Research articles

By Dr. Ari Massoudi
Corresponding Author Dr. Ari Massoudi
Institute of Developmental Biology, - France 06250
Submitting Author Dr. Ari Massoudi

Stem cells, Regeneration, Differentiation, Determination, Pluripotency, Multipotency, Totipotency, Myogenesis, Adipocytes, Duchenne, Dystrophy, Myopathology

Massoudi A. Human Adipose Tissue-Derived Mesenchymal Stem Cells Acquire Muscle Identity Only after Spontaneous Fusion with Myoblasts. WebmedCentral STEM CELL RESEARCH 2011;2(9):WMC002229
doi: 10.9754/journal.wmc.2011.002229
Submitted on: 22 Sep 2011 08:15:00 AM GMT
Published on: 22 Sep 2011 06:49:02 PM GMT


Mesenchymal stem cells have an intrinsic capacity to differentiate into various cell types in culture or after transplantation. Previously, mesenchymal stem cells derived from human adipose tissue (hMADS cells) have been reported to restore dystrophin expression after transplantation in mdx mice. In the currentstudy, we investigated the mechanism of skeletal muscle phenotype acquisition by hMADS cells. We first tested various culture conditions to induce an intrinsic myogenic program in hMADS cells. Neither myotube formation nor Pax7, myf-5, MyoD and myogenin expression could be detected under these conditions. In contrast, when co-cultured with myoblasts, some rare hMADS cells reproducibly contributed to hybrid myotube formation, expressed several muscle differentiation markers and had the ability to restore dystrophin expression in human dystrophic background. Interestingly, this myogenic conversion was not associated with the expression of key transcription factors for muscle termination/differentiation.
Therefore, our results clearly demonstrated that acquisition of myogenic identity by hMADS cells did not occur via a bona fide determination/differentiation process. We propose a model of myogenic conversion of human mesenchymal stem cells derived from adipose tissue in which, only after fusion, transcription factors of myoblasts were able to program hMADS genome toward the skeletal muscle lineage.
Key Words: Human mesenchymal stem cells, myogenesis, Duchenne Muscular Dystrophy.


Mesenchymal stem cells (MSCs), one type of adult stem cell, isolated from the stroma of several organs have been proposed to contribute to the regeneration of different tissues including liver, blood, skeletal tissues, heart, and skeletal muscle (1-5). From a cell therapy perspective adipose tissue presents several advantages as a source of MSCs compared to other organs as it is very large and can be easily harvested
by surgery with little trouble (6-10).
A mesenchymal stem cell population, named hMADS cells (human Multipotent Adipose tissue-Derived Stem cells) has been isolated from human infant adipose tissue. hMADS cells exhibit fundamental stem cell properties at the single cell level such as self-renewal and multipotency (11, 12). Indeed, they can be
propagated in vitro without apparent alteration or transformation while maintaining their differentiation capacities. These cells are able to differentiate under specific inductive media into adipocytes, osteocytes, and chondrocytes. In addition, more detailed investigations showed that adipocyte and osteocyte differentiation of hMADS cells are temporally regulated by the expression of determination/differentiation transcription factors specific of these two lineages (13, 14). Interestingly,
these adipose tissue-derived stem cells seem to display potentialities for skeletal muscle regeneration. Indeed, after injection into skeletal muscle of mdx mice, hMADS cell nuclei and human dystrophin protein were detected in fully mature myofibers (12). Mdx mice are a model of the Duchenne Muscular Dystrophy (DMD), a fatal X-linked myopathy. In this disease, DMD symptoms are caused by mutations of dystrophin gene leading to the absence of dystrophin and dystrophin-associated proteins (15-17).
Although skeletal muscle contribution of MSCs after in vivo transplantation has been well documented, little evidence of functional commitment to the myogenic phenotype has been reported (18, 19). It remains unclear whether MSCs can be either programmed to differentiate into fully functional specialized cells or recruited by surrounding cells via cell fusion, thereby acquiring cellular identity and functions (20-25).
Skeletal muscle development requires well defined steps including cell to cell fusion. Indeed, after activation, determined progenitor cells, termed myoblasts, commit into the myogenic lineage. Differentiating myoblasts - termed myocytes - then fuse and form multinucleated syncitial cells termed myotubes. Ultimately, myotubes undergo terminal differentiation to form myofibers that express a cohort of myofibrillar proteins organized in sarcomeres (26). Determination, self-renewal and differentiation of
myoblasts are strictly controlled by a set of transcription factors, i. e. Myogenic Regulatory Factors (MRFs), such as Pax3, Pax7, MyoD, myf-5, myogenin and MRF4 (27).
In an attempt to delineate further the in vivo data obtained with hMADS cells, we investigated their myogenic potential by in vitro assays. First, their intrinsic myogenic potential was assessed using various culture conditions. Then, we performed lineage tracking to monitor incorporation of hMADS cells into multinucleated skeletal muscle myotubes. Finally, in order to discriminate between a pre-fusion determination or a post-fusion conversion, expression of numerous skeletal muscle genes encoded by
hMADS cells was investigated during the myogenic process. Our results led to propose a model of myogenic conversion of human adult MSCs derived from adipose tissue.

Methods and Materials

Cell culture media were purchased from Cambrex, fetal calf serum (FCS) and horse serum from Dutscher S.A., and chick embryo extract from PAA laboratories. FGF2 and EGF were from Euromedex. Matrigel was a product of BD Biosciences. The mouse monoclonal IgG primary anti-human δ-sarcoglycan (NCL-dSARC) and anti-dystrophin (NCL-DYS3) antibodies were from Novocastra. Mouse monoclonal IgG anti-human nuclei antigen and anti-human nestin from Chemicon, anti-human MyoD (G106-647), anti-Connexin43 (clone 2) and anti-M-Cadherin (Clone 5) were from BD Biosciences. The following monoclonal antibodies, mouse IgG anti-Pax7 (PAX7), mouse IgG anti-Myogenin (F5D), mouse IgG anti-desmin (D3), mouse IgM anti-titin (9D10), rat IgG anti-human α5-integrin (BIIG2) and rat IgG anti-human β1-integrin (AIIB2) were purchased from Developmental Studies Hybridoma Bank (University of Iowa). Goat polyclonal IgG anti-
MyoD (S-17) was purchased from Santa Cruz. Alexa Fluor 594-conjugated secondary antibodies,
streptavidin-coupled Alexa Fluor 596 and Hoechst-34580 were purchased from Molecular Probes. HRP or Biotin-conjugated secondary antibodies were from Jackson ImmunoResearch. HRP-ABC kit and DAB detection kit were from Vector Laboratories. CMV-nlsLacZ and PGK-eGFP-lentivirus vectors were kindly provided by Dr. C. Bagnis (Etablissement Français du Sang Alpes Méditerranée, Marseille, France). MMLV reverse transcriptase, RQ1 RNase-free DNase and GoTaq DNA polymerase were purchased from Promega, and TRI-Reagent from Euromedex. All the other products were purchased from Sigma–Aldrich.
Cell cultures
hMADS cells: The establishment, characterization and cultures of hMADS cells have been previously described by (11, 12). hMADS cells used in this report were provided by Dr. Christian Dani (Nice,
France) and were isolated from the pubic region fat pad of a 5-year old male donor. Cells were inoculated at a density of 4500 cells/cm2 in proliferation medium (PM), Dulbecco’s modified Eagle’s medium (DMEM, low glucose) supplemented with 10% FCS, 2.5 ng/ml hFGF2, penicillin/streptomycin. PM was changed every 2-3 days. hMADS cells were sub-cultured when they reach 9000 cells/cm2 (~80% confluence). One passage before co-culture experiments, FGF2 was omitted in PM.
C2C12 cells: C2C12 cells (ATCC) were inoculated at 2500 cells/cm2 and maintained as undifferentiated myoblasts in PM [DMEM (high glucose) supplemented with 20% FCS, penicillin/streptomycin]. They were sub-cultured when they reached 25 000 cells/cm2 (~80% confluence). human DMD and wt-myoblasts: Human DMD immortalized but not transformed myoblastic cell line (named here DMD myoblasts) was kindly provided by Pr. D. Trono (Geneva, Swizerland). These cells were cultured as previously described (28). The human wild-type myoblasts derived from a healthy patient were kindly provided by Dr. G. Butler-Browne (Institut de Myologie, Paris, France) and cultured , in the same PM than DMD myoblasts, i. e. Ham’s F10 supplemented with 20% and penicillin/streptomycin. Mouse primary satellite cells: Proliferating primary satellite cells were obtained from mdx or wild-type mouse isolated myofibers of tibialis anterior muscle as described previously by (29). Briefly, muscles were incubated for 60 min at 37°C in 0.2% (w/v) type-1 collagenase in DMEM (high glucose) supplemented with penicillin/streptomycin. Myofibers were then isolated by trituration. Isolated myofibers were placed individually in wells of 24 well-plates coated with 1 mg/ml Matrigel. PM [DMEM with 10% (v/v) horse serum and 0.5% (v/v) chick embryo extract and penicillin/streptomycin] was added. Under these conditions, myofibers adhere to the substrate and satellite cell-derived myoblasts migrate away from the fiber and proliferate. All listed-cells were cultured at 37°C in a humidified gassed incubator in 5% CO2 atmosphere.
hMADS/C2C12 cells co-cultures: proliferating hMADS cells and C2C12 cells were mixed in a ratio of
3:1, and inoculate at 20 000 cells/cm2 in hMADS PM. In a couple of day after plating, cells reached
100% confluence and PM was shifted to differentiation medium (DM) [DMEM (high glucose)
supplemented with 10 μg/ml insulin, 5 μg/ml transferrin and 1μM dexamethasone and
penicillin/streptomycin]. DM was changed every 2 days. From initial plating until day 4 in DM, a ratio of 0.4-0.5 ml media per cm2 culture area was used. Highly mature myotubes (i.e. > 6 days in DM) were obtained as previously described (30) with some modifications. Cells were plated onto collagen/Matrigel-coated [0.1 mg/ml rat tail collagen, 0.3 mg/ml Matrigel] coverslips. Since day 4 to day 10-14 in DM a ratio of 0.3 ml media per cm2 culture area was used. hMADS/DMD cells co-cultures: the same methods as above were used with some modifications. hMADS and DMD cells were mixed at a ratio of 1:1. DM was DMEM (high glucose) supplemented with 10 ng/ml EGF, 0.05% (w/v) BSA, 1 mM creatine, 110 μg/ml pyruvate, and 50 μg/ml uridine and penicillin/streptomycin.
hMADS/human wt-myoblasts co-cultures: hMADS cells and wt-myoblasts were mixed at a ratio of 1:1. DM was DMEM (high glucose) supplemented with 10 μg/ml insulin, 5 μg/ml transferrin and
penicillin/streptomycin. hMADS/mouse primary satellite cells co-cultures : when a sufficient number of satellite cells has migrate from myofibers (~40% confluence), ~8 500 hMADS cells were added per well onto satellite cells grown on matrigel-coated 24-well plates. Twenty four hours after hMADS cell addition, PM was shifted to DM (DMEM supplemented with 10 % HS, 2% FCS, 0.5 % chick embryo extract and penicillin/streptomycin).

Cells plated on plastic of 48 or 24-well plates, or on coverslips, or collagen/Matrigel-coated coverslips were fixed and assessed for immunostaining. Briefly, cells were rinsed twice with cold phosphate buffer saline (PBS), and then subjected to fixation in 4% paraformaldehyde in PBS for 15 min at room temperature (RT). Free aldehydes were quenched with 100 mM Tris-pH 8.5, 150 mM NaCl for 5mn. Fixed cells were permeabilized with 0.005% (w/v) digitonin in PBS for 15 min.

Permeabilization step was omitted when antigens were at the cell surface (i.e. β1-integrin, α5-integrin, M-Cadherin). Unspecific reactions were blocked with 1% BSA plus or not 1% normal goat serum (depending of secondary antibody origin) in PBS for 30 min. Cells were then incubated with primary antibody 1h at RT. After 3 washes 1% BSA in PBS, Alexa Fluor 594, HRP-conjugated or biotin-conjugated secondary antibodies were incubated 45 mn at RT (in the dark for the former). Antibodies were diluted at 1-10 μg/ml final in 0.1 % carrageenan-λ in PBS (antibody buffer). Antibody buffer for the Alexa Fluor 594-conjugated antibody were supplemented with 0.5 μg/ml Hoechst-34580 for nuclei counterstaining. Biotin-conjugated secondary antibodies were detected with streptavidin-coupled Alexa Fluor 596 or HRP-ABC signal amplification Kit followed by DAB detection. Fluorescent-immunostained coverslips were mounted on slides using Mowiol 4-88 solution. Micrographs of DAB-stained cells were directly taken with a numerical camera (DSC-D75, Sony) under a light transmission microscope (Axiovert25, Zeiss).

Confocal image capture
Fluorescent immunostained cells were viewed on a confocal microscope (LSM510 META, Zeiss) using
Plan-Neofluar x40/1.3 and Plan-Achromat x63/1.4 oil-immersion objectives. Images were optimized globally for contrast and brightness using ImageJ software.
Semi-quantitative RT-PCR
Total RNA was extracted using TRI-Reagent according to manufacturer’s instructions and quality was
assessed on 1% agarose gel to eliminate any degraded RNA. Reverse transcription was carried out with MMLV reverse transcriptase and random primers on 1 μg RNA treated with RQ1 RNase-free DNase. PCR was performed with GoTaq DNA polymerase in a total volume of 25 µl. Cycle numbers (n) were optimized for each primer pair to be in the linear range amplification. For normalization all cDNA amounts under comparison were carefully adjusted between 1/25 to 1/50 of reverse transcription solution to give the same quantity of amplified cDNA with mouse hypoxanthine guanine phosphoribosyl transferase (HPRT) or human β-actin primers. General PCR conditions were the following: 3 min at 95°C followed by n cycles of 30 s at 95°C, 30 s 55°C, and 50 s at 72°C with a final elongation step of 5 min at 72°C. 20 μl of PCR products were analyzed on 1-1.5 % ethidium bromide stained agarose gels. Mouse primers did not anneal with human cDNAs and human primers did not anneal with mouse cDNAs. Specific PCR conditions for each primer pair were as follows. Mouse HPRT: 249 bp, 35 cycles, forward 5’- GCTGGTGAAAAGGACCTCT-3’, reverse 5’- CACAGGACTAGAACACCTGC-3'. Human α-actin: 746 , bp, 23 cycles, forward 5’- AGCCATGTACGTTGCTA-3’, reverse 5’- AGTCCGCCTAGAAGCA-3’. Human desmin: 519 bp, 35 cycles, forward 5’-CCTACTCTGCCCTCAACTTC-3’, reverse 5’- AGTATCCCAACACCCTGCTC-3’. Human enolase-3: 502 bp, 40 cycles, forward 5’- TGACTTCAAGTCGCCTGATGATCCC-3’, reverse 5’- TGCGTCCAGCAAAGATTGCCTTGTC-3’. Human muscle creatine kinase: 720 bp, 40 cycles, forward 5’-GGCACAATGACAACAAGAGC-3’, reverse 5’-GAAAAGAAGAGGACCCTGCC-3’. Human dystrophin: 778 bp, 33 cycles, forward 5’- TTCTCAGCTTATAGGACTGCC-3’, reverse 5’-GGAGTGCAATATTCCACCAT-3’. Human sarcospan: 466 bp, 33 cycles, forward 5’-GGGCTGGGATCATTGTCTGCT-3’, reverse 5’-
GGAATTCTTAGATCTTTTGCTGGGG-3’. Human Pax7: 465 bp, 35 cycles, forward 5’-
417 bp, 35 cycles, forward 5’- TGAGAGAGCAGGTGGAGAACTAC-3’, reverse 5’-GCCTTCTTCGTCCTGTGTATTAG-3’. Human MyoD1: 503 bp, 35 cycles, forward 5’-AAGCGCCATCTCTTGAGGTA-3’, reverse 5’-GCGCCTTTATTTTGATCACC-3’. Human myogenin: 364
bp, 35 cycles, forward 5’-AGCGCCCCCTCGTGTATG-3’, reverse 5’- TGTCCCCGGCAACTTCAGC-3’.
Lentiviral transduction protocol
Transduction of hMADS cells was accomplished as follows. Lentiviral vectors (CMV-nlsLacZ and PGKeGFP vectors) were diluted into 5 ml of PM (for infection in a 100-mm dish). Polybrene was added at a concentration of 10 mg/ml. The viral vector/polybrene mix was left at RT for 15 mn and then added to the cells. After 15 h, cells were washed once in PBS, and fresh medium was added. Three days after infection, transgene expression was assessed. Transgenic nuclear β-galactosidase activity was detected by X-Gal staining. Trangenic GFP expression was assessed by confocal microscopy and FACS analysis. More than 95% of cells were positive for the expression of nuclear β-galactosidase or GFP at a MOI of 10 viruses by cell.


Intrinsic myogenic potential of hMADS cells
In order to evaluate the intrinsic ability of hMADS cells to differentiate into the skeletal muscle lineage, various culture conditions were tested. In vitro, no myotube formation occurred when hMADS cells were cultured in various media supplemented with hormones and cytokines related to myogenesis, as well as with chemical activators or inhibitors of signaling pathways. Moreover, we did not observe any myotube formation by treating hMADS cells with conditioned media from proliferating or differentiating myoblasts of human or mouse origin, or those from regenerating mouse mdx or wild-type skeletal muscles. We checked the expression of Pax7, MyoD and myogenin proteins after treating hMADS cells with different putative myogenic media. Immunostaining data indicated no detectable expression under these various conditions even in the presence or in absence of serum (sup S1). The effect of extracellular matrix components was also assessed. No myotubes were observed when hMADS cells were cultured on dishes coated with collagen, Matrigel, laminin or gelatin.
Thus, hMADS cells are unable to form myotubes and to express MRFs when maintained in various
myogenic media. Therefore, we set up a co-culture assay with myoblasts in order to investigate in details mechanisms by which hMADS cells acquire a myogenic potential.
hMADS cells co-cultured with myoblasts contribute to myotube formation
hMADS cells cells were maintained in co-culture with primary satellite cells isolated from myofibers of mdx mice and the formation of hybrid myotubes was evaluated several days after maintaining cells in differentiation medium. The presence of human nuclei within myotubes was detected by immunostaining using an anti-human nuclei antibody (Figure 1a, a’). Co-cultures with primary satellite cells obtained from myofibers of wild-type mice yielded similar results (not shown).
For a better estimate of hMADS cell myotube contribution, co-cultures were performed with proliferating nlsLacZ-hMADS cells and C2C12 myoblasts and percentages of hybrids were evaluated after myotube formation (Figure 1b). Although, hybrid myotube formation proved to be reproducible, the yield remained rather very low. Hybrid myotubes represented approximately 1-3 % of total myotubes, in which hMADS nuclei represented 1-2 nuclei of total myonuclei. These results demonstrate that less than 1% of hMADS cells plated at t0 of co-culture experiments have fused with myoblasts.
Then we investigated whether hMADS cells could also participate into the formation of human myotubes. For this purpose myoblasts derived from a Duchenne Muscular Dystrophy (DMD) patient (28), was cocultured with nlsLacZ-hMADS cells (Figure 1c). Similarly to co-cultures performed with murine myoblasts, hMADS cells contributed to myotube formation as shown by the nuclear β-galactosidase activity. Hybrid myotubes were similarly obtained using human wild-type myoblasts derived from a healthy patient (not shown). The yield of human-human hybrid myotube formations was in the same range than hMADS/mouse myonlasts hybrids. No homogenous hMADS cell-derived myotubes could be observed in any experiments described above.
Interestingly, positive nuclei in all hMADS cells/myoblasts hybrid myotubes showed an anti-human
nuclei staining or β-galactosidase activity with a gradient intensity starting in a large nucleus (Figure 1, blue arrows) surrounded by small and weakly stained nuclei (Figure 1, black arrows). The large nuclei were identified like coming from hMADS cells whereas the small ones coming from myoblasts (Sup S2). Therefore, this observation indicated that nuclear proteins expressed by hMADS cell nuclei could be transferred in mouse nuclei in hybrid myotubes.
Expression of muscle genes encoded by hMADS cells in co-cultures
Expression of several muscle differentiation markers i. e. desmin, enolase-3, muscle creatine-kinase, sarcospan and dystrophin was investigated in co-cultures of hMADS/C2C12 cells by using semiquantitative RT-PCR and primer combinations that were specific for human transcripts (Figure 2A). This set of genes was induced in a sequence that accurately reflects their regular temporal order during myogenic differentiation. The myogenic conversion of hMADS cells was then evaluated at the protein level for -δsarcoglycan and dystrophin known to be expressed only in myotubes.
Forteen days after maintenance in differentiation medium (DM), hMADS/C2C12 co-cultures expressed human δ-sarcoglycan and confocal microscopy revealed both a striated sarcoplasmic reticulum (Figure 3, blue arrow) and a sarcolemmal staining (Figure 3, yellow arrow) pattern in accordance with the subcellular localisation observed in skeletal muscle (31, 32). Next, the ability of hMADS cells to complement dystrophin in a human dystrophic background was assessed. For this purpose, co-cultures were performed with hMADS cells and human DMD myoblasts. After 10 days under DM, cultures were stained with anti-dystrophin (Figure 4 a,b) or anti-δ-sarcoglycan
antibodies (Figure 4c,d). In both cases, positive staining was observed. Dystrophin-positive myotubes numbers were in the same range than hMADS myotube contributions. The number of δ-sarcoglycanpositive myotubes was higher than that of dystrophin-positive myotubes (see Discussion). As expected, control experiments with myotubes formed from DMD myoblasts presented no dystrophin (Fig. 4b) and lowδ-sarcoglycan (Fig. 4d) stainings. Human myoblasts were used as a positive control for antibody reactions (not shown). Altogether, these data indicate that fusion with human or murine myoblasts led hMADS cells to express a functional myogenic program.

Expression of MRFs in hMADS/C2C12 cell co-cultures
Some hMADS cells could have acquired myoblasts identity by the complex inductive co-culture microenvironment, and then committed to differentiate and fuse with myoblasts. To test this hypothesis, GFP hMADS/C2C12 cells co-cultures were carried out and expression of the MRFs Pax7, MyoD and myogenin was assessed. Immunostaining using the highly sensitive labeled-streptavidin /biotin-coupled secondary antibody detection system did not allow the detection of MRFs in mono-nucleated GFPhMADS cells at different time-points, i.e growing phase, cell confluency and day 1, 2 and 3 in differentiating medium; Fig. 5 is a representative illustration of these data. Similar results were obtained in co-cultures of hMADS cells and DMD myoblasts, excluding species-specific barrier in preventing MRF induction. This lack of expression (including myf-5) was confirmed at mRNA level using human specific RT-PCR from hMADS/C2C12 co-cultures (Figure 3B). Control experiments of hMADS cells cultured alone under the same conditions showed no expression of these genes neither at the mRNA level (not shown) nor at the protein level (sup S1).
After two days in DM, myotubes nuclei expressed myogenin, as expected. Surprisingly GFPhMADS/
C2C12 hybrid myotubes (Fig. 6a) showed also a weak nuclear staining for myogenin protein in
hMADS-derived nuclei (Fig. 6b). The anti-myogenin antibody used in this assay recognized mammal myogenin as no human specific myogenin antibodies are available in our knowledge. Expression of myogenin in hMADS nuclei could not be due to de novo synthesis in the myotube context as RT-PCR failed to detect myogenin mRNA from human origin (Figure 3B). This data strongly suggested that transcription factors of the host cells (myoblasts) could target to nuclei of donor cells (hMADS cells) and induced the myogenic program.
Desmin and nestin expression in hMADS/C2C12 co-cultures
Although hMADS cells did not express MRFs, the complex muscle micro-environment could have
induced the expression of some muscle markers in hMADS cells before fusion (33, 34). To test the
hypothesis of a “partial commitment”, we assessed the expression of the muscular intermediate filament (IF) desmin and the neuro-muscular IF nestin. Before myotube formations, all C2C12 cells were desmin-positive in contrast to GFP-hMADS cells (sup S3). As expected all myotubes were desmin-positive (not shown). As RT-PCR experiments revealed that human desmin mRNA was indeed expressed (Figure 3a), these data suggested that human desmin may contribute to this staining in hybrid myotubes. Nestin is known to be expressed in motor endplates and myo-tendinous junctions of skeletal muscle, and to play a redundant function with the muscle specific IF desmin (35-38). Using a human specific antinestin antibody, we first checked by immunostaining that nestin was expressed in both human wild-type myoblasts (not shown) and DMD myoblasts (sup S4). Then, co-cultures of GFP-hMADS cells and C2C12 myoblasts were analyzed for human nestin expression at different time-points (Figure 7A a-e). Mono-nucleated GFP-hMADS cells were always negative before fusion. However, human nestin was clearly detected in GFP-hMADS/C2C12 hybrid myotubes as soon as they formed (2 days in DM). All of the GFP-positive myotubes were human nestin positive and contained human nuclei. Reciprocally, hybrid myotubes containing hMADS-derived nuclei detected by Hoechst staining proceeded to be always positive for GFP and human nestin. In addition, the staining pattern of human nestin in hybrid myotubes (Figure 7B), i.e. with a higher intensity at the edges, was in accordance with its reported location in skeletal muscle myotubes (35).


This study, using hMADS cells, evaluates the mechanism of human adult MSCs myogenic contribution. First, numerous culture conditions failed to induce myotube formation of hMADS cells even on longterm treatment. This was confirmed by gene expression analysis where no evidence of MRFs expression was found. Moreover, hMADS cells-derived clones, and provided from several donors, failed also to express MRFs (not shown). Second, when co-cultured with human or mouse myoblasts, incorporation of some rare hMADS cells into multinucleated skeletal myotubes was observed by lineage tracking and specific gene expression. Myogenic potential of adipose tissue-derived stroma-vascular cells from mouse origin has been recently investigated (39). These authors observed a restricted subpopulation of cells possessing a complete myogenic determination/ differentiation ability. Interestingly, the group of Braun T. (40, 41) has recently shown that Bone-Marrow derived-Mesenchymal Stem Cells from mouse origin (mBM-MSCs) can
express MRFs and skeletal muscle markers when co-cultured with feeder-cell layers producing Wnt
proteins or, by treating mBM-MSCs with chromatin remodelling agents i. e. 5-azacytidine (AZA, a DNA demethylating agent), and Trichostan (an inhibitor of histone desacetylases). Nevertheless, myotube formation was not observed. Under our culture conditions, we tested activation of Wnt-pathway by using lithium chloride and the chemical Bio, two well-known inhibitors of glycogen synthase kinase-3 described as a negative regulator of this pathway (42-44). In both cases, hMADS cells neither expressed MRFs nor did form myotubes (sup S1). Similarly, after treatment with sodium butanoate, valproic acid (two inhibitors of histone desacetylases) or AZA, hMADS cells did not express MRFs, neither form myotubes. In addition, no differentiation into skeletal muscle was observed when human BM-MSCs was treated with AZA (45, 46). Altogether, differences observed with “human” MSCs could be explained by a higher plasticity of MSCs from “murine” origin. At the cellular level, this decrease of MSCs plasticity during evolution might participate in the weaker organ regenerative capacities of phylogenetically newer mammals such as human (47). Recently, comparative experiments performed between human “fetal” versus “adult” MSCs have been reported with respect to myotube formation (45). In contrast to fetal MSCs but consistent with our data, these authors failed to differentiate adult MSCs into myotubes. In addition, MSCs obtained from human embryonic stem cells (48, 49), after treatment with conditioned medium from differentiating C2C12 cells can express MRFs (49). Thus, along the transition of MSCs from an “embryo-fetal” to an “infant-adult” stage, there might be a lineage restriction that could explain differences in myogenic potential of MSCs (50). Therefore, bona fide myogenic determination/differentiation capacity of “adult” MSCs from human remains to be clearly demonstrated. Consequently, the next issue was to gain molecular insights into the in vivo contribution of hMADS cells and human dystrophin expression in mdx mice that we had reported after transplantation in the tibialis anterior muscle (12). Most likely, exogenous hMADS cells transplanted into regenerating mdx mouse muscle interacted preferentially with activated satellite cells (51, 52) whereas interactions with other cell types should have remained of low magnitude, suggesting that hMADS/myoblasts in vitro co-cultures were relevant in term of cell fate recapitulation. Our results clearly demonstrate the fusion of some rare hMADS cells with myoblasts independently of species origin. These hMADS cells participate in myotube formation which takes place only in the presence of murine or human fusing-myoblasts. In addition, the existence of hybrid pre-myotubes (3-5 nuclei per syncititum) suggests that hMADS cells have fused at the beginning of the differentiation process. Nevertheless, the percentage of hMADS-derived hybrid myotubes remains very low. Two hypotheses could explain these results. Either, a minor sub-population of hMADS cells has the ability to fuse. Or, all hMADS cells share a weak fusogenic ability under our culture conditions. Some candidate genes reported to be involved in myoblast fusion which are expressed in many cell types, such as some integrins (53-56) and connexin-43 (57-60), are also expressed in hMADS cells (sup S5A, B and C). In contrast, expression of muscle specific cadherin (M-Cadherin) known to have a role in myoblast fusion (61, 62) was observed in C2C12 cells but not in hMADS cells (sup S5D). Further investigations should be carried out for better defining fusion capacity of hMADS cells.
Various cells have been reported to be unable to fuse with myoblasts, whereas other cells are able to form hybrids (34, 63-68). Among all these latter cells a low yield of hybrid myotube formation could be noted, suggesting that myoblasts preferentially fused together rather with other cells. However, fusion ability per se may not be sufficient to lead to muscle gene expression (69-72). In contrast to cells that fuse with myoblasts but do not express muscle genes or any genes, some cells after fusion disturb myogenic program (64, 73). Expression of skeletal muscle genes encoded by adult stem cells genome in hybrid myotubes is a key issue with respect to transplantation in humans. hMADS cells exhibit a broad myogenic conversion after fusion as shown by expression of mature δ-sarcoglycan and nestin proteins of human origin. Other muscle markers encoded by hMADS cell genome were also induced. In addition, any hMADS cells that fused with myoblasts followed a muscle fate, as revealed by the co-expression of GFP and human nestin protein in GFP-hMADS/C2C12 hybrid myotubes. No GFP-negative and human nestin-negative hybrid myotubes were detected containing hMADS-derived nuclei. Altogether, these data indicate that fusion does not impair expression of muscle genes in host cells and induces myogenic program of hMADS cells.
Dystrophin expression is a key issue when considering DMD disease cytotherapy. Our results show
that dystrophin was indeed expressed, at odds with a previous report showing the absence of dystrophin expression in hybrid myotubes derived from human BM-MSC/DMD myoblasts (74). Indeed, dystrophin is a very late marker, and we were able to detect a clear linear staining only in some rare highly differentiated myotubes (since day 10 in DM). Under five days in DM, no positive myotubes could be found in accordance with previous reports (30, 75). The discrepancy concerning the number of dystrophin and δ-sarcoglycan-positive myotubes found in hMADS/DMD co-cultures could be due to the late expression of dystrophin, compared to that of δ-sarcoglycan that is easily detectable in myotubes as early as day 4 under DM, whereas, dystrophin protein begins to be synthesized at day 5-6 (30). After this stage, it cannot be ruled out that dystrophin expression remains restricted to mature producing myo-nuclear domains (65). In contrast, δ-sarcoglycan synthesized earlier in an immature syncitium may have been spread within all the myotubes due to vesicular trafficking leading to an apparent larger staining area (76, 77). The restoration of other dystrophin associated-proteins in hMADS/DMD hybrid myotubes should be investigated.
Finally, we have demonstrated that the contribution of hMADS cells to myotube formation is neither due to a determination/differentiation process, i.e. the de novo expression of muscle “key” transcription factors, nor to a “partial” commitment process, i.e. the expression of muscle markers independently of MRFs expression. Indeed, neither Pax7, MyoD, myf-5, or myogenin are expressed by hMADS cells in co-cultures. Rather, by using the marker nestin expressed at all stages of myogenesis, we found that hMADS myogenic conversion occurred only after fusion.
Forced acquisition of muscle identity was first reported by Blau et al. in amniocytes/myoblasts PEGinduced heterocaryons (78). In our model, heterocaryonic hybrids formed spontaneously without PEG.

Moreover, in hybrid myotubes, myogenin protein encoded by C2C12 nuclei was detected in hMADS
nuclei. This result strongly supports a direct transactivation of hMADS-encoded skeletal muscle genes by MRFs in myotubes, i.e. myogenin, MEF2D, Myf-5, MRF4. Myogenin, is known to induce MRF4 gene (79), and it could also transactivate this gene in hMADS nuclei; this hMADS-encoded MRF4 could then participate in later stage of muscle conversion. A model of hMADS cell myogenic conversion is proposed in Figure 8. Concomitantly, non-muscle loci could be repressed in hMADS nuclei after fusion with myoblasts, as it has been recently reported in human lymphocyte-B or human keratynocyte/C2C12 PEG-induced hybrids (80, 81). This possibility is currently difficult to confirm in our model because specific MSC markers have not yet been identified (82).
Interestingly, human nuclei antigen and β-galactosidase activity encoded by hMADS genes were also detected in mouse-derived nuclei in hybrids. This result suggests that nuclear proteins encoded by hMADS genome and synthesized in the myotube can be targeted into myoblast-derived nuclei. Such hMADS-encoded nuclear proteins could alter in an unknown manner the transcriptome of myoblast derived nuclei within hybrids. Most likely, these putative transcriptional modifications should not disturb myotube/myofiber identity as shown by the numerous skeletal muscle genes expressed in hMADS-derived hybrids [such as titin (sup S2Ab), skeletal muscle myosin, desmin, α-sarcomeric actin, M-Cadherin and α-sarcoglycan proteins (not shown)].
In conclusion, at odds with Rodriguez article (12), our results indicate that adipose tissue-derived
mesenchymal stem cells are not an attractive tool for myo-cytotherapy due to their low fusogenic
capacity. Muscle cell therapy based on hMADS cells transplantation will not be possible if the yield of fusion with myoblasts can not be enhanced.


We are grateful to Pr. Didier Trono and Drs. Claude Bagnis, Gillian Butler-Browne for providing
respectively human DMD myoblast cell line, lentivectors and human primary myoblasts. Special thanks are due to Drs. Christian Elabd, Laure-Emmanuelle Zaragosi, Florence Massiera, Nathalie Billon, Pascal Peraldi for critical review of the manuscript. This work was supported by “Association Française contre les Myopathies (AFM)”, and Ali Massoudi was a scholarship recipient of an AFM fellowship.


MSCs- Mesenchymal Stem Cells
hMADS- human Multipotent Adipose-tissue Derived Stem
DMD- Duchenne Muscular Dystrophy
MRFs- Myogenic Regulatory Factors
PM- Proliferation Medium
DM- Differentiation medium
GFP- Green Fluorescent Protein
IF- Intermediate Filament
PEG- PolyEthylene Glycol.


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This work was supported by “Association Française contre les Myopathies (AFM)”, and Ari Massoudi was a recipient of an AFM fellowship.

Competing Interests



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