Myostatin genetic inactivation inhibits myogenesis by muscle-derived stem cells in vitro but not when implanted in the mdx mouse muscle
© Tsao et al.; licensee BioMed Central Ltd. 2013
Received: 12 July 2012
Accepted: 3 January 2013
Published: 7 January 2013
Stimulating the commitment of implanted dystrophin+ muscle-derived stem cells (MDSCs) into myogenic, as opposed to lipofibrogenic lineages, is a promising therapeutic strategy for Duchenne muscular dystrophy (DMD).
To examine whether counteracting myostatin, a negative regulator of muscle mass and a pro-lipofibrotic factor, would help this process, we compared the in vitro myogenic and fibrogenic capacity of MDSCs from wild-type (WT) and myostatin knockout (Mst KO) mice under various modulators, the expression of key stem cell and myogenic genes, and the capacity of these MDSCs to repair the injured gastrocnemius in aged dystrophic mdx mice with exacerbated lipofibrosis.
Surprisingly, the potent in vitro myotube formation by WT MDSCs was refractory to modulators of myostatin expression or activity, and the Mst KO MDSCs failed to form myotubes under various conditions, despite both MDSC expressing Oct 4 and various stem cell genes and differentiating into nonmyogenic lineages. The genetic inactivation of myostatin in MDSCs was associated with silencing of critical genes for early myogenesis (Actc1, Acta1, and MyoD). WT MDSCs implanted into the injured gastrocnemius of aged mdx mice significantly improved myofiber repair and reduced fat deposition and, to a lesser extent, fibrosis. In contrast to their in vitro behavior, Mst KO MDSCs in vivo also significantly improved myofiber repair, but had few effects on lipofibrotic degeneration.
Although WT MDSCs are very myogenic in culture and stimulate muscle repair after injury in the aged mdx mouse, myostatin genetic inactivation blocks myotube formation in vitro, but the myogenic capacity is recovered in vivo under the influence of the myostatin+ host-tissue environment, presumably by reactivation of key genes originally silenced in the Mst KO MDSCs.
The lipofibrotic degeneration of skeletal muscle (that is, excessive deposition of endomysial collagen, other extracellular matrix, and fat), characterizes muscle dystrophy, and in particular Duchenne muscular dystrophy (DMD) [1, 2], as seen also in its animal model, the mdx mouse [3–5]. This process, associated with inflammation and oxidative stress , is partially responsible for the severe muscle contractile dysfunction in DMD and the mdx mouse, caused mainly by the bouts of myofiber necrosis due to dystrophin genetic inactivation. In the gastrocnemius, these processes are rather mild in young animals but become particularly severe after 8 to 10 months of age . Dystrophic muscle fibrosis not only is a major factor for DMD mortality, but also hampers the uptake and survival of cells implanted for potential therapeutic approaches  and/or may drive their differentiation into myofibroblasts . Therefore, trying to ameliorate this process while stimulating myogenesis constitutes an ancillary strategy to favor repair and regeneration of dystrophic muscle tissue, even under ineffective or absent dystrophin replacement.
Although pharmacologic approaches to combat muscle lipofibrotic degeneration and the underlying chronic inflammation are being widely investigated, biologic factors such as myostatin, the main negative regulator of muscle mass , are also potential key targets. Myostatin, a member of the TGF-β family, aggravates muscle dystrophy not only as an antimyogenic agent but also as a profibrotic and adipogenic factor [9–14]. Inhibition of myostatin by using its propeptide, shRNA, or specific antibodies, improves myogenesis and reduces fibrosis in the mdx mouse [15–17] and also in the rat . The same effects are generated in response to genetic deletion of myostatin in the myostatin knockout (MST KO) mouse, in which myofiber hypertrophy is associated with less fat and reduced fibrosis [19–23].
It is assumed that in the dystrophic or injured muscle, tissue repair and the opposite process of lipofibrotic degeneration involve not only the differentiation of progenitor satellite cells and fibroblasts into myofibers and myofibroblasts, respectively, but also the modulation of lineage commitment by stem cells present in the adult muscle [24–26]. These stem cells have been isolated from the rodent and human skeletal muscle and named, in general, muscle-derived stem cells (MDSCs), because they have the ability to differentiate in vitro into multiple cell lineages and to generate myofibers, osteoblasts, cardiomyocytes, or smooth muscle cells after implantation into the skeletal muscle, bone, heart, corpora cavernosa, or vagina, respectively [27–31]. They are not satellite cells and may act also by secreting paracrine growth factors that are believed to modulate the differentiation of endogenous stem cells or the survival of differentiated cells in the tissue [32–34]. However, the roles of MDSCs in the biology and pathophysiology of the skeletal muscle are largely unknown.
Myostatin modulates the differentiation of pluripotent cells in vitro, albeit in some cases, with conflicting outcomes [14, 35–37]. It also inhibits the proliferation and early differentiation of both satellite cells from the skeletal muscle and cultured myoblasts, and blocking its expression improves the success of their in vivo transplantation [38–40]. To our knowledge, no reports are available on myostatin effects on MDSC differentiation, either in vitro or in the context of repairing the exacerbated lipofibrosis in the injured muscle of aged mdx mice.
MDSCs obtained from wild-type (WT) mice have been tested experimentally, aiming to trigger repair of the mdx muscle with variable results [31–45], but they appear to be superior in this respect to myoblasts or satellite cells . However, some of the main limitations of myoblast therapy, when translated from the murine models into DMD and other human muscle dystrophies, may also affect the MDSCs and other types of stem cells . Therefore, it is a therapeutic goal to enhance the repair capacity of WT MDSCs by in vitro or in vivo modulation of their multilineage potential, and to stimulate or even awake endogenous stem cells of dystrophic muscle to regenerate myofibers while avoiding differentiation into cells responsible for lipofibrotic degeneration. Such an approach may be provided by the use of MDSCs where myostatin is genetically inactivated (that is, obtained from the Mst KO mouse), under the assumption that myogenesis would be stimulated and the undesired lineage commitment reduced, even when implanted into a host tissue environment with normal myostatin expression. No reports are available on the in vitro and in vivo differentiation of these MDSCs and how this affects, even paracrinely, muscle repair.
Potential in vitro modulation of MDSCs, or the effects that myostatin or dystrophin gene inactivation exert on this balance
In the current study, we have investigated the in vitro myogenic versus fibrogenic and adipogenic differentiation of Mst KO MDSCs vis-à-vis the WT counterpart, and the effects of manipulation of these processes by modulating myostatin expression or activity, and by other putative regulators of muscle mass and fibrosis. Their differential in vitro features in terms of the expression of some key stem cell and myogenic genes, and the repair ability of Mst KO MDSCs in the injured mdx muscle, also were studied. The ultimate goal is to gain a preliminary insight into how in vitro preconditioning of MDSCs by pharmacologic or gain-of-function approaches may modulate their capacity to repair dystrophic skeletal muscle, to design in vivo pharmacologic interventions that may mimic these processes, and even myostatin blockade in the host muscle to activate myogenesis in the endogenous dystrophin-negative MDSCs.
Materials and methods
Mst knockout mice (C57BL/6J/Mst-/-), referred to here as Mst KO, are regularly maintained and bred in our vivarium on a BL/6 background , derived from the original strain on a Balb/c background. Aged-matched wild-type control mice (C57BL/6J), referred to here as WT, were from Jackson Laboratories (Bar Harbor, ME, USA). Hindlimb muscles from the WT and Mst KO male mice (12 to 16 weeks old) were subjected to the preplating procedure to isolate MDSCs [49, 50], by using a modification of a well-validated method that has led to extensively characterized stem cell populations [5, 27–30, 46, 49]. Tissues were dissociated by using sequentially collagenase XI, dispase II, and trypsin, and after filtration through 60-μm nylon mesh and pelleting, the cells were suspended in plating medium (PM), containing Dulbecco Modified Eagle Medium (DMEM), with 10% fetal bovine serum (FBS), 10% horse serum, and 0.5% chick embryo extract (US Biological, Marblehead, MA, USA). Cells were plated onto collagen I-coated flasks for 1 hour (preplate 1 or pP1), and 2 hours (preplate 2 for pP2), followed by sequential daily transfers of nonadherent cells and replatings for 2 to 6 days, until preplate 6 (pP6). The latter is the cell population containing MDSCs. Sca1+ cells were selected with immunobeads (Milteni, Auburn, CA, USA) coated with antibody against Sca1 as small cells with a large nucleus that easily form clusters/spheroids [24–27]. Cells were subjected to flow cytometry, as described later, for the MDSC standard markers Sca1, CD34, and CD44, and for the key stem-cell gene, Oct 4 , maintained in growth medium (GM) GM-20 (DMEM, with 20% FBS) on regular culture flasks (no coating) and used in passages 14 to 28. WT MDSCs have been maintained in our laboratory for at least 40 generations with the same, or even increasing, growth rate.
MDSC and KO cells were grown in GM-20, washed twice with Hanks, disaggregated by repeated pipetting in Cell Stripper (Mediatech, Manassas, VA, USA), pelleted, and resuspended in staining buffer consisting of PBS, 3% FBS, 0.01% Na azide (SB). Cells were incubated in the presence of antibodies for 30 minutes on ice, washed twice with SB, and finally resuspended in SB for flow cytometry on an LSR II (BD Biosciences, San Jose, CA, USA). Data analysis and plotting were done by using FACSDiva Version 6.1.1 software. All fluorophore-conjugated antibodies and isotype controls were from eBioscience (San Diego, CA, USA), as follows: CD44-APC-eFluor 780; CD34-eFluor 660; Sca1-PE; Oct 4-PE (performed separately, after cell permeabilization with BD CytoFix/CytoPerm Kit), and the appropriate rat isotype controls IgG2b-APC-eFluor 780, IgG2a-eFluor 660, and IgG2a-PE. BD CompBeads (rat) were used for compensation.
Stem cell characterization, differentiation, and modulation
MDSC cultures were analyzed for the expression of stem cell markers, as described later, on collagen-coated six-well plates and eight removable-chamber plates. Multipotency was analyzed in 2-week incubations with GM-20 or GM-10 (GM with 10% FBS) supplemented or not with 10 nM DMSO or 5 ng/ml TGF-β1, or, to induce myofiber formation, after reaching confluence, for 2 to 3 weeks with GM-HC (DMEM, 10% FBS, 5% horse serum, and 50 μm hydrocortisone to promote proliferation, a key event in myogenic differentiation) [44, 45], or as described. In certain cases, cultures were treated with or without 20 μM 5'-azacytidine (AZCT) in GM-20 for 3 days to induce multipotency, before switching them to the appropriate medium [11, 14, 51].
For the tests on the modulation of MDSCs skeletal myotube formation by various factors, cells were allowed to reach confluence, switched to GM-HC, and incubated for 2 weeks with 2 μg/ml recombinant 113-amino acid myostatin protein (R-Mst), a biologically active recombinant 16-kDa protein containing 113 amino acid residues of the processed human myostatin protein (BioVendor Laboratory Medicine Inc., Palackeho, Czech Republic) [14, 52, 53], or with a recombinant mouse follistatin protein (RD Systems, Minneapolis, MN, USA) at 0.2 μg/ml [11, 14], changing medium twice a week. In other experiments, incubations with the monoclonal (Chemicon International, Temecula, CA, USA) and polyclonal (Millipore Corp, Billerica, MA, USA) antibodies against myostatin (1:20) were substituted for the previous treatments. Alternatively, the adenoviruses expressing the mouse myostatin full-length cDNA under the CMV promoter (AdV-CMV-Mst375) and an shRNA, which targets myostatin RNA and inhibits more than 95% of myostatin gene expression [11, 14, 18] (AdV-Mst shRNA) were transduced into MDSCs at 80% confluence. Then cells were switched to GM-HC medium, as described earlier.
Implantation of MDSCs into skeletal muscle
Male mdx mice (C57BL/6/10ScSn-Dmdmdx), referred to here as "mdx", obtained from Jackson Laboratories (Bar Harbor, ME, USA) were allowed to reach 10 months of age, to allow lipofibrotic degeneration to become more evident, not only in the diaphragm but also in the gastrocnemius. In contrast, in young animals (12 to 16 weeks of age), the first round of muscle necrosis and regeneration had already subsided (stable phase).
Mice were treated according to National Institutes of Health (NIH) regulations with an Institutional Animal Care and Use Committee-approved protocol. In one experiment, the WT and mdx MDSCs (0.5 to 1.0 × 106 cells/50 μl saline) were labeled with the nuclear fluorescent stain, 4',6-diamidino-2-phenylindole (DAPI) [27–30], and implanted aseptically under anesthesia into the surgically exposed tibialis anterior. The muscle had been cryoinjured by pinching it for 10 seconds with a forceps cooled in liquid nitrogen immediately before implantation. Control mice with the same cryoinjury received saline. Mice were killed after 2 weeks, and the tibialis excised and subjected to cryoprotection in 30% sucrose, embedding in OCT, and cryosectioning.
In another experiment, the DAPI-labeled WT and Mst KO MDSCs (0.5 × 106 cells/50 μl GM) were implanted into the central region of the surgically exposed left gastrocnemius of 10-month-old mdx mice, which 4 days earlier had been injured with two injections of notexin in both tips of the muscle (total: 0.2 μg in 10 μl saline). Control muscle-injured mice were injected with saline (n = 5/group). Mice were killed at 3 weeks, the gastrocnemius excised, and a section around the site of notexin injection was used for cryosectioning. The remaining tissue was kept frozen at -80°C.
Immunocytochemistry and dual immunofluorescence
Cells on collagen-coated eight-well removable chambers, fixed in 2% p-formaldehyde, and 10 μm unfixed frozen tissue sections, were reacted [10, 11, 14, 18, 29, 30] with some of the following primary antibodies against (a) human myosin heavy-chain fast, detecting both MHC-IIa and MHC-IIb); monoclonal, 1:200 Vector Laboratories, Burlingame, CA, USA), a marker for skeletal myotubes and myofibers; (b) human ASMA (mouse monoclonal in Sigma kit, 1:2, Sigma Chemical, St. Louis, MO, USA), a marker for both SMCs and myofibroblasts; (c) neurofilament 70 (NF70; mouse monoclonal, 1:10, Millipore); (d) Dystrophin (rabbit polyclonal, 1:200 Abcam, Cambridge, Massachusetts, USA); (e) Sca-1 (mouse monoclonal, 1:100, BD Pharmingen, San Jose, CA, USA) and M.O.M blocking kit (Vector, Burlingame, CA, USA); and (f) Oct 4 (rabbit polyclonal, 1:500, BioVision, Mountain View, CA, USA). When MDSCs in eight-well chambers were not previously tagged with DAPI, all nuclei were stained with coverslips with DAPI antifading emulsion.
Cultures or tissue sections not involving DAPI labeling were subjected to immunohistochemical detection by quenching in 0.3% H2O2, blocking with goat (or corresponding serum), and incubated overnight at 4°C with the primary antibody. This was followed by biotinylated anti-mouse IgG (Vector Laboratories), respectively, for 30 minutes, the ABC complex containing avidin-linked horseradish peroxidase (1:100; Vector Laboratories), 3,3' diaminobenzidine, and counterstaining with hematoxylin, or no counterstaining. For cells labeled with DAPI, fluorescent detection techniques were used. The secondary anti-mouse IgG antibody was biotinylated (goat, 1:200, Vector Laboratories), and this complex was detected with streptavidin-Texas Red. After washing with PBS, the sections were mounted with Prolong antifade (Molecular Probes, Carlsbad, CA, USA). Negative controls in all cases omitted the first antibodies or were replaced by IgG isotype. In the case of Oct 4, streptavidin-FITC was used.
In tissue cryosections for experiments involving DAPI-labeled cells (10 μm), tissue sections were processed in regions where the DAPI+ cells could be detected. Muscle fibers were either stained with hematoxylin/eosin, or by MHC-II antibody, either by Texas red fluorescence as previously described, or with the diaminobenzidine tetrahydrochloride-based detection method (Vectastain-Elite ABC kit; Vector Labs), counterstaining with Harris hematoxylin. Tissue sections that were incubated with mouse IgG instead of the primary antibody served as negative controls. The sections were viewed under an Olympus BH2 fluorescent microscope, and cell cultures, under an inverted microscope. In some cases, the cytochemical staining was quantitated by image analysis by using ImagePro-Plus 5.1 software (Media Cybernetics, Silver Spring, MD, USA) coupled to a Leica digital microscope bright-field light fluorescence microscope/VCC video camera. After images were calibrated for background lighting, integrated optical density (IOD, area × average intensity) was calculated.
Gene transcriptional expression profiles
Pools of total cellular RNA from three T25 flasks for each MDSC cultured in DM-20 were isolated with Trizol-Reagent (Invitrogen, Grand Island, NY, USA) and subjected to DNAse treatment, assessing RNA quality by agarose gel electrophoresis. cDNA gene microarrays (SuperArray BioScience Corp., Frederick, MD, USA) [11, 24, 41] were applied, by using the mouse stem cell (OMM-405), Oligo GEArray microarray. Biotin-labeled cDNA probes were synthesized from total RNA, denatured, and hybridized overnight at 60°C in GEHybridization solution to these membranes. Chemiluminescent analysis was performed per the manufacturer's instructions. Raw data were analyzed by using GEArray Expression Analysis Suite (SuperArray BioScience Corp.). Expression values for each gene based on spot intensity were subjected to background correction and normalization with housekeeping genes, and then fold changes in relative gene expression were calculated. Microarray data were deposited in the Gene Expression Omnibus (GEO) public repository (accession number GSE28986).
The expression of some of the down- or upregulated genes detected earlier was examined on 1 μg RNA isolated from consecutive similar incubations performed in triplicate by reverse transcription (RT) by using a 16-mer oligo(dT) primer, as previously described [11, 27], and the resulting cDNA was amplified using PCR in a total volume of 20 μl. The locations of the primers used for the quantitative estimation of mouse myostatin mRNA were nts 136 to 156 (forward) and 648 to 667 (reverse), numbering from the translation initiation codon, as previously described. For mouse GAPDH primers, sequences were from the mRNA sequence NM_008084.2, with a forward primer spanning nts 778-797 and reverse primer spanning nts 875-852, with a product length of 98 nt.
Additional primers were designed by using the NCBI Primer Blast program applied to mRNA sequences and synthesized by Sigma-Aldrich. Numbering refers to the length in NT from the 5' end of the mRNA: (a) Acta1 (skeletal muscle actin) NM_009606.2 (forward 501 to 520 and reverse 841 to 822; product length, 341); (b) Actc1 (cardiac actin) NM_009608.3 (forward 38 to 58 and reverse 554 to 530, product length 517); (c) MyoD NM_010866.2 (forward 515 to 534 and reverse 1013 to 994, product length 499); and (d) Pax3 NM_008781.4 (forward 1164 to 1183 and reverse 1893 to 1874, product length 730). The number of PCR cycles used for each primer set is stated in parenthesis, as follows: Actc1 (30), Acta1 (30), MyoD1 (33), Pax3 (28), and GAPDH (26). All primers were designed to include an exon-exon junction in the forward primer, except for GAPDH and MyoD1. Negative controls omitted cDNA.
Protein expression by Western blots
Cells were homogenized in boiling lysis buffer (1% SDS, 1 mM sodium orthovanadate, 10 mM Tris pH 7.4 and protease inhibitors, followed by centrifugation at 16,000 g for 5 minutes [10, 11, 14, 18, 29, 30]. Then 40 μg of protein was run on 7.5% or 10% polyacrylamide gels, and submitted to transfer and immunodetection with antibodies against (a) human ASMA (monoclonal, 1:1,000; Calbiochem, Billerica, MA, USA); (b) Oct 4, as for immunohistochemistry; (c) MyoD (rabbit polyclonal 1:200, Santa Cruz Biotechnology, Dallas, TX, USA); (d) MHC (fast), as for immunohistochemistry; (e) TGF-β1 (rabbit polyclonal, 1:1,000; Promega Corporation, Madison, WI, USA); (f) myostatin (rabbit polyclonal 1:1,000; Chemicon International Inc), (g) ActRIIb (monoclonal, 1:1,000, Abcam); and (h) GAPDH (mouse monoclonal, 1:3,000, Chemicon). Membranes were incubated with secondary polyclonal horse anti-mouse or anti-rabbit IgG linked to horseradish peroxidase (1:2,000; BD Transduction Laboratories, Franklin Lakes, NJ, USA, or 1:5,000, Amersham GE, Pittsburgh, PA, USA), and bands were visualized with luminol (SuperSignal West Pico; Chemiluminescent, Pierce, Rockford, IL, USA). For the negative controls, the primary antibody was omitted.
Values are expressed as the mean (SEM). The normality distribution of the data was established by using the Wilk-Shapiro test. Multiple comparisons were analyzed with a single-factor ANOVA, followed by post hoc comparisons with the Newman-Keuls test. Differences among groups were considered statistically significant at P < 0.05.
MDSC cultures from the Mst KO resemble their counterparts from WT mice in morphology, replication, cell markers, and multipotent differentiation
WT MDSCs (pP6 fraction) formed in vitro the most robust skeletal myotubes (see next section) at about passage 13, and WT MDSCs and Mst KO MDSCs were compared from passages 10 through 28. The morphology of the proliferating cultures was similar, but the replication times for the Mst KO MDSCs were slower than those for the WT MDSCs (27.0 versus 19.8 hours, respectively). This morphology and replication pattern continued throughout the 13- through 28-passages period of study.
The stem cell nature of the nuclear Oct 4A expression was confirmed by the detection of the 45-kDa Oct 4A transcriptionally active protein accompanied to a lesser extent by the 33-kDa Oct 4B of cytoplasmic origin (Figure 1B bottom).
Some stem cell-related genes are transcribed similarly in MDSCs, irrespective of myostatin inactivation
Pou domain (Oct4)
Alkaline phosphatase 2
Alkaline phosphatase 5
Telomerase reverse transcriptase
Undifferentiated embryonic cell TP1
Leukemia inhibitory factor
Peroxisome proliferator-activated receptor γ
The genetic inactivation of myostatin is, however, associated with the loss of the ability of MDSCs to form myotubes in vitro, and with the downregulation of key myogenic genes
Some skeletal myogenesis-related genes are downregulated in MDSCs by myostatin genetic inactivation, whereas others remain unchanged
Secreted phosphoprotein 1 (osteopontin)
Myogenic differentiation 1
Myogenic factor 5
Notch gene homolog 2
Bone morphogenetic receptor 2
Bone morphogenetic receptor 1a
Bone morphogenetic receptor 1b
Bone morphogenetic protein 4
Insulin-like growth factor 1
Frizzled homolog 1
Notch gene homolog 1
Notch gene homolog 3
Myotube formation cannot be induced in Mst KO MDSCs by stem cell-reactivating agents, and the WT MDSCs are also refractory to positive or negative modulation of myostatin expression
This suggests that the WT MDSC ability to form myotubes is refractory to the modulation by myostatin, and this was confirmed by transfection with the AdV Mst cDNA construct, or alternatively, with the AdV Mst shRNA, which also expresses β-galactosidase, which did not inhibit or stimulate this process, although myostatin and β-galactosidase were respectively expressed (not shown). The suppression of myotube formation in the Mst KO MDSCs by myostatin genetic inactivation and the lack of response to demethylating agents suggests that this is a complex imprinting process occurring during their embryologic generation, of a different nature than the resistance to paracrine and autocrine myostatin modulators observed in the WT MDSCs.
Mst KO MDSCs stimulate myofiber repair in the injured gastrocnemius of the aged mdx mouse, but the absence of myostatin in these cells does not confer on them a distinctive advantage over the WT MDSCs
To our knowledge, this is the first report testing the myogenic capacity of MDSCs isolated from transgenic mice with inactivation of the myostatin gene, in comparison to the WT MDSC, both in vitro and in the injured muscle of the aged mdx mice in vivo. Our main findings were (a) in contrast to WT MDSCs, Mst KO MDSCs were unable to form myotubes in vitro, although no major differences were found between both MDSC cultures in terms of morphology, replication rates, expression of most members of a subset of key embryonic-like stem cell and other markers, and nonmyogenic multilineage differentiation; (b) however, a fundamental difference is that the expression of key genes in myogenesis seen in WT MDSCs such as actc1, acta1, and myoD, was virtually obliterated in Mst KO; (c) surprisingly, both types of MDSCs were refractory in vitro to the modulation or induction of myotube formation by well-known regulators of this process, or of myofiber number in vivo, such as demethylating agents, myostatin inhibition or overexpression, or follistatin, although myostatin receptors are expressed in MDSC cultures; (d) the myofiber regeneration and anti-lipofibrotic capacities of WT MDSCs were evident even in the environment of a severely injured mdx gastrocnemius at an age at which lipofibrotic degeneration is considerable; (e) in turn, these capacities, blocked in cell culture, were recovered in Mst KO MDSCs when they were implanted in the injured mdx aged-muscle setting, even if not at the level expected from the supposed paracrine effects triggered in the MDSCs by the absence of myostatin.
It should be noted that although notexin-induced injury is not clinically relevant for DMD, it is experimentally convenient by stimulating cell engraftment on implantation and also inducing more lipofibrotic degeneration both in mdx and Mst KO mice [56, 57], thus providing an adequate environment for testing the MDSC-repair effects. The high variability in the repair response that is often associated with notexin injection was not observed in the current work.
The WT MDSC used here as control, fulfill all the criteria that have been extensively defined as potential tools for skeletal muscle, cardiac, and osteogenic repair on implantation into the target organs [29, 34]. In the current work, MDSCs were isolated as the pP6 fraction by using a modification of the extensively validated preplating procedure on collagen-coated flasks and Sca1 selection, and shown to have the expected morphology, rapid replication for at least 50 passages, express MDSC markers such as Sca1, CD44, and CD34, and the stem cell gene Oct 4, and the ability to differentiate in vitro into multiple cell lineages. The latter capability includes a robust formation of multinucleated and branched myotubes that is assumed to translate in vivo into their ability to donate their nuclei to injured skeletal myofibers or most likely to stimulate paracrinely their regeneration through paracrine trophic effects [32–34]. This is evidenced by a much higher number of centrally located nuclei, and even some central location of the DAPI-labeled implanted nuclei. In previous studies, we showed that WT MDSC generate at least smooth muscle and epithelial cells when implanted into urogenital tissues [27, 28], adding to the extensive demonstration of their stem cell nature [7, 12, 26, 58] related to their putative origin as myoendothelial stem cells in the muscle and other tissues .
Another novel finding here is that WT MDSCs have some embryonic-like stem cell features, mainly the expression of nuclear Oct 4 A, myc, LIF, and other embryonic stem cell genes. Oct 4 is a key not only for embryonic stem cell programming, but also for iPS generation, where it can act virtually by itself . Our MDSC cultures contain some tiny rounded cells similar to the very small embryonic-like stem cells (VSELs) described in many adult organs , and other larger ones.
An important finding is the unexpected observation that myotube formation by the WT MDSCs in vitro is refractory to modulation by agents that are well known to affect this process, or skeletal muscle mass in vivo. The fact that myotube formation by WT MDSCs was not influenced by (a) demethylating agents like azacytidine that stimulate 'stemness" in cell lines ; (b) downregulation or overexpression of myostatin, despite the detectable expression of its receptor (ActIIb); (c) counteracting myostatin activity by the respective antibodies or follistatin, that in vivo stimulate myofiber growth [17, 19, 20]; poses questions related to the role of MDSCs during normal myogenesis. A study showing that myostatin stimulated fibroblast proliferation in vitro and induced its differentiation into myofibroblasts, while increasing TGF-β1 expression in C2C12 myoblasts, did not examine MDSC differentiation . The claim of a small inhibitory effect of myostatin on the fusion index in MDSCs  may indicate less fusion efficiency but might not entirely reflect the actual effects on the number and size of myotubes, as determined here. This question requires further clarification in terms of the actual modulation of MDSC differentiation.
It may be speculated that satellite cells rather than MDSCs are the only myogenic progenitors during normal myofiber growth, as opposed to repair of damaged fibers . Therefore the selected in vitro conditions may not mimic the repair process, or alternatively, unknown in vivo paracrine or juxtacrine modulators may modify the response of MDSCs to the better-characterized agents tested in this work. Another possibility is that myostatin and other modulators investigated here would stimulate in vivo satellite cell replication and fusion to the adjacent myofibers to induce hypertrophy, without truly affecting MDSC differentiation or fusion.
We are unaware of any report on the isolation or characterization of MDSCs from the Mst KO. Therefore, it is also both novel and unexpected to find that these cells obtained from the same skeletal muscles as the WT MDSCs, by using identical procedures, and displaying rather similar nonmyogenic pluripotency and stem cell-marker features, are however completely unable to form myotubes in vitro. In fact, our prediction was that the Mst KO MDSCs should be more myogenic than the WT MDSCs because of the absence of the myogenic inhibitor myostatin, The fact that Mst replenishment, either as recombinant protein or as cDNA, does not counteract the unexpected myogenic blockade found in the Mst KO MDSCs, suggests speculatively that these cells have been imprinted in the embryo by the myostatin genetic inactivation through downstream pathways that have become unresponsive to the in vitro myostatin modulation that we explored here. This may involve genes in other myogenic pathways whose expression may be altered, as we observed in Mst KO MDSCs. However, validation of this assumption requires further investigation.
An interesting corollary is the activation of the in vitro-suppressed myogenesis in Mst KO MDSCs, and/or their ability to fuse with preexisting myofibers, after their implantation into the notexin-injured mdx gastrocnemius. At the age selected (10 months), this muscle experiences the considerable damage that occurs in the diaphragm much earlier [3, 4], and this is compounded by injury. It may be speculated that the restoration of myotube (myofiber) formation by Mst KO MDSCs in this setting occurs by paracrine or juxtacrine modulation, possibly of some of the key genes silenced in these cells. Estimation of their products and proof-of-function approaches may elucidate this issue. The fact that although Mst KO MDSCs are able to fuse with or differentiate into new myofibers, they do not increase the muscle-repair process in a clearly more efficient way than do WT MDSCs, may possibly result from the persistent myostatin expression in the fibers that may counteract its absence in Mst KO MDSCs. This suggests the need to block myostatin systemically in the host muscle [63, 64], not just in the implanted MDSCs, and our findings do not contradict the potential use of this approach
One of the genes that may be involved in the silencing of Mst KO MDSC myogenesis in vitro and its reactivation in vivo is the cardiac α-actin (Actc), the major striated actin in fetal skeletal muscle and in adult cardiomyocytes, but strongly downregulated in adult skeletal muscle to 5% of the total striated actin , and whose mRNA is highly expressed in the proliferating (nondifferentiating) WT MDSCs but at very low level in the Mst KO MDSCs. The same applies to the α1-actin (Acta1) mRNA, the adult protein encoding thin filaments . Because actins are so crucial for cell division, motility, cytoskeleton, and contraction, and mutations are associated with severe myopathies, it would not be surprising that their downregulation could cause the lack of myogenic commitment in vitro in Mst KO.
Similarly, the striking transcriptional downregulation of myoD, a critical early gene in skeletal myogenesis , confirmed at the protein level, and of secreted phosphoprotein 1, or osteopontin, a gene mostly involved in ossification, inflammation, and fibrosis, but postulated recently to participate in early myogenesis and skeletal muscle regeneration , may also trigger the absence of myogenic capacity in Mst KO. Interestingly, the fact that Pax 3 mRNA, upstream of MyoD in the myogenic signaling  is expressed in Mst KO MDSCs at higher levels than in WT MDSCs, suggests that the myogenic commitment of Mst KO and mdx MDSC is arrested at some point between these genes. Because a critical regulator of skeletal muscle development, Mef2a (myocyte enhancer factor 2a) , is expressed similarly in both MDSCs (as is Pax 3), albeit at very low levels, the silencing may occur at the level of the satellite cell marker, Pax 7. Therefore, it is not surprising that expression of a member of the cadherin family (cadherin-15) that is involved in later stages, such as myoblast differentiation and fusion , is so downregulated in Mst KO MDSCs.
Our results show that MDSCs obtained from wild-type and Mst KO mice lacking myostatin express Oct 4 and other embryonic-like stem cell genes and appear similar in most features, except for the null or poor expression in Mst KO MDSCs of some critical early genes. These genes encode factors critical for myogenesis and for maintaining the integrity of myotubes and myofibers, thus possibly leading to their inability to form myotubes in vitro. The cross-talk of Mst KO MDSCs with myofibers and other cell types in the host injured mdx muscle may release the pertinent gene silencing and restore the typical myogenic ability of the MDSCs. Although our results do not prove the initial working hypothesis that myostatin inactivation would enhance the myogenic capacity of MDSCs, this possibility still needs further in vivo testing by blocking myostatin, not just in the implanted MDSCs, but also in the host muscle with follistatin, shRNA, antibodies, or other procedures. Finally, systemic muscle-targeted WT MDSC implantation that was previously shown as a promising approach to stimulate repair in the adult dystrophic muscle [5, 12, 45, 46], may even be effective in the setting of an injured aged dystrophic skeletal muscle with severe bouts of necrosis .
adenovirus construct expressing the mouse myostatin full-length cDNA under the CMV promoter
- AdV-Mst shRNA:
shRNA against myostatin RNA
α-smooth muscle actin
Duchenne muscular dystrophy
muscle-derived stem cell
- Mst KO:
myostatin knockout mouse
quantitative image analysis
transforming growth factor β1
very small embryonic-like stem cell
This work was supported by DOD W81XWH-07-1-0181 grant, and partially by an NIH R21DK-070003 grant.
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