- Open Access
Loss of niche-satellite cell interactions in syndecan-3 null mice alters muscle progenitor cell homeostasis improving muscle regeneration
© The Author(s). 2016
- Received: 20 September 2015
- Accepted: 26 August 2016
- Published: 4 October 2016
The skeletal muscle stem cell niche provides an environment that maintains quiescent satellite cells, required for skeletal muscle homeostasis and regeneration. Syndecan-3, a transmembrane proteoglycan expressed in satellite cells, supports communication with the niche, providing cell interactions and signals to maintain quiescent satellite cells.
Syndecan-3 ablation unexpectedly improves regeneration in repeatedly injured muscle and in dystrophic mice, accompanied by the persistence of sublaminar and interstitial, proliferating myoblasts. Additionally, muscle aging is improved in syndecan-3 null mice. Since syndecan-3 null myofiber-associated satellite cells downregulate Pax7 and migrate away from the niche more readily than wild type cells, syxndecan-3 appears to regulate satellite cell homeostasis and satellite cell homing to the niche.
Manipulating syndecan-3 provides a promising target for development of therapies to enhance muscle regeneration in muscular dystrophies and in aged muscle.
- Satellite cells
- Muscle regeneration
- Muscular dystrophy
- Cell adhesion
- Cell migration
Muscular dystrophy is a family of genetic disorders characterized by progressive muscle loss, chronic inflammation and replacement of muscle tissue with fibrotic tissue [1, 2]. In several types of muscular dystrophy, the continuous myofiber damage caused by the primary genetic defect imposes a high demand for myofiber repair, which is sustained by muscle progenitors. The proliferative potential of the resident muscle progenitors, named satellite cells, is presumed to be prematurely exhausted in muscular dystrophy, abrogating muscle regeneration and leading to fibrosis [3–7]. Although a thorough understanding of the molecular mechanisms regulating satellite cells in muscular dystrophy is incomplete, cell-intrinsic mechanisms, such as telomerase expression , cell cycle regulators , and cell-intrinsic disruptions of self-renewal and cell division upon dystrophin loss from satellite cells  as well as non cell autonomous regulation, including the extracellular environment [11–14], play critical roles in regulating satellite cell homeostasis.
During aging, myofiber size progressively decreases with an accompanying loss of fast twitch myofibers, leading to reduced overall muscle mass and strength that, when severe, results in sarcopenia. Loss of muscle mass and strength is accompanied by increased matrix deposition (fibrosis) and increased fat infiltration. Skeletal muscle regeneration is impaired in aged muscle and associated with cell-intrinsic deficits in satellite cell function [15–20]; however, satellite cell contribution to sarcopenia has been recently questioned, although a contribution of satellite cell loss to aging-associated fibrosis is supported .
Satellite cells in G0 phase reside within the musculature and are poised to rapidly activate in response to injury [22–26]. Upon activation, satellite cells re-enter the cell cycle, migrate away from their niche, and proliferate as myoblasts, eventually undergoing terminal differentiation into myocytes that fuse into pre-existing damaged muscle fibers or fuse to one another generating new muscle fibers . During regeneration, a portion of satellite cells returns to its niche, re-enters quiescence, and expresses Pax7 but no other myogenic transcription factors [27–29]. The transmembrane heparan sulfate proteoglycan syndecan-3, a component of the satellite cell niche, controls satellite cell homeostasis by regulating signaling pathways within the niche [12, 14, 30–32]. Moreover, members of the Syndecan family regulate cell-cell adhesion and cell-matrix adhesion via interaction with integrins and cadherins . Following a muscle injury, syndecan-3 null (Sdc3 −/− ) satellite cells fail to replenish the resident pool of quiescent satellite cells within the niche  and therefore syndecan-3 appears to regulate satellite cell homeostasis .
We show that syndecan-3 loss alters satellite cell adhesion to the myofiber, altering interactions with the niche and (i) improves muscle regeneration upon repeated acute muscle injuries, (ii) rescues muscle histopathology and function in dystrophic muscle tissue, and (iii) improves muscle aging with a reduction in fibrosis. The lifelong improvement in muscle regeneration observed in Sdc3 −/− muscle arises in part by altered satellite cell homeostasis and changes in satellite cell adhesiveness to the myofiber.
Mice were housed in a pathogen-free facility at the University of Colorado at Boulder, USA, or at the University of Liverpool, UK. All injuries and other procedures were performed at the University of Colorado, and protocols were approved by the IACUC at the University of Colorado. Animals housed at the University of Liverpool were used in accordance with the Animals (Scientific Procedures) Act 1986 and the EU Directive 2010/63/EU and after local ethical review and approval by Liverpool University’s Animal Welfare and Ethical Review Body (AWERB). Sdc3 −/− mice were donated by Dr. Heikki Rauvala, University of Helsinki, Finland. Mdx 4cv mice were donated by Dr. Jeffrey Chamberlain, University of Washington, Seattle, USA. Generation of double mutant colonies is described in details in Additional file 1. In all experiments, wild type and mdx 4cv ;Sdc3 +/+ controls were all siblings or closely related, inbred, sex- and age-matched animals for all transgenic lines.
Tissue samples were collected and either immediately frozen in liquid nitrogen-cooled isopentane or fixed in 10 % formalin. For all immunofluorescence staining except Myf5 and Pax7, sections were fixed with 4 % paraformaldehyde (PFA) in phosphate buffered saline (PBS) for 10 min at room temperature. For Myf5 staining, sections were fixed for 10 min with acetone at −20 °C. For Pax7 staining, sections were either fixed and stained using an anti-Pax7 rabbit polyclonal antibody (Genetex) or non fixed, processed for antigen retrieval, and stained with an anti-Pax7 mouse monoclonal antibody (DSHB). The antibodies used were as follows: rabbit polyclonal anti-Pax7 (Genetex) at 1:250; rabbit polyclonal anti-laminin (Sigma) 1:150; rat polyclonal anti-laminin α2 (Sigma) 1:100; rat anti-F4/80 (Genetex) 1:200; rat anti-BrdU (Serotec) 1:100; mouse anti-Pax7 monoclonal (DSHB) 1:200; rabbit anti-myogenin (SCBT) 1:50; rabbit anti Myf5 (SCBT) 1:200; rat anti-CD31 (BD Biosciences) 1:100; rabbit anti-NG2 (Chemicon) 1:200; rabbit anti-Ki67 (Abcam) 1:400; rat anti-Sca1 (unconjugated, PE-conjugated, APC-Cy7-conjugated and FITC-conjugated were all from BD Biosciences), 1:100; rabbit anti-GFP (BD Biosciences), 1:400. Secondary antibodies conjugated with Alexa594, Alexa555, Alexa488, or Alexa647 (Molecular Probes) were used at 1:500 dilution. Vectashield with DAPI (Vector Laboratories) was used as a mounting medium.
Sirius red staining
Flash-frozen sections were fixed for 1 h at 56 °C in Bouin’s fixative, washed in water, stained for 1 h in Master*Tech Picro Sirius Red, washed in 0.5 % acetic acid, dehydrated, equilibrated with xylene, and mounted using Permount™.
Trichrome staining was performed according to standard protocols by Premier Laboratory LLC, Boulder, CO, on paraffin-embedded tissues fixed in 10 % formalin in neutral buffered saline and preserved in 70 % ethanol.
Myofiber cross-sectional area and numbers in uninjured and injured TA muscles were quantified as previously described . The fibrotic index (% collagen + area in Sirius Red staining relative to total section area) was quantified by selecting red pixels in Adobe Photoshop, deleting all non-red pixels, converting the resulting image to a binary image, and counting red pixels using the ImageJ Analyze Particles function. The necrotic index was calculated by counting the number of mIgG+ myofibers and normalizing to total number of myofibers in the image. Capillary density was calculated by measuring the numbers of capillary around each fiber on alternate fibers in order to avoid overlapping scorings. Ten sections per mouse for three different mice were scored.
Female and male mice of different genotypes were individually housed in cages equipped with a training wheel connected to a bicycle computer (Schwinn) with ad libitum access to food and water for 3 weeks. Time and distance run were recorded daily.
Mice were anesthetized with 2,2,2-tribromoethanol (Sigma) such that they were insensitive to tactile stimuli. Peak isometric force of the TA muscle was analyzed in situ via nerve stimulation. First, we found the maximum force-producing capacity of each muscle at its optimum length according to maximal stimulation over 300 ms to elicit tetanic contraction. The peak force was then divided by the unit area of muscle to obtain specific force (kN/m2) using the equation: specific force = peak force × muscle length × 0.6 × 1.04/muscle weight . Next, we measured protection from contraction-induced injury. The force-producing capacity of the muscle was measured immediately prior to increased length changes during maximal stimulation at 20-s intervals. Length changes were increased in 5 % increments from 5 to 45 % of muscle fiber length to produce injury. The rate of length change was 2 lengths/s.
Quadriceps were homogenized in 20 mM HEPES, 50 mM KCl, 1 mM DTT, 2 mM MgCl2, 0.5 mM EDTA, 0.5 % NP40 supplemented with protease inhibitor cocktail (Complete, Roche), and phosphatase inhibitors (1 mM Na3VO4 + 1 mM NaF) using an UltraTurrex homogenizer followed by incubation on ice for 20 min and then cleared by centrifugation at 13,000 rpm for 10 min at 4 °C. Western blot was performed as previously described . The antibodies used were as follows: rabbit polyclonal anti-dystrophin (Abcam) at 1:1000; rabbit polyclonal anti-utrophin (kindly donated by Dr. Froehner, University of Washington, Seattle) 1:2000. Anti-rabbit conjugated secondary antibodies (Santa Cruz) were used at 1:10,000, and HRP activity was visualized using the ECL plus system (Amersham).
Dystrophin forward: CAGCTGCAGAACAGGAGTT.
Dystrophin reverse: GCATCTACTGTGTGAGGACC.
Mice were anesthetized with isofluorane and the right TA muscle was injected with 50 μL of 1.2 % BaCl2  in three places along the length of the muscle and then both the injured muscle and the contralateral uninjured muscle harvested at the indicated time points. For repeated injuries, the same TA muscle was injured as above for a total of three times with 3-week intervals between injuries. The injured TA muscle and the contralateral uninjured TA muscle were harvested 3 weeks after the last injury.
Fluorescence-activated cell sorting
Hindlimb muscles of 3–6-month-old Sdc3 −/− and littermate wild type female mice were dissected, minced, and digested in 400 U/mL collagenase type I in Ham’s F-12C (F12 + 0.4 mM CaCl2) at 3 °C for 1 h, gently vortexing every 10 min. Collagenase was diluted 1:3 with F12C + 15 % horse serum (HS) and tissue debris removed by centrifugation at 30×g for 5 min (pellet contains large debris) followed by straining of the supernatant (containing mononucleated cells and smaller debris) through 40-μm cell strainers (BD Falcon). Flow through was then centrifuged at 300×g, and the cell pellets were re-suspended in PBS + 5 % fetal bovine serum (FBS) and incubated for 45 min at 4 °C with 1:100 phycoerythrin (PE)- or fluorescein isothiocyanate (FITC)-directly conjugated rat anti-Sca1 antibody (BD Biosciences) and 1:500 chicken anti-Sdc4  followed by an incubation for 45 min at 4 °C with Alexa 647-conjugated anti-chicken IgY (Molecular Probes). Sca1+, Sca1+/Sdc4−, and Sca1−/Sdc4+ cells were sorted on a MoFlo XDP Cell Sorter (Dako Cytomation) into Ham’s F12C + 15 % horse serum (HS) and cultured in a myogenic growth medium (F12C + 15 % HS + 2 nM FGF2) or transplanted, see transplantation details below. To assess the expression of Pax7, Pax3, and Myf5 and MyoD, fibro-adipogenic progenitors (FAPs) were sorted as Hoechstmid PIlo CD45− CD31− Sca1+ CD34+ cells and muscle progenitors (MPs) were sorted as Hoechstmid PIlo CD45− CD31− Sca1− CD34+ as previously described  directly into lysis buffer (CellsDirect Resuspension & Lysis Buffer, Life Technologies).
Droplet Digital PCR
Following RNA isolation (CellsDirect Resuspension & Lysis Buffer, Life Technologies) and reverse transcription (High Capacity cDNA Reverse Transcription Kit, Life Technologies) according to the manufacturer’s instructions, complementary DNA (cDNA) was diluted five times in TE buffer and 5 μL were used in a reaction mix containing Droplet Digital™ PCR Supermix (BioRad), 1× TaqMan probes from Life Technologies [Pax7 (Mm03053796-s1), Myf5 (Mm00435125-m1), Hprt (Mm00446968_m1), Pax3 (Mm00435493_m1), and Myod1 (Mm00440387_m1)] and H2O. Droplets were generated with a QX100 droplet generator (BioRad), after mixing 20 μL of reaction mix and 70 μL of droplet generator oil (BioRad). The emulsified samples were loaded onto 96-well plates, and endpoint PCRs were performed in C1000 Touch thermal cycler (BioRad) at the following cycling conditions (95 °C for 10 min, followed by 45 cycles of 94 °C for 30 s and 60 °C for 1 min, followed by 98 °C for 10 min). The droplets from each sample were read through the QX100 droplet reader (BioRad). Resulting PCR-positive and PCR-negative droplets were counted using QuantaSoft software (BioRad). Expression levels were normalized to Hprt.
Sca1+ cells were FACS-isolated as described above from Sdc3 +/+ ;β-actin-GFP and Sdc3 −/− ;β-actin-GFP mice, centrifuged, and washed twice with sterile 0.9 % NaCl to remove the serum, re-suspended into 0.9 % NaCl at the concentration of 2400 cells/μL and 30 μL (~70,000 cells) immediately injected into the right TA muscle of wild type mice which had been injured 4 h before with an injection of 30 μL of 1.2 % BaCl2. Three weeks after animals were sacrificed, the right (injured and transplanted) and left (uninjured, untransplanted) TA muscles were dissected and cryopreserved for subsequent histological analysis.
Myofiber isolation and culture
The gastrocnemius muscles of wild type and Sdc3 −/− mice were dissected and incubated with 400 U/mL collagenase type I in F12C at 37 °C, with gentle mixing by inversion every 15 min for 1 h 30 min, after which collagenase was diluted 1:5 with F12C + 15 % HS and muscles gently rocked at room temperature for 15 min to allow for myofiber release from the digested muscle. Individual myofibers were manually picked and transferred to fresh F12C + 15 % HS using a sterile, flame-polished Pasteur pipette. Myofibers were cultured in suspension in F12C + 15 % HS + 2 nM FGF2 in non-coated sterile petri dishes unless otherwise specified and transferred every 24 h to fresh medium.
Microscopy, image processing, and figure preparation
Micrographs were taken with a Leica TCS SP2 AOBS confocal microscope using dedicated Leica software, or with a Nikon (Eclipse E800) epifluorescence microscope using Slidebook v4.1 acquisition software (Intelligent Imaging Innovations Inc.) coupled to a Cooke Sensicam digital camera or with an EVOS-FL inverted microscope (Life Technologies). Lenses used with the Leica confocal microscope were either HC PL APO 20×/0.70 IMM CORR CS or HCX PL APO 40×/1.25–0.75. Lenses used with the Nikon Eclipse microscope were Nikon Plan Fluor either 40×/0.75 DIC M or 20×/0.50 Ph1 DLL. Lenses used with the EVOS microscope were PL FL, either 10× LWD PH, 0.25NA/9.2WD or 40× LWD PH, 0.56NA/1.6WD. All digital microscopic images were acquired at room temperature. For figure preparation, images were exported in Adobe Photoshop, if necessary brightness and contrast adjusted and the background removed for the entire image, the image cropped and individual color channels extracted (when required) without color correction or gamma adjustments.
To assess statistical significance, two-tailed, unpaired Student’s t test or one-way analysis of variance (ANOVA) were performed. p < 0.05 was considered significant. At least three different animals per genotype and per age group were used in all experiments. Cell culture experiments (both myofiber and myoblast cell cultures) were repeated three independent times using three different animals per genotype group. For force measurements, five to seven animals per genotype were used. For muscle function testing (voluntary wheel), three to seven animals per genotype group were used.
Dystrophic mice lacking syndecan-3 show improved muscle histopathology and function
Improved muscle function in syndecan-3 null dystrophic mice accompanied the improved muscle histopathology as compared to mdx 4cv ;Sdc3 +/+ mice. Both male and female mdx 4cv ;Sdc3 −/− mice ran for longer distances and for longer periods of time than mdx 4cv ;Sdc3 +/+ controls when assayed on a voluntary wheel, performing similar to the times and distances recorded for wild type mice (Fig. 1g, j). This was not due to an intrinsically increased propensity of Sdc3 −/− mice to perform better in endurance training tests as no significant differences in time and distance run were recorded for Sdc3 −/− non-dystrophic mice compared to wild type mice. The diaphragm muscle, which was severely affected following voluntary running in mdx 4cv ;Sdc3 +/+ mice, was dramatically improved in mdx 4cv ;Sdc3 −/− mice following voluntary exercise (Fig. 1k). Amelioration of the dystrophic phenotype in mdx 4cv ;Sdc3 −/− mice was likely maintained throughout life as in 14-month-old (Additional file 1: Figure S1B, C) and 19-month-old (Additional file 1: Figure S1D, E) muscle, collagen deposition is reduced in mdx 4cv ;Sdc3 −/− compared to mdx 4cv ;Sdc3 +/+ mice (Additional file 1: Figure S1B-E).
Regeneration is improved in dystrophic muscle lacking syndecan-3
Contraction-induced injury and muscle force measurement are carried out on individual muscles and represent a measure of muscle performance prior to damage. If syndecan-3 loss improved exercise performance by improving myofiber integrity, then both contraction-induced injury and muscle force would be improved. Instead, we find the opposite: neither contraction-induced injury nor muscle force are improved in mdx 4cv ;Sdc3 −/− muscles. Therefore, loss of syndecan-3 in dystrophic muscle does not prevent myofiber rupture in response to stretch, yet the overall muscle histology is improved and is associated with an overall improvement in exercise performance. These apparently conflicting results could be explained if muscle regeneration was improved in mdx 4cv ;Sdc3 −/− enhancing muscle function, improving fatigue resistance during exercise and reducing fibrosis.
We asked if enhanced regeneration in syndecan-3 null dystrophic muscle ameliorates the dystrophic phenotype. We observed an increase in myofiber area (Fig. 2h) accompanied by increases in centrally located nuclei (Fig. 2i) and numbers of myofibers with two or more centrally located nuclei (Fig. 2j) in mdx 4cv ;Sdc3 −/− muscles compared to mdx 4cv ;Sdc3 +/+ muscles. These observations, together with our previous data that Sdc3 −/− satellite cells generate larger myotubes ex vivo  support the hypothesis that syndecan-3 loss enhances myofiber regeneration in chronically injured muscles by increasing muscle progenitor contribution to damaged myofibers.
Fibro-adipogenic progenitors (FAPs) can convert to myogenic progenitors in a dystrophic environment . To test whether myogenic conversion of FAPs was responsible for increased muscle regeneration and decreased fibrosis observed in mdx 4cv ;Sdc3 −/− mice, we isolated FAPs from mdx 4cv ;Sdc3 +/+ and mdx 4cv ;Sdc3 −/− mice and profiled them by qPCR for expression of myogenic markers. Although MyoD, Pax7, and Myf5 expression was detected in prospective satellite cells isolated from mdx 4cv ;Sdc3 +/+ or mdx 4cv ;Sdc3 −/− muscle, no MyoD, Pax7, Pax3, and Myf5 expression was detected in FAPs isolated from either mdx 4cv ;Sdc3 +/+ or mdx 4cv ;Sdc3 −/− muscle (Additional file 1: Figure S2). These results support the conclusion that in vivo conversion of FAPs to myogenic progenitors is negligible or absent in the mdx 4cv dystrophic background and is not enhanced by syndecan-3 loss.
Syndecan-3 loss improves regeneration repeatedly injured muscle and muscle aging
During aging, a progressive loss of satellite cells occurs via loss of satellite cell self-renewal [15, 16, 45], which is thought to contribute to age-associated muscle fibrosis . To determine if syndecan-3 loss affects fibrosis and muscle aging, we measured the levels of extracellular matrix deposition in 2-year-old wild type and Sdc3 −/− muscles. Although an occasional accumulation of lipid droplets was previously described in aged Sdc3 −/− muscle , a significant decrease in collagen was observed in aged Sdc3 −/− muscle compared to aged wild type muscle (Fig. 3e, f). Reduced muscle fibrosis in old Sdc3 −/− muscle was associated with increased numbers of myogenin + cells (Fig. 3g) and increased numbers of centrally nucleated fibers (Fig. 3h), suggesting that depletion of the pool of Pax7+ satellite cells upon activation in Sdc3 −/− mice does not exhaust muscle regenerative capacity. Instead, syndecan-3 loss is associated with improved muscle in aged mice and improved regeneration in repeatedly injured muscle and in dystrophic muscle.
Muscle progenitors distinct from satellite cells contribute minimally to muscle regeneration in the absence of syndecan-3
In addition to interstitial, non-myogenic cells, Sca1 is also expressed in a subpopulation of satellite cells marked by syndecan-4 [49, 51] and is induced upon satellite cell activation in a subpopulation of satellite cells that self-renew [52, 53]. Therefore, we asked if differences exist between wild type and Sdc3 −/− muscle-derived Sca1+/Sdc4− cells (Fig. 4g). When Sca1+/Sdc4− cells from wild type and Sdc3 −/− mice were isolated by FACS and cultured at clonal density, a higher percentage of myogenic Sca1+/Sdc4− clones was present in Sdc3 −/− muscle as opposed to wild type muscle (Fig. 4h).
Satellite cell homeostasis is altered in mice lacking syndecan-3
Total numbers of myogenic progenitors are increased in uninjured Sdc3 −/− muscle compared to wild type muscle
Maintenance of interstitial Myf5+ cells in Sdc3 −/− mice may reflect changes in Sdc3 −/− cell adhesion since syndecans are adhesion molecules. Sdc3 −/− myofiber-associated satellite cells appear less adhesive than wild type satellite cells (Additional file 1: Figure S4A-B). When isolated myofibers from wild type and Sdc3 −/− muscles were cultured in suspension and then transferred onto gelatin-coated dishes, twofold more Sdc3 −/− myoblasts adhered to the gelatin-coated surface than wild type myoblasts 4 h post-transfer (Fig. 5h, i and Additional file 1: Figure S4C). The propensity of Sdc3 −/− satellite cells to migrate away from their native niche is consistent with the finding that the majority of Myf5+ and Ki67+ cells observed in regenerated Sdc3 −/− muscle are located in the interstitial space, and supports the idea that the My5+ myoblast population observed in regenerated Sdc3 −/− muscle is derived from satellite cells that migrated away from their niche.
In adult wild type muscle, Pax7+ satellite cells are quiescent and indispensable for muscle regeneration [56, 57]; Pax7 is necessary to maintain this population [58, 59]. Satellite cell niche components including Notch, syndecan-4, integrin-α7, Wnt, FGFs, HGF, the calcitonin receptor, and fibronectin play critical roles in maintaining satellite cells in their niche [12, 14, 15, 27, 60–64]. Syndecan-3, a transmembrane proteoglycan expressed in satellite cells and involved in regulating satellite cell responses to growth factors and to Notch [12, 14, 30], appears to promote satellite cell identity, the association of satellite cells with their niche and satellite cell quiescence.
Since Sdc3 −/− satellite cells proliferate slowly and show increased rates of cell death due to a defect in Notch signaling , the process of myonuclear accretion is slow and in the short lifespan of a mouse does not lead to an appreciable increase in muscle size. Nonetheless, a significant increase in satellite cell contribution to myofibers, shown by the presence of centrally nucleated myofibers, accompanied by a significant reduction in muscle fibrosis, is observed in wild type or dystrophic aged mice lacking syndecan-3. Thus, syndecan-3 loss appears to provide a lifelong benefit to muscle regenerative capacity in mice.
Although other potential myogenic progenitors, such as pericytes, myoendothelial cells, and side population cells, which are increased in Sdc3 −/− muscle and show increased myogenicity in vitro, may contribute to interstitial and sublaminar myoblasts, the relative contribution of these cells appears low and may possibly be due to satellite cell contamination of the interstitial cell preparation. We cannot directly lineage trace the Myf5+ interstitial cells identified in regenerated Sdc3 −/− muscle due to (i) the close proximity of Pax7 and syndecan-3 on the same chromosome, (ii) the lower levels of Pax7 in Sdc3 −/− satellite cells, and (iii) the co-expression of MyoD and Myf5 by activated satellite cells and the interstitial myoblasts in Sdc3 −/− muscle.
Loss of muscle regenerative capacity in the muscular dystrophies is often attributed to satellite cell exhaustion [3–9, 71]; however, there are only few experiments directly supporting this hypothesis. We utilized mdx 4cv mice [37, 72], which develop a more severe form of muscular dystrophy than mdx mice that is exacerbated when challenged with exercise . The dystrophy becomes more severe as the mice age, presumably due to the lower numbers of revertant fibers in mdx 4cv mice than in mdx mice . Loss of syndecan-3 in dystrophic mice reduces muscle fibrosis while improving exercise performance without ameliorating myofiber fragility or increasing the specific force. Since myofiber damage appears equivalent in dystrophic muscle with or without syndecan-3, we postulate that muscle regeneration is enhanced, leading to improved exercise performance. This conclusion is supported by the finding that mdx 4cv ;Sdc3 −/− muscles contain more regenerating myofibers than mdx 4cv ;Sdc3 +/+ muscles and enhanced myonuclear accretion, consistent with a role for syndecan-3 in supporting Notch signals which promotes self-renewal while inhibiting myoblast fusion .
The Sdc3 −/− satellite cell phenotypes appear cell autonomous as they occur in culture as well as in dystrophic mice lacking syndecan-3 and in aged Sdc3 −/− mice. Overall the mechanism responsible for the enhancement of regeneration in double mutant mdx 4cv ;Sdc3 −/− mice, the amelioration of the dystrophic phenotype, and the improvement of muscle maintenance in aged mice appears to be the failure of Sdc3 −/− satellite cells to return to quiescence and re-home to their niche after activation, which maintains an expanding population of interstitial Myf5+ myoblasts. The numbers of Sdc3 −/− myoblasts increase over time leading to an expanded muscle progenitor population in the muscle interstitium that eventually generates large, centrally nucleated myofibers (Fig. 6).
Sdc3 −/− mice maintain lifelong muscle regenerative capacity and resist injury-induced loss of regenerative capacity by maintaining a population of activated, Myf5+Pax7− satellite cells and a proliferating myoblast population in the myofiber interstitium. Sdc3 −/− satellite cells do not appear exhausted in either dystrophic muscle or aged muscle apparently enhancing muscle regenerative capacity, identifying a new potential therapeutic target for the treatment and management of muscular dystrophies, repeated acute injuries and muscle aging.
We thank Dr. Jeffrey Chamberlain for the mdx 4cv mice and Dr. Heikki Rauvala for the Sdc3 −/− mice. We also thank Dr. Michelle Doyle for critical reading of the manuscript and insightful discussions; Dr. Malea Murphy for help with development of histology techniques and automated image quantification and Ms. Tiffany Antwine for technical help with histology and flow cytometry. This work was supported by the MDA, The Ellison Medical Foundation, and NIH Grants AR049446 and AG040074 to BBO, by a Wellcome Trust ISSF and a Marie Curie IEF to AP, an MDA development grant to GBB, CIHR grant MOP-97856 to FMVR, and a 4YF fellowship from UBC to FB.
AP designed and performed all experiments except those shown in Figs. 2a, c, d–f, 4d, e, and 5a, Additional file 1: Figure S1B and D. AP also carried out data analysis and drafted the manuscript. GBB and JSC designed and performed the experiments shown in Fig. 2a, d, e and contributed to the manuscript preparation. FB and FMVR designed and performed the experiments shown in Fig. 4d, e and Additional file 1: Figure S2. BBO participated in experimental design and in drafting of the manuscript. All authors read and approved the final manuscript.
The authors declare that they have no competing interests.
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