tissue, and physically dissociated by mincing, followed by enzymatic digestion for 1 h at 37°C in Ham’s F-12 medium containing collagenase type IV (Worthington) and liberase (Sigma-Aldrich). After filtration, cells were recovered in PromoCell Skeletal Muscle Cell Growth Medium (cat. no. C-23060), containing 20% FBS, and supplemented with 50 µg/mL fetuin (bovine), 10 ng/ml EGF (recombinant human), 1 ng/mL basic FGF (recombinant human), 0.4 µg/mL dexamethasone, 10 µg/mL insulin (recombinant human), and 1% penicillin-streptomycin. Cells were preplated for 24 h; then the nonadherent myoblasts were passaged to gelatin-coated plates for expansion. Myoblasts were maintained at 37°C in a humidified incubator at 5% CO₂ and 95% air. Upon reaching 70% confluence, myoblasts were frozen as aliquots in freezing medium (Ham’s F-12 nutrient mixture) containing 15% horse serum and 20% DMSO for further studies. Differentiation was induced when cells were fully confluent by replacing the growth medium with DMEM supplemented with gentamycin (50 μg/mL) and insulin (10 μg/mL). To knock down SLN expression in dog primary myoblasts, cells were infected with AAV.cshSLN at 1 × 10¹⁰ viral genomes per milliliter of culture medium 24 h before differentiation. The uninfected normal and DMD myoblasts were used as controls. For autophagy experiments, chloroquine (15 mmol/L) was added to the differentiation medium 24 h before harvesting the cells.

Alamar blue assay.

Cell viability was measured by Alamar blue assay. Briefly, C₂C₁₂ and the dog primary myoblasts grown on 96-well plates were infected with Ad.SLN, Ad.LacZ, or AAV.cshSLN as described above. After 4 (dog myoblasts) or 5 (C₂C₁₂) days of differentiation, the media were replaced with 0.1 mL of media containing 10% (vol/vol) Alamar blue, and cells were incubated for 90 min at 37°C, 5% CO₂ and 95% air. The plate was then read for absorbance at 570 and 600 nm. Wells containing media only and wells containing media plus Alamar blue were used for correcting background absorbance in test wells. Overall viability of the cells was calculated over the negative wells with no cells. Data are presented as relative fluorescence units.

Oxygen consumption rate assays using a Seahorse analyzer.

The respiratory complex activities in differentiated dog myotubes were measured using a Seahorse XF24 analyzer. Briefly, primary myoblasts from the normal and DMD dogs were plated at a density of 6,000 cells per well on collagen-coated Seahorse plates. Cells were infected or not infected with AAV.cshSLN and differentiated as described above. Four days after differentiation, the medium was replaced with unbuffered DMEM supplemented with 1 mg/mL glucose, 1 mM pyruvate, and 4 mM glutamine for 1 h in a non-CO₂ incubator. The Mito Stress test was performed as per the manufacturer’s recommendations (Seahorse; Agilent Technologies). Oxygen consumption rate (OCR) was measured at baseline and after the sequential addition of the following: 1) oligomycin (1 µM final), a complex V ATP synthase inhibitor used to measure ATP-dependent oxygen consumption; 2) FCCP (1.5 µM final), an uncoupler carbonyl cyanide 4-(trifluoromethoxy) phenylhydrazone used to measure maximum respiratory capacity; and 3) antimycin A and rotenone (each 1 µM final), which are complex I and III inhibitors, respectively, used to measure nonmitochondrial OCR. Calculation of basal OCR, percent spare respiratory capacity, proton leak, coupling efficiency, nonmitochondrial respiration, and extracellular acidification rate (ECAR) was performed using Excel/Mito Stress Report generator (Seahorse; Agilent Technologies).

Quantitative real-time polymerase chain reaction.

Total RNA was extracted from the cells using Qiagen RNA isolation kit. Complementary DNA (cDNA) was prepared from 1 μg of total RNA using the High Capacity cDNA Reverse Transcription Kit (Thermo Fisher Scientific). The quantitative RT-PCR analysis was performed using SuperScript RT-PCR kit (Invitrogen). The primer sequences are as follows: SLN forward, TCAGGAAGTGAAGACAAGCC; SLN reverse, GGAGCCACATAAGGAGAACG; myogenin forward, GCAATGCACTGGAGTTCG; myogenin reverse, ACGATGGACGTAAGGGAGTG; GAPDH forward, GTCGTGGATCTGACGTGCC; and GAPDH reverse, ATGCCTGCTTCACCACCTTC. The PCR products were verified by 2% agarose gel electrophoresis. The mRNA levels of SLN and myogenin were normalized to the mRNA levels of glyceraldehyde-3-phosphate dehydrogenase (GAPDH).

SR Ca²⁺ load and Ca²⁺ transient measurements.

The SR Ca²⁺ load and Ca²⁺ transients were measured using fluo-4 AM at 34–36°C, as described earlier (58). C₂C₁₂ cells and primary myoblasts were cultured on collagen-coated coverslip dishes (no. P35Gcol-1.5-14C; MatTek) and differentiated for 5 and 4 days, respectively. Cells were then loaded with the Ca²⁺ indicator fluo-4 AM (F-14201; Molecular Probes), placed in a heated chamber on an inverted microscope, and field stimulated at 0.5 Hz to maintain consistent SR Ca²⁺ load. The fluo-4 fluorescence was excited at ∼485 nm, and the emissions were measured at ∼530 nm using a Nikon Eclipse TE200 inverted microscope with a Fluor ×40 oil objective lens (numerical aperture 1.3). The fluorescence signals were recorded using an Andor Ixon charge-coupled device (CCD) camera (Andor Technology) operated with Imaging Workbench software (INDEC BioSystems) at 50 frames per second with a spatial resolution of 500 × 400 pixels. Fluorescence intensity was measured as the ratio of the fluorescence (F) to the basal diastolic fluorescence (F₀). The amplitude of the 10 mmol/L caffeine-induced Ca²⁺ transient was used as a measure of total SR Ca²⁺ content. Fractional SR Ca²⁺ release was calculated by dividing the height of the last twitch transient by the height of the caffeine transient.

Immunofluorescence.

C₂C₁₂ and the dog primary myoblasts grown on collagen-coated coverslip dishes were infected with Ad.SLN, Ad.LacZ, or AAV.cshSLN and differentiated as described above. After differentiation, cells were fixed in 4% paraformaldehyde for 20 min at room temperature, washed twice in phosphate-buffered saline (PBS), and blocked for 45 min in 1% BSA and 2% horse serum in PBS. The cells were stained for myosin using the mouse monoclonal myosin heavy chain antibody MF20 (1:10; Developmental Studies Hybridoma Bank) overnight at 4°C, followed by Alexa Fluor 488 or 594 secondary antibody (Thermo Fisher Scientific) for 1 h, and processed. Images were obtained using Olympus BX51 microscope and Cell Sense standard software, respectively. The fusion index for C₂C₁₂ was calculated as the percentage of nuclei within myotubes containing at least 3 nuclei as a ratio of total nuclei (Ad.LacZ = 1,635 ± 63 nuclei per field and Ad.SLN = 2,167 ± 36 nuclei per field) in 8–15 randomly selected, nonoverlapping fields per coverslip. For dystrophin-deficient dog myoblasts, the fusion was quantitated by calculating the percentage of nuclei present in myosin-positive myotubes with the indicated number of nuclei (≤3 or >3 nuclei) in 5–8 randomly selected nonoverlapping fields per coverslip.

Western blot analysis.

Total protein extract was prepared by lysing the cells with radioimmunoprecipitation assay buffer (in mmol/L: 50 Tris, pH 7.4; 150 NaCl; 1 EDTA; and 0.5% Nonidet P-40) supplemented with PMSF (1 mmol), NaVO₃ (5 mmol), okadaic acid (10 nmol), NaF (1 mmol), and benzamidine (1 mmol). Tissue homogenization was performed in lysis buffer (in mmol/L: 50 Tris, pH 7.4; 150 NaCl; 1 EDTA; and 0.5% Nonidet P-40) supplemented with PMSF, NaVO₃, okadaic acid, NaF, and benzamidine. Equal amounts of protein extracts prepared from the cells/tissues were separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) along with prestained molecular weight markers and transferred to nitrocellulose membranes for 1 h at room temperature. After transfer, the membranes were stained with Ponceau S and cut into strips based on the molecular weight of each protein studied. The membrane strips were then blocked with 3% milk in PBS and probed overnight at 4°C using antibodies specific for SLN (anti-rabbit, 1:3,000; 3); SERCA1 (anti-rabbit, 1:2,000, custom-made; 3); SERCA2a (anti-rabbit, 1:5,000, custom-made; 3); calsequestrin (CSQ, anti-rabbit, 1:5,000, no. PA-1-913; Thermo Fisher Scientific), which recognizes both cardiac and skeletal isoforms; myoblast determination protein-1 (MyoD, anti-mouse, 1:1,000, SC-377460; Santa Cruz Biotechnology); dystrophin (anti-mouse, 1:100, no. mandys8; Developmental Studies Hybridoma Bank); myogenin (anti-mouse, 1:100, F5D-s; Developmental Studies Hybridoma Bank); caveolin-3 (anti-mouse, 1:2,000, SC-5310; Santa Cruz Biotechnology); utrophin (anti-mouse, 1:100, 8A4; Developmental Studies Hybridoma Bank); pan-calcineurin A (anti-rabbit, 1:1,000, no. 2614; Cell Signaling Technology); stabilin 2 (Stab2, anti-rabbit, 1:2,000, no. PA-5-55447; Thermo Fisher Scientific); mitofusin 2 (Mfn2, anti-mouse, 1:1,000, SC-515647/F-5; Santa Cruz Biotechnology); sequestosome 1 (SQSTM1/P62, anti-mouse, 1:5,000, H-00008878-M01; Abnova); dynamin-1-like protein [DLP1, also referred to as dynamin-related protein-1 (Drp1); anti-mouse, 1:2,000, no. 611738; BD Biosciences]; phospho-Drp1-S616 (anti-rabbit, 1:2,000, no. 3455; Cell Signaling Technology); microtubule-associated protein 1A/1B-light chain 3 (LC3, anti-rabbit, 1:2,000; NB-100-2220; Novus Biologicals); β-actin (anti-goat, 1:2,000, SC-1615/C-11; Santa Cruz Biotechnology); myomaker [also referred to as transmembrane protein 8C (TMEM8C); anti-rabbit, 1:500; NBP-2-34175; Novus Biologicals]; or GAPDH (anti-mouse, 1:10,000, G-8795; Sigma). Membranes were incubated with appropriate secondary antibodies for 45 min at room temperature and visualized with SuperSignal West Dura Substrate Kit (Thermo Fisher Scientific) using Bio-Rad ChemiDoc MP Imaging System. Western blot analysis of subunits of oxidative phosphorylation (OXPHOS) protein complexes were performed using OXPHOS Rodent WB Antibody Cocktail (anti-mouse, 1:1,000, ab-110413; Abcam). Quantitation of signals was performed using Image Laboratory version 5.1 software and then normalized to GAPDH or β-actin levels. Western blot analysis was repeated at least three times.

SR Ca²⁺ uptake.

SR Ca²⁺ uptake was measured following the Millipore filtration technique as previously described (48). Briefly, ~150 μg of total protein extract were incubated at 37°C in 1.5 mL of Ca²⁺ uptake medium (in mmol/L: 40 imidazole, pH 7.0; 100 KCl; 5 MgCl₂; 5 NaN₃; 5 potassium oxalate; and 0.5 EGTA) and various concentrations of CaCl₂ to yield 0.03–3 μmol/L free Ca²⁺ (containing 1 μCi/μmol ⁴⁵Ca²⁺). To obtain the maximal stimulation of SR Ca²⁺ uptake, ruthenium red was added to a final concentration of 1 μmol immediately before the addition of the substrates to begin the Ca²⁺ uptake. The reaction was initiated by the addition of ATP to a final concentration of 5 mmol and terminated at 1 min by filtration. Each assay was performed in duplicate. The rate of SR Ca²⁺ uptake and the Ca²⁺ concentration required for EC₅₀ were determined by nonlinear curve-fitting analysis using GraphPad Prism v6.01 software.

Statistical analysis.

Each experiment was repeated at least three times, and the data were analyzed in an unbiased and blinded manner to ensure that these results are consistent and reproducible. All statistical analyses were performed using GraphPad Prism v6.01 software. Differences were determined using a two-tailed, unpaired Student’s t test with Welch’s correction. Two-way analysis of variance (ANOVA) with post hoc Bonferroni correction was used for multigroup comparison when necessary. A value of P < 0.05 was considered significant.

RESULTS

Overexpression of SLN impairs SR Ca²⁺ handling in C₂C₁₂ cells.

SLN protein expression was undetectable by Western blot analysis in C₂C₁₂ myoblasts as well as in myotubes after 5 days of differentiation (Fig. 1A). The quantitative RT-PCR analysis shows that SLN mRNA expression was at a very low level in C₂C₁₂ myoblasts as evidenced by the low PCR cycle threshold values (quantitation cycle >36 cycles). SLN mRNA expression further declined upon differentiation (Supplemental Fig. S1A; all Supplemental Material is available at https://doi.org/10.6084/m9.figshare.8041427.v3). On the other hand, the mRNA levels of myogenin, a myogenic marker, were increased by 7.5-fold in 2 days of differentiation and then declined as differentiation proceeded (Supplemental Fig. S1A). Western blot analysis further confirmed these changes at protein level (Supplemental Fig. S1B). Therefore, to determine the direct effect of SLN on myogenic differentiation, we chose to study the C₂C₁₂ cell line.

Fig. 1.

Sarcolipin (SLN) overexpression inhibits sarco(endo)plasmic reticulum (SR) Ca²⁺-ATPase (SERCA) function and alters SR Ca²⁺ handling in C₂C₁₂ cells. A: representative Western blots and box-and-whisker plots showing the protein levels of SLN, SERCA isoforms, and calsequestrin (CSQ) in C₂C₁₂ myoblasts and in myotubes after 5 days of differentiation infected with either adenovirus expressing LacZ gene encoding β-galactosidase (Ad.LacZ) or adenovirus expressing SLN (Ad.SLN). The protein levels of SERCA2a and CSQ are significantly increased in Ad.SLN compared with that of Ad.LacZ controls; n = 6 samples per group. B: Ca²⁺-dependent Ca²⁺ uptake is decreased in the Ad.SLN-infected C₂C₁₂ myotubes (n = 5 experiments) compared with that of Ad.LacZ-infected controls (n = 4 experiments). C–F: box-and-whisker plots showing the summary data for twitch Ca²⁺ transients (elicited by field stimulation at 0.5 Hz; C), SR Ca²⁺ content measured as the height of the caffeine (10 mmol/L)-induced Ca²⁺ transient (D), time taken for 50% decay of Ca²⁺ (T₅₀; E), and fractional SR Ca²⁺ release, calculated by dividing the height of the last twitch transient by the height of the caffeine transient (F), in control (Ad.LacZ) and SLN-overexpressing C₂C₁₂ myotubes; n = 12 myotubes per group. Statistical significance was determined by unpaired Student’s t test with Welch’s correction. P values are shown within the histograms. F/F₀, ratio of the fluorescence (F) to the basal diastolic fluorescence (F₀); NS, not statistically significant.

To overexpress SLN, C₂C₁₂ myoblasts were infected with Ad.SLN during differentiation. Ad.LacZ-infected cells were used as controls. The cells were analyzed 5 days after differentiation. We first determined the cell viability in Ad.LacZ- and Ad.SLN-infected C₂C₁₂ myotubes by Alamar blue assay. Results show that the cell viability was not different between Ad.LacZ- and Ad.SLN-infected C₂C₁₂ myotubes indicating that SLN overexpression did not affect the viability of C₂C₁₂ myotubes (Supplemental Fig. S2). We then determined the effect of SLN overexpression on SERCA function and ${\text{Ca}}_{\text{i}}^{\text{2+}}$ homeostasis in cells after 5 days of differentiation. Western blot analysis and quantitation show that the protein levels of SERCA2a and CSQ were significantly increased in SLN-overexpressing cells (Fig. 1A). There was a small increase in SERCA1 protein levels in SLN-overexpressing cells, but this increase was not significantly different from Ad.LacZ-infected control cells. We next determined the SERCA function in SLN-overexpressing cells by Ca²⁺-dependent Ca²⁺ uptake assays. Results show that overexpression of SLN resulted in decreased Ca²⁺-dependent Ca²⁺ uptake (Fig. 1B); however, the maximal velocity of Ca²⁺ uptake was not significantly different between the control and SLN-overexpressing myotubes [Ad.LacZ = 171 + 9 (n = 4) vs. Ad.SLN = 146 + 10 (n = 5) nmol Ca²⁺/mg, P = 0.12]. On the other hand, the EC₅₀ values for Ca²⁺ activation were significantly increased in SLN-overexpressing cells [Ad.LacZ = 265 ± 16 (n = 4) vs. Ad.SLN = 474 ± 58 (n = 5) nmol Ca²⁺/mg, P < 0.05] indicating decrease in the apparent affinity of SERCA pumps for Ca²⁺. We next determined the effect of SLN overexpression on SR Ca²⁺ load and Ca²⁺ transients (Supplemental Fig. S3). The twitch Ca²⁺ transient (Fig. 1C) and the SR Ca²⁺ content as measured by caffeine-induced Ca²⁺ release (Fig. 1D) were significantly decreased in the SLN-overexpressing C₂C₁₂ cells. The duration of twitch transient [expressed as the time of 50% decline (T₅₀); Fig. 1E] and the fractional Ca²⁺ release (Fig. 1F) in SLN-overexpressing cells were not significantly different from the Ad.LacZ-infected control cells.

SLN overexpression inhibits C₂C₁₂ myogenic differentiation.

We next examined the myotube formation capability of C₂C₁₂ myoblasts infected with Ad.SLN by staining with pan-myosin heavy chain antibody. On day 5, the control myoblasts had formed larger myotubes, whereas SLN overexpression resulted in small and thin myotubes with less branching (Fig. 2A). The fusion index, defined as the percentage of nuclei within myotubes containing >3 nuclei, was significantly reduced in SLN-overexpressing cells compared with that of controls (P < 0.0005; Fig. 2B). These findings indicate that SLN overexpression can impair the ability of C₂C₁₂ cells to fuse together and form larger myotubes.

Fig. 2.

Sarcolipin (SLN) overexpression prevents myoblast fusion and differentiation. A: representative immunofluorescence images of C₂C₁₂ myotubes after 5 days of differentiation infected with adenovirus expressing LacZ gene encoding β-galactosidase (Ad.LacZ) or adenovirus expressing SLN (Ad.SLN), stained with the myosin heavy chain antibody MF20. Original magnification is ×10. Scale bars = 100 μm. B: box-and-whisker plots showing the percent fusion index calculated as the percentage of cells containing at least 3 nuclei within a differentiated myotube. A minimum of 8 different nonoverlapping areas from each coverslip slide were used for quantitation. Here, n = 4 experiments per group. C–F: representative Western blots and box-and-whisker blots showing the protein levels of myogenin, myoblast determination protein-1 (MyoD), caveolin 3 (Cav3), stabilin 2, calcineurin, utrophin, and dystrophin as fold changes in myoblasts and in C₂C₁₂ cells after 5 days of differentiation infected with either Ad.LacZ or Ad.SLN; n = 6 samples per group. Statistical significance was determined by unpaired Student’s t test with Welch’s correction. P values are shown within the histograms. NS, not statistically significant.

To explore the mechanisms responsible for the defects in differentiation of SLN-overexpressing cells, we measured the protein levels of the myogenic regulatory factors myogenin and MyoD. Results show that myogenin level was significantly increased in SLN-overexpressing cells after 5 days of differentiation compared with that of controls (Fig. 2, C and D). On the other hand, MyoD level was significantly decreased upon differentiation but to the same extent in both control and SLN-overexpressing C₂C₁₂ cells (Fig. 2, C and D).

It has been shown that stabilin 2 (Stab2) can modulate myoblast fusion during differentiation via the calcineurin (CaN)-nuclear factor of activated T cells (NFAT) pathway (35). A recent study showed that ablation of SLN in mdx mice is associated with decreased levels of Stab2 and CaN (13). Similarly, overexpression of caveolin 3 (Cav3) is shown to inhibit myoblast fusion (56). We therefore next determined whether altered expression of these proteins contributes to the impaired myogenic differentiation of SLN-overexpressing C₂C₁₂ cells. Results show that the Stab2 protein level was increased upon differentiation but to the same extent in both Ad.LacZ- and Ad.SLN-infected C₂C₁₂ cells (Fig. 2E and Supplemental Fig. S4A), whereas CaN expression was decreased in SLN-overexpressing C₂C₁₂ myotubes (Fig. 2, E and F). Cav3 expression was undetectable in proliferating myoblasts, but its expression increased upon differentiation. This increase was more significant in SLN-overexpressing C₂C₁₂ myotubes compared with that of Ad.LacZ-infected controls (Fig. 2, C and D).

SLN overexpression has no effect on dystrophin and utrophin expression in C₂C₁₂ cells.

Ablation of SLN expression is shown to be associated with decreased expression of utrophin in muscles of mdx mice (13). We therefore determined whether SLN overexpression has any effect on the expression levels of utrophin and dystrophin. Results show that utrophin levels were not different between myoblasts and myotubes after 5 days of differentiation. Furthermore, SLN overexpression had no effect on the utrophin protein expression (Fig. 2E and Supplemental Fig. S4B). Dystrophin expression, which was undetectable in proliferating myoblasts, significantly increased upon myogenic differentiation but to the same extent in both control and SLN-overexpressing C₂C₁₂ cells (Fig. 2E and Supplemental Fig. S4C).

SLN is upregulated in dystrophic dog myoblasts.

To determine the role of SLN in dystrophic myoblast differentiation, we studied the primary myoblasts isolated from muscles of normal and dystrophin-deficient DMD dogs. We first examined the expression levels of myogenic markers and SR Ca²⁺-handling proteins in dystrophic myoblasts. Results show that SLN protein expression was detectable in both normal and dystrophin-deficient myoblasts; however, dystrophic myoblasts expressed higher levels of SLN than normal myoblasts (1.8 ± 0.25-fold vs. normal; Fig. 3, A and B). The protein levels of SERCA1, CSQ, and myogenic markers such as myogenin and MyoD were similar between normal and dystrophic myoblasts (Fig. 3A). The SERCA2a isoform was undetectable by Western blot analysis in both normal and dystrophic myoblasts.

Fig. 3.

Reduction in sarcolipin (SLN) expression improves the sarco(endo)plasmic reticulum (SR) Ca²⁺ handling in primary myoblasts from the dog model of Duchenne muscular dystrophy (DMD) after 4 days of differentiation. A: representative Western blots showing the protein levels of SLN, SR Ca²⁺-ATPase 1 (SERCA1), calsequestrin (CSQ), myogenin, and myoblast determination protein-1 (MyoD) in normal and DMD dog myoblasts. B: quantitation showing that SLN is upregulated in the primary myoblasts from the DMD dog; n = 4 samples per group. C: Alamar blue assay showing the viability in normal (N) myoblasts, normal myoblasts treated with adeno-associated virus expressing short hairpin RNA specific for canine SLN (AAV.cshSLN; N/shSLN), DMD myoblasts, and DMD myoblasts treated with AAV.cshSLN (DMD/shSLN) after 4 days of differentiation. Values shown are percent relative fluorescence units relative to normal controls; n = 10 experiments per group. D–F: representative Western blots (D) and box-and-whisker plots (E and F) showing the protein levels of SLN and SERCA isoforms in normal and DMD (D) myoblasts treated with AAV.cshSLN after 4 days of differentiation; n = 4 samples per group. G–J: box-and-whisker plots showing the summary data for twitch Ca²⁺ transients (elicited by field stimulation at 0.5 Hz; G), time taken for 50% decay of Ca²⁺ (T₅₀; H), SR Ca²⁺ content measured as the height of the caffeine (10 mmol/L)-induced Ca²⁺ transient (I), and fractional SR Ca²⁺ release, calculated by dividing the height of the last twitch transient by the height of the caffeine transient (J), in normal and DMD myotubes; n = 14 myotubes per group. Statistical significance was determined by unpaired Student’s t test with Welch’s correction. P values are shown within the histograms. F/F₀, ratio of the fluorescence (F) to the basal diastolic fluorescence (F₀); NS, not statistically significant.

Reduction in SLN expression alters SR Ca²⁺ handling in dystrophic myotubes.

To determine the effect of SLN reduction on dystrophic dog myoblast differentiation, we knocked down the SLN expression in both normal and dystrophic dog myoblasts using AAV.cshSLN. The cells were analyzed 4 days after induction of differentiation. We first determined whether the AAV treatment and SLN knockdown affected the cell growth and viability in normal and DMD myoblasts during differentiation using Alamar blue assay. Results (Fig. 3C) show that there was no difference in cell viability between normal and DMD myotubes after 4 days of differentiation. Furthermore, AAV.cshSLN treatment had no effect on the cell viability. In dystrophic myotubes, SLN protein level was significantly increased compared with that of normal controls (1.45 ± 0.06-fold vs. normal). AAV treatment, on the other hand, reduced the SLN protein levels ~50% in both normal and dystrophic myotubes (Fig. 3, D and E). The SERCA2a level was increased upon differentiation but to a lesser extent in dystrophic myotubes (0.59 ± 0.05-fold vs. normal; Fig. 3, D and F). The expression level of SERCA1 was significantly reduced in dystrophic myotubes compared with that of normal controls (0.41 ± 0.08-fold vs. normal; Fig. 3, D and F). In normal myotubes, AAV treatment had no significant effect on the SERCA2a expression, whereas the expression of SERCA1 increased upon AAV treatment. In dystrophic myotubes, AAV treatment reduced the SERCA2a expression but increased the SERCA1 expression compared with that of untreated controls (Fig. 3, D and F). CSQ levels were similar in both normal and dystrophic myotubes (Fig. 3D).

We next determined the SR Ca²⁺ load and Ca²⁺ transients in primary myoblasts after 4 days of differentiation. The twitch Ca²⁺ transient (Fig. 3G) was not different between normal and dystrophic myotubes, whereas the duration of twitch transient [expressed as the time of 50% duration (T₅₀); Fig. 3H] was significantly longer in the dystrophic myotubes. AAV treatment had no effect on the Ca²⁺transients obtained from normal and dystrophic myotubes. The SR Ca²⁺ content as measured by caffeine-induced Ca²⁺ release (Fig. 3I) was significantly higher in the dystrophic myotubes, whereas AAV treatment significantly reduced the SR Ca²⁺ content and T₅₀ in the dystrophic myotubes compared with that of untreated controls (Fig. 3, H and I). The fractional Ca²⁺ release was significantly reduced in the dystrophic myotubes, whereas AAV treatment normalized these changes (Fig. 3J).

Reduction in SLN expression improves myogenic differentiation of dystrophic myoblasts.

Loss of dystrophin expression in dystrophic myotubes was verified by Western blot analysis (Fig. 4A). Results also show that the expression levels of utrophin were unaltered between normal and dystrophic myotubes. Furthermore, AAV treatment had no effect on the utrophin protein expression (Fig. 4A).

Fig. 4.

Reduction in sarcolipin (SLN) expression improves dystrophic myoblast fusion. A: representative Western blots showing the protein levels of myogenic markers and utrophin in normal (N) and Duchenne muscular dystrophy (DMD; D) myoblasts treated with adeno-associated virus expressing short hairpin RNA specific for canine SLN (shSLN; AAV.cshSLN) after 4 days of differentiation. B: quantitation showing that myogenin is downregulated in the AAV.cshSLN-treated DMD myotubes; n = 4 samples per group. *P < 0.05 for myogenin in the DMD/shSLN group vs. other groups. C: representative immunofluorescence images of control and AAV.cshSLN-treated primary myoblasts after 4 days of differentiation from the normal and DMD dogs, stained with the myosin heavy chain antibody MF20. Original magnification is ×10. Scale bars = 100 μm. Arrows indicate the multinucleated myotubes; n = 4 experiments per group. D: box-and-whisker plots showing the myosin-positive cells containing ≤3 and >3 nuclei in DMD myoblasts and in DMD myoblasts treated with AAV.cshSLN. A minimum of 5 different nonoverlapping areas from each coverslip slide were used for quantitation; n = 4 experiments per group. Cav3, caveolin 3.

We next determined the effect of SLN reduction on the expression levels of myogenic markers. MyoD protein was undetectable in both normal and dystrophic myotubes. There was also no difference in the expression levels of myogenin in both normal and dystrophic myotubes. However, AAV-treated dystrophic myotubes had significantly reduced levels of myogenin (Fig. 4, A and B). The protein levels of CaN, myomaker, and Cav3 were not significantly different between normal and dystrophic myotubes. Furthermore, AAV treatment had no effect on the expression levels of these proteins (Fig. 4A). There was a slight decrease in the protein levels of Stab2 in dystrophic myotubes, but this decrease was not statistically different from normal myotubes. AAV treatment increased the Stab2 expression in normal, but not dystrophic, myotubes (Fig. 4, A and B).

We next examined the effect of SLN reduction on myotube formation. Figure 4C shows the myosin-stained normal and dystrophic myotubes after 4 days of differentiation. Unlike immortalized mouse myoblasts, the primary myoblasts from normal dog produced very thin but multinucleated and branched myotubes, whereas many of the myotubes derived from dystrophic myoblasts were thin and bi- or trinucleated. Furthermore, dystrophic myotubes were typically less branched than normal myotubes. Reduction in SLN expression in dystrophic myoblasts by AAV treatment, on the other hand, resulted in an increased number of multinucleated and highly branched myotubes like that of normal myotubes. The fusion index quantitated by calculating the percentage of nuclei present in myosin-positive myotubes with myotubes containing >3 nuclei showed approximately threefold increase in fusion in AAV-treated dystrophic myotubes (Fig. 4D). AAV treatment had no effect on the differentiation of normal myoblasts.

Reduction in SLN expression improves autophagy in dystrophic myotubes.

Autophagy and mitophagy are shown to play an important role during muscle differentiation. Autophagy is also severely impaired in muscles of patients with DMD and in animal models (12, 33). We therefore first examined whether autophagy is impaired in dystrophic myotubes. Western blot analysis shows that microtubule-associated protein 1A/1B-light chain 3-II (LC3II), often used as a marker for autophagy, was significantly decreased in dystrophic myotubes (Fig. 5A). Treatment with chloroquine, a lysosomal inhibitor that blocks the autophagosome degradation, significantly increased the LC3II levels in both normal and dystrophic myotubes. However, this increase was significantly higher (P < 0.05) in chloroquine-treated normal myotubes compared with that of chloroquine-treated dystrophic myotubes (Fig. 5A). The autophagy receptor protein sequestosome 1 (SQSTM1), which acts as a substrate during autophagic degradation, was increased in dystrophic myotubes (Fig. 5B). These findings validate the autophagic defect in dystrophin-deficient cells. We next determined the effect of SLN reduction on autophagy. Western blot analysis shows that upon AAV treatment, the LC3II level was significantly increased in dystrophic myotubes (Fig. 5, B and C). LC3II levels were unaltered between the control and AAV-treated normal myotubes upon chloroquine treatment, whereas chloroquine treatment significantly increased the LC3II levels in AAV-treated dystrophic myotubes (1.4 ± 0.2-fold vs. control; n = 4; P < 0.05) compared with that of control dystrophic myotubes (Fig. 5D). In addition, AAV treatment significantly reduced the SQSTM1 protein levels in dystrophic myotubes (Fig. 5, B and E). Taken together, these findings suggest that reduction in SLN expression improved the autophagic flux in dystrophic myotubes.

Fig. 5.

Reduction in sarcolipin (SLN) expression restores autophagy in differentiated dystrophic myotubes. A: representative Western blots showing the microtubule-associated protein 1A/1B-light chain 3-II (LC3II) levels in normal and dystrophic myotubes with or without chloroquine treatment. Quantitation showing that the LC3II-to-LC3I ratio was significantly reduced in dystrophic myotubes after 4 days of differentiation. Chloroquine treatment (Chloro) significantly increased the LC3II levels and the LC3II-to-LCI ratio in normal (N) and Duchenne muscular dystrophy (DMD) myotubes; n = 4 samples per group. **P < 0.005 vs. respective untreated group. B: representative Western blots showing the LC3II, sequestosome 1 (SQSTM1), mitofusin 2 (Mfn2), dynamin-related protein-1 (Drp1), and phospho-Drp1 (pDrp1; S616) protein levels in normal myoblasts, normal myoblasts treated with adeno-associated virus expressing short hairpin RNA specific for canine SLN (AAV.cshSLN; N/shSLN), DMD (D) myoblasts, and DMD myoblasts treated with AAV.cshSLN (D/shSLN) after 4 days of differentiation. C: quantitation showing that the LC3II-to-LC3I ratio is significantly reduced in differentiated DMD myoblasts, whereas AAV.cshSLN treatment restores these changes; n = 4 samples per group. Statistical significance was determined by unpaired Student’s t test with Welch’s correction. *P < 0.05 vs. all other groups. D: representative Western blots showing LC3I and LC3II levels in normal, normal/shSLN, DMD, and DMD/shSLN myotubes with or without chloroquine treatment. E–G: box-and-whisker plots showing the SQSTM1, Mfn2, and Drp1 protein levels as fold changes in control and AAV.cshSLN-infected normal and DMD myoblasts differentiated for 4 days; n = 4 samples per group. Statistical significance was determined by unpaired Student’s t test with Welch’s correction. *P < 0.05 vs. N and N/shSLN groups. H: representative Western blots showing the protein levels of oxidative phosphorylation complex I (CI) through complex V (CV) subunits in control and AAV.cshSLN-infected primary myoblasts from normal and DMD dogs after 4 days of differentiation; n = 4 samples per group. I: oxygen consumption rate (OCR) obtained from normal, normal/shSLN, DMD, and DMD/shSLN myoblasts after 4 days of differentiation; n = 5 experiments per group. ATP5A, ATP synthase subunit-α; MTCO1, cytochrome c oxidase subunit I; NDUFB8, NADH-ubiquinone oxidoreductase 1 β-subcomplex 8; SDHB, succinate dehydrogenase complex, subunit B, iron sulfur protein; UQCRC2, ubiquinol-cytochrome c reductase core protein II. NS, not statistically significant.

We next determined whether mitophagy is affected in dystrophic dog myotubes. The proteins involved in mitochondrial fission and fusion such as mitofusin 2 (Mfn2), dynamin-related protein-1 (Drp1), and Drp1 phosphorylated at serine 616 (activated Drp1) were significantly increased in dystrophic myotubes compared with that of normal myotubes, whereas AAV treatment reduced the expression levels of these proteins near to normal in dystrophic myotubes (Fig. 5, B, F, and G). In normal myotubes, AAV treatment significantly increased the Mfn2 and Drp1 protein levels.

We next determined the mitochondrial function in differentiated dog myotubes by measuring the levels of oxidative phosphorylation proteins using the OXPHOS Western blot antibody cocktail kit and the basal OCR by Seahorse analysis. Results show that the expression levels of representative proteins from complex I through complex V (Fig. 5H) and the OCR (Fig. 5I) were not different between normal and dystrophic myotubes. In addition, AAV treatment had no effect on the expression levels of these proteins or the OCR in both normal and dystrophic myotubes.

DISCUSSION

Muscle degeneration and improper muscle regeneration are the major contributors of muscle wasting in DMD. Therefore, better understanding of mechanisms that cause these conditions would allow us to design novel therapeutic strategies for DMD. In this study, using C₂C₁₂, a mouse myoblast cell line, and primary myoblasts from a dog model of DMD, we investigated the role of SLN in myogenic differentiation. The key findings demonstrated that SLN upregulation is one of the early molecular changes in dystrophin-deficient myoblasts, which causes defects in ${\text{Ca}}_{\text{i}}^{\text{2+}}$ cycling and subsequently contributes to defects in myogenic differentiation. Reduction in SLN expression restores these changes and improves dystrophic myoblast differentiation.

The present study on C₂C₁₂ myoblast differentiation provides direct evidence for the role played by SLN in myoblast fusion during differentiation. Overexpression of SLN in C₂C₁₂ leads to increased expression of SERCA2a and CSQ. These changes could be compensatory alterations to improve the SR Ca²⁺ uptake in the presence of high levels of SLN. Despite these changes, SLN overexpression resulted in decreased SR Ca²⁺ uptake, reduced SR Ca²⁺ load, and reduced ${\text{Ca}}_{\text{i}}^{\text{2+}}$ transient amplitude. Most remarkably, SLN overexpression impaired the fusion of myoblasts during differentiation as evident from the formation of thin myotubes with fewer nuclei. Together, these findings suggest that SLN upregulation in mouse C₂C₁₂ myoblasts could contribute to poor muscle differentiation and regeneration possibly via chronic SERCA inhibition and abnormal ${\text{Ca}}_{\text{i}}^{\text{2+}}$ cycling.

Although the above findings provide direct evidence for the role played by SLN in muscle differentiation, unlike dystrophic myoblasts, C₂C₁₂ cells express dystrophin. Therefore, we tested our hypothesis in dystrophin-deficient myoblasts from large mammals. Unlike mouse myoblasts, normal dog myoblasts express SLN. In addition, SLN expression is significantly increased both in myoblasts and differentiated myotubes from the DMD dog. These findings indicate that SLN upregulation is an intrinsic secondary change in dystrophin-deficient muscles. The increased duration of twitch transient (T₅₀), decreased fractional Ca²⁺ release, and increased SR Ca²⁺ content suggest that the SR Ca²⁺ uptake and release mechanisms are impaired in the dystrophic dog myotubes. The decreased SERCA1 levels and increased SLN expression could contribute to the defect in SR Ca²⁺ uptake, which could subsequently affect the SR Ca²⁺ release. Reduction in SLN expression restored these changes in dystrophic myotubes. The most important finding is that reduction in SLN expression helped facilitate dystrophic myoblast fusion during differentiation as evident from the formation of more-branched myotubes. All these findings are consistent with our previous data, which demonstrate that normalization of SLN expression is sufficient to improve the SR function and muscle regeneration in a mouse model of DMD (55).

Myogenin plays a critical role in the maintenance and regeneration of adult skeletal muscles (20). Myogenin is not expressed in proliferating myoblasts, but it is expressed early in the terminal differentiation process (14). In C₂C₁₂, consistent with an earlier report (35), myogenin expression peaks during early differentiation and then starts to decline (Supplemental Fig. S1, A and B). On the other hand, myogenin levels are very high in SLN-overexpressing C₂C₁₂ cells, even 7 days after induction of differentiation, indicating that these cells are in the early stages of differentiation (Supplemental Fig. S1B). On the other hand, in dystrophic dog myotubes, reduction in SLN expression is associated with decreased levels of myogenin. Consistent with these findings, reduction or ablation of SLN expression reduced the myogenin expression in muscles of mdx mice (Supplemental Fig. S5). It has also been demonstrated that reducing myogenin expression could partially restore muscle function in DMD (30). At present we do not know the mechanism by which SLN levels modulate myogenin expression in dystrophic muscles. Future studies are needed to explore these mechanisms.

Cav3 is unaltered in dystrophic dog myotubes, and reduction in SLN expression has no effect on Cav3 expression. This is in contrast to reports that show that Cav3 is upregulated in dystrophin-deficient skeletal muscles of mdx mice (26, 54) and in the quadriceps of patients with DMD (39). It has also been demonstrated that overexpression of Cav3 in skeletal muscle fibers is sufficient to induce a Duchenne-like muscular dystrophy phenotype in mice (56). In addition, overexpression of Cav3 inhibits myoblast fusion to multinucleated myotubes, and lack of Cav3 enhances this fusion process (56). Consistent with these reports, SLN overexpression in C₂C₁₂ is associated with Cav3 upregulation and poor differentiation. Thus, Cav3 levels could contribute to the defects in mouse myoblast differentiation, but not in dystrophic dog myoblast differentiation.

Activation of the CaN-NFAT pathway has been shown to play an important role in muscle fusion (24, 25, 35, 37). The decreased expression of CaN in SLN-overexpressing C₂C₁₂ thus suggests that the decreased CaN activity could contribute to the defect in C₂C₁₂ myoblast fusion. However, in dystrophic dog myotubes, reduction in SLN expression has no effect on the expression levels of CaN or Stab2, another protein shown to be involved in myoblast fusion. This is in contrast to findings that show reduced levels of CaN and Stab2 in the muscles of mdx mice on SLN-null background (13). At present, we did not find a possible explanation for the discrepancy in CaN expression and SLN levels, although in the present study we did not completely ablate SLN expression in dystrophic dog myotubes. It is also important to note that unlike in rodents, in large mammals, SLN is expressed in all muscles (3), and its expression is increased in DMD (55). Furthermore, it is possible to speculate on the likely possible difference in myogenic differentiation during DMD pathogenesis in rodents and in higher mammals.

Autophagy and mitophagy play important roles in myogenic differentiation (50). A recent study shows that the maintenance of basal level autophagy is critical for myogenesis, particularly for the fusion of myoblasts (16). During myogenic differentiation, mitochondrial function is dramatically increased to meet the metabolic shift from glycolysis to oxidative phosphorylation. To meet this energetic demand, mitochondria undergo fission and fusion cycles along with autophagy to remove the damaged mitochondria and restore them with healthy tubular mitochondria (50). We found that in dystrophic dog myotubes, mitochondrial fission and fusion are increased, whereas autophagy is decreased. Our study also corroborates the decreased autophagy (12, 33) and increased levels of mitochondrial fission and fusion proteins (34, 44) reported in muscles of DMD models. In dystrophin-deficient dog myotubes, the decreased autophagy could cause more frequent cycles of mitochondrial fusion and fission as an adaptation to meet metabolic needs during differentiation. This is also evident from the unaffected electron transport chain complexes as well as the unaltered respiratory capacity seen in the dystrophic dog myotubes. A recent study from Periasamy’s group has demonstrated that SLN expression activates mitochondrial biogenesis and oxidative metabolism in muscles via Ca²⁺-dependent signaling pathways (28). They have also shown that SLN overexpression improved muscle performance by increasing oxidative capacity (52). On the other hand, metabolic dysfunction and altered mitochondrial dynamics are present in muscles of mdx:utr^−/− mice (34), where SLN is significantly upregulated (43, 55). Thus, SLN upregulation could be rather detrimental and not beneficial to dystrophin-deficient muscles. In support of this notion, normalizing SLN expression restored autophagy and mitochondrial dynamics in dystrophic dog myotubes (present findings) and also improved muscle function and ameliorated severe muscular dystrophy in mouse models of DMD (27, 55). Taken together, our studies suggest that SLN upregulation in dystrophin-deficient myoblasts could be an upstream pathological event causing defects in autophagy and mitophagy and thereby contributes to muscle pathophysiology.

In summary, the present findings implicate a key and previously unknown mechanism that links SLN upregulation to poor or improper myogenic differentiation. Although the exact signaling mechanism by which SLN overexpression interferes with muscle differentiation is yet to be identified, our findings suggest that reduction in SLN expression can improve myoblast fusion, a key process for muscle regeneration in DMD. Consistent with these observations, we found improved muscle regeneration in mdx:utr^−/− mice deficient in SLN (55). Therefore, reducing SLN expression may represent a novel therapeutic strategy for improving muscle structure and function in DMD.

GRANTS

This work was supported by National Institute of Arthritis and Musculoskeletal and Skin Diseases (NIAMS) Grant 1R01AR069107 (to G. J. Babu) and partially supported by NIAMS Grant 1R01AR070517 (to D. Duan and G. J. Babu), Jesse’s Journey-The Foundation for Gene and Cell Therapy (to D. Duan and G. J. Babu), and Jackson Freel DMD Research Fund (to D. Duan).

DISCLAIMERS

The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

DISCLOSURES

D. Duan is a member of the scientific advisory board for Solid Biosciences and equity holders of Solid Biosciences. The Duan laboratory has received research supports unrelated to this project from Solid Biosciences.

AUTHOR CONTRIBUTIONS

G.J.B. conceived and designed research; N.N., S.M., Y.T., K.K., N.F., and A.V. performed experiments; N.N., S.M., L.-H.X., and G.J.B. analyzed data; N.N., S.M., D.D., and G.J.B. interpreted results of experiments; N.N. and G.J.B. prepared figures; N.N., S.M., and G.J.B. drafted manuscript; K.K., L.-H.X., D.D., and G.J.B. edited and revised manuscript; N.N., S.M., Y.T., K.K., N.F., A.V., L.-H.X., D.D., and G.J.B. approved final version of manuscript.