TY - JOUR
AU - Wang, Yaming
AB - Abstract Certain lower organisms achieve organ regeneration by reverting differentiated cells into tissue-specific progenitors that re-enter embryonic programs. During muscle regeneration in the urodele amphibian, postmitotic multinucleated skeletal myofibers transform into mononucleated proliferating cells upon injury, and a transcription factor-msx1 plays a role in their reprograming. Whether this powerful regeneration strategy can be leveraged in mammals remains unknown, as it has not been demonstrated that the dedifferentiated progenitor cells arising from muscle cells overexpressing Msx1 are lineage-specific and possess the same potent regenerative capability as their amphibian counterparts. Here, we show that ectopic expression of Msx1 reprograms postmitotic, multinucleated, primary mouse myotubes to become proliferating mononuclear cells. These dedifferentiated cells reactivate genes expressed by embryonic muscle progenitor cells and generate only muscle tissue in vivo both in an ectopic location and inside existing muscle. More importantly, distinct from adult muscle satellite cells, these cells appear both to fuse with existing fibers and to regenerate myofibers in a robust and time-dependent manner. Upon transplantation into a degenerating muscle, these dedifferentiated cells generated a large number of myofibers that increased over time and replenished almost half of the cross-sectional area of the muscle in only 12 weeks. Our study demonstrates that mammals can harness a muscle regeneration strategy used by lower organisms when the same molecular pathway is activated. Stem Cells 2014;32:2492–2501 Cell cycle, Adult stem cells, Cellular therapy, In vivo optical imaging, Muscle stem cells, Reprogramming, Skeletal muscle, Transcription factors Introduction Unlike the strategies that have led to the current nuclear reprogramming protocols to create pluripotent cells from differentiated cells or converting lineage committed cells to mature cells of other lineages [1-3], urodele amphibians and zebrafish regenerate lost organs using a different nuclear reprogramming strategy. In response to injury, their differentiated cells re-enter the cell cycle and instead of acquiring pluripotency, the dedifferentiated cells retain their original tissue identities and reform these specific lost tissues during regeneration [1, 4, 5]. Whether it is possible to leverage this primitive regenerative strategy to induce new tissue and organ formation in mammals has been a longstanding question; however, there has been much debate about this possibility. In the past few years, studies using different methods demonstrated that postmitotic mammalian multinucleated myotubes could be induced to dedifferentiate into mononuclear proliferating cells. Ectopic expression of Msx1, the transcription factor that is upregulated and drives muscle cell dedifferentiation in urodele amphibians [6], has been shown to dedifferentiate multinucleated C2C12 myotubes into proliferating mononuclear cells [7, 8]. Interestingly, these dedifferentiated mononuclear cells display properties that were more primitive than C2C12 cells. However, due to the tumorigenic nature of C2C12 cells, whether these dedifferentiated mouse muscle cells possessed the regeneration capability of their amphibian counterparts was not explored. In an attempt to induce dedifferentiation without overexpression of Msx1, Pajcini et al. showed that concomitant transient inactivation of Arf and Rb led mammalian myotubes (myocytes) to cellularize and re-enter the cell cycle. The mononuclear clones derived from these myotubes were capable of fusing with existing muscle [9]. It has also been shown that treatment of differentiated muscle cells with small molecules such as the cyclohexylaminopurine reversine, induces a proliferative response, mainly though downregulation of cyclin-dependent kinase inhibitors or tyrosine phosphatases [10, 11]. These cells have been shown to be multipotent, and are able to fuse into existing muscle after cardiotoxin injury. More recently, it was shown that downregulation of myogenin, one of the myogenic regulatory factors, can reverse the differentiation state of terminally differentiated mouse myotubes and initiate their cleavage into mononucleated cells [12]. However, whether or not these dedifferentiated mammalian muscle cells possessed long-term regeneration capability that is similar to their amphibian counterparts was not explored. We therefore set out to examine whether ectopic overexpression of Msx1 could drive primary multinucleated murine myotubes to re-enter the cell cycle and furthermore to determine whether and how these dedifferentiated progenitor cells regenerate skeletal muscles after transplanting them into different in vivo microenvironments. Materials and Methods Cell Culture and Gene Transduction Primary myoblasts were isolated from hind limb muscles of 4-week-old C57BL/10 male mice as described previously [13, 14]. Cells were expanded in Ham’s F-10 medium supplemented with 20% fetal calf serum and 5 ng/ml basic fibroblast growth factor (growth media) on collagen-coated plates. Before transduction, the myogenic identity of cells was verified with anti-desmin antibody through immunocytochemistry. Retroviral vectors LINX-Msx1-fwd and LINX-Msx1-rev (kind gifts of Dr. Shannon Odelberg) were packaged as described elsewhere and the sequence was driven by a Tet-off inducible system [8]. Primary myoblasts at passage 5 were transduced with either LINX-fwd or -rev virions. The transduced cells were selected using G418 and expanded clonally in the presence of doxycycline 3 µg/ml. A proportion of cells from all three groups were transduced with either eGFP or nl-GFP lenti-viral virions as described elsewhere [15]. The nl-GFP is specifically targeted to the nuclei and was used to visualize the myonuclei in real-time microscopic imaging in this study (Supporting Information Movies). eGFP transduced cells were used for the ectopic and intramuscular cell transplantation experiments in severe combined immunodeficiency (SCID) mice. Induction of Myotube Dedifferentiation To induce myogenic differentiation of primary myoblasts, cells were cultured in differentiation medium (Dulbecco’s modified Eagle’s medium with 2% horse serum) with 3 µg/ml doxycycline on collagen-coated plates. On the fourth day differentiation in the presence of doxycycline, myotubes were removed from the plate with trypsin, filtered through a 100 µm filter and then through a 40 µm secondary filter. The remaining myotubes on the sieve were collected and then diluted with differentiation medium. Single myotubes were picked up manually by a pipette and placed in 48-well plates with 1 myotube/well. On the next day, all wells were examined carefully by visual inspection for the presence of any residual mononuclear cells (Supporting Information Fig. S2). Contaminated mononuclear cells were destroyed using a 25Ga needle, or the wells were excluded from further observation. The induction of dedifferentiation was then initiated by withdrawing doxycycline and switching to growth medium with addition of 1× Insulin-Transferrin-Selenium (Invitrogen, Inc., Carlsbad, CA, http://www.lifetechnologies.com/us/en/home/brands/invitrogen.html). The area containing each myotube was circled on the bottom of the plate (Supporting Information Fig. S2). In a separate set of experiments, the myotubes were cultured in methylcellulose semisolid medium. MethoCult (StemCell Technologies, British Columbia, Canada, http://www.stemcell.com/?_kk=stemcell%20technologies&_kt=757f97cd-b99e-4620-a657-06741856726e&gclid=CJqk2IC3n74CFU5afgodAn0AGQ) mixed with liquid growth medium in 40:60 ratio with the final concentrations of Fetal Calf Serum (FCS) and growth factors adjusted to be the same as liquid growth medium. Once the dedifferentiation occurred, the mononuclear proliferating cells in the well were picked out after a typical clone was formed. These cells (mononucleated cells formed by Msx1 induced dedifferentiation of myotubes [MIDCs]) were then cultured in the growth medium supplemented with Leukemia Inhibitory Factor (LIF) 100 µl/10 ml on noncoated plates. Real-Time Microscopic Image Acquisition of Myotube Dedifferentiation For microscopic time-lapse studies on a 37°C heated stage (Omega Engineering, Inc., Stamford, CT, http://www.omega.com/?gclid=CLnyptO3n74CFUiEfgodiSwABQ), 100–200 myotubes were individually placed in a 35 mm collagen-coated dish and cultured in HEPES buffered Hams F-10 media with no HCO3 (Invitrogen) having a mineral oil lid. For studies in an on-stage mini-CO2 incubator (Pathology Devices, Westminster, MD, http://www.pathologydevices.com/), myotubes were plated in collagen-coated 48-well plastic dishes in standard growth media. Imaging was accomplished on an automated Nikon TE2000E inverted fluorescence microscope (Nikon U.S., Inc., Melville, NY, http://www.nikonusa.com/). Excitation illumination was supplied by an X-Cite mercury lamp (Lumen Dynamics, Mississauga, Canada, http://www.ldgi.com/x-cite/) and attenuated by a neutral density filter. Images were captured on a Roper HQ CCD camera (Photometrics, Inc., Tucson, AZ, http://www.photometrics.com/) using 4 × 4 binning for fluorescence and 2 × 2 binning for phase contrast images. IPLab software (BioVision, Inc., Exton, PA, http://www.biovision.com/) controlled automated image capture of 200–400 ×20 fields at each time point. Time-lapse image acquisition was begun 8–12 hours after induction of msx-1 expression and continued for 25–66 hours at 30-minute intervals and was analyzed using IPLab software. Multipotency Assays of MIDCs In Vitro For adipogenic assays, cells were cultured in conditions described previously [16] and then subjected to Oil Red labeling. In osteogenic assays, cells were cultured in conditions described elsewhere without addition of BMP2 [17] and then stained for alkaline phosphatase. In chondrogenic assays, cells were seeded on PEG polymer and cultured in the medium described previously [18] and examined for the presence of collagen II. For the myogenic assay, cells were cultured in differentiation medium for 4 days followed by immunocytochemistry for the expression of myosin heavy chain. During all these assays, the expression of Msx1 was suppressed using doxycycline beginning at day 0 for each assay. Gene Expression Analysis by RT-PCR, Western Blot, Immunocytochemistry, and Immunohistochemistry For RT-PCR, total RNA was extracted using a GenElute Mammalian Total RNA Miniprep kit (Sigma, Sigma Aldrich, St. Louis, MO, http://www.sigmaaldrich.com/united-states.html) and reverse-transcribed into cDNA using TaqMan Reverse Transcription Reagents (Roche, Roche Diagnostics, Indianapolis, IN, http://lifescience.roche.com/). Primer sequences are listed in Supporting Information Figure S18. For qRT-PCR, SYBR Premix Ex Taq Kit (TaKaRa, Shiga, Japan, http://www.clontech.com/takara/US/Support/) on ABI 7300 PCR and Detection System (Applied Biosystems, Molecular Biosystems, Grand Island, NY, www.lifetechnologies.com/us/en/home/brands/applied-biosystems.html) were used for qRT-PCR. Mouse GAPDH was amplified as an internal standard. The relative values were calculated using ΔΔCt method and normalized against endogenous GAPDH. Primer sequences are listed in Supporting Information Figure S19. For Western blot analysis, total proteins were extracted and expression levels of Pax3, Pax7, Myf5, and MyoD were measured using antibodies of anti-Pax3 (MAB2457, R&D, MN, http://www.rndsystems.com/product_results.aspx?k=MAB2457), Pax7 (Contributed by Kawakami, Hybridoma Bank, Developmental Hybridoma Studies Bank, Iowa City, IO, http://dshb.biology.uiowa.edu/), MyoD (Sc-760, Santa Cruz, Santa Cruz Biotechnology, Santa Cruz, California) in 1:1,000, and Myf5 (ARP32134, Avivasysbio, Aviva Systems Biology, San Diego, CA, http://www.avivasysbio.com) in 1:2,000 dilutions. For immunocyto- and immunohisto-chemistry myotubes were immunostained with anti-mouse Myosin Heavy Chain (MEDCLA66, Accurate Chemical & Scientific Corp, Westbury, NY, http://www.accuratechemical.com/) followed by incubation with a CyTm3-conjugated secondary antibody (Jackson Lab, Jackson Immuno Research, West Grove, PA, http://www.jacksonimmuno.com/) (all secondary antibodies here and below were used at a dilution of 1:500). Muscle sections from injected mdx mice were incubated with anti-dystrophin (ab15277, Abcam, Cambridge, MA, http://www.abcam.com/) in 1:500 dilution followed by incubation with a CyTm3-conjugated secondary antibody. Muscle sections of SCID mice were incubated with mouse-anti-Pax7 antibody in 1:50 dilution followed by the CyTm3-conjugated secondary antibody and then double labeled with rabbit-anti-Laminin (Sigma, MO) in 1:500 followed by incubation with Alexa-488-conjugated secondary antibody (Invitrogen). Cell Transplantation, In Vivo Fluorescence Imaging, and Tissue Preparation For cell transplantation, the cultured MIDCs and myoblasts were trypsinized and collected. Animal experiments were carried under the guidance and approval of the Harvard Medical School standing committee on animals. Three days prior to cell transplantation, the hind legs of 4–8-week-old mdx mice (Jackson Lab) were given 18 Gy irradiation. Animals were anesthetized with Ketamine/Xylazine (100 mg/15 mg in 5 ml saline) at a dosage of 100 ml/20 g of b.wt. and then 1 × 105 MIDCs or the same number of control myoblasts in 10 µl Hanks’ balanced saline solution were injected into tibialis anterior (TA) muscles of mdx mice at three positions. Similarly, 1 × 105 eGFP-labeled MIDCs were injected into the TA muscles of unirradiated 8–12-week-old SCID mice or nude mice (Taconic). The TA muscles of SCID mice were injured with cardiotoxin in a dosage 10 ng/muscle 3 days prior to cell injection. eGFP-labeled MIDCs (1 × 105) or control myoblasts were injected subcutaneously at the thigh reign of SCID mice at one position. At 60 days post-cell injection, cardiotoxin (10 ng/leg, Sigma) was injected to selectively destroy myofibers formed by injected cells. All the animals were given doxycycline 1 mg/ml starting 3 days prior to cell transplantation. Suppression of Msx1 after transplantation was confirmed by RT-PCR (Supporting Information Fig. S17). The fate of eGFP-labeled cells was monitored by in vivo fluorescence imaging periodically. For in vivo fluorescence imaging, the injected SCID mice were anesthetized as described above. The imaging procedure and parameters were described as our previous report [19]. Mid-TA cross-sections and in some cases serial cross-sectional tissue samples were harvested at 2, 4, 8, and 12 weeks post-cell injection and in age matched uninjected muscles from the 12-week group for histology analysis including analysis of whether donor-derived cells formed fat, cartilage, or osteogenic tissue. Mice were deeply anesthetized with xylazine and ketamine and had their left ventricles perfused with Phosphate Buffered Saline (PBS) and 4% (wt/vol) paraformaldehyde (Sigma). The TA muscles and sites of the subcutaneous engraftment were then carefully removed, postfixed in 4% Paraformaldehyde (PFA) over 12 hours, submersed in 30% sucrose overnight, frozen in OCT embedding compound, and Cryo-sectioned coronally (10 µm). Cryo-sections were thaw mounted onto gelatin-coated slides and stored at −20°C. Statistical Analysis Quantitative data are presented as mean ± SD. A Student’s t test (assuming equal variances) was performed to determine the statistical significance between two experimental groups. A p value less than .05 was considered to be statistically significant. Results Ectopic Overexpression of Msx1 Reprograms Terminally Differentiated Mouse Myotubes to Become Proliferating Mononuclear Cells We first explored whether differentiated mouse myotubes could be induced to reenter the cell cycle and undergo cellularization to form proliferating mononuclear cells by expressing Msx1. Primary myoblasts were transduced with retrovirus where either Msx1 expression was under the control of a tetracycline-off promoter or the vector containing a reversed Msx1 insert (Supporting Information Fig. S1). Permanently transduced clonal cells were expanded and differentiated into myotubes in the presence of doxycycline. Single myotubes were then isolated and cultured in either liquid or semisolid medium after exclusion of all mononucleated cells (Supporting Information Fig. S2). Analysis of individual myotubes over time revealed that 1%–5% of these myotubes produced proliferating mononucleated cells in 1–5 days in both types of media, after initiation of the expression of Msx1 by removal of doxycycline and addition of growth media. No mononuclear cells were produced by myotubes expressing the reversed vector or that were not transduced (Supporting Information Fig. S3, Table l). Using static fluorescence microscopy (Fig. 1A) and real-time microscopic imaging, we were able to capture the detailed kinetics of transformation as Msx1 transduced myotubes underwent cellularization (Supporting Information Movies S4–S6), and confirmed that this does not occur in control myotubes (Supporting Information Movie S7). Analysis of these recordings revealed that Msx1-induced cellularization results from extension of the leading edge of the myotube cells accompanied by nuclear migration, and finally by pinching off of a small portion of the cytoplasm containing a single nucleus. These newly formed mononucleated cells can translocate along the surface of existing multinucleated myotubes, or move freely over the culture substrate as single cells (Supporting Information Movies S4, S6). Some of them also can undergo repeated rounds of cell division and form new mononucleated daughter cells that retain the ability to migrate individually (Supporting Information Movies S5, S6). These results provide unequivocal evidence that expression of Msx1 can induce primary mouse myotubes to revert into proliferating mononuclear cells, as previously observed in myotubes formed by immortalized C2C12 cells [7, 8]. Open in new tabDownload slide Evidence of Msx-1-induced dedifferentiation. (A): Representative pictures of the kinetics of cellularization of Msx1-expressing myotubes. Cellularization begins 24 hours after induction of Msx-1 expression. This continues through day 2 and daughter cell expansion begins. By day 3, cellularization is complete and daughter cell expansion continues, with a doubling time of less than 24 hours and a clone of daughter cells is ready for further expansion by day 5. For real-time observations over 49.5 hours see Supporting Information Movie S5. (B): Western blot showing the expression levels of MyoD, Myf5, Pax7, and Pax3 in control myoblasts (Cont Mb), Msx1-expressing myoblasts (Msx1-Mb), and two MIDC clones (MIDC-C1 and C2). (C): Quantitative Western blot analysis normalized with GAPDH. *, p < .05 and ω, p < .01 versus Cont-Mb Student’s t test. n = 3–4. (D): Myotubes formed by Cont-Mbs and MIDCs stained with adult myosin heavy chain (MyHC) antibody. Scale bar = 100 µm. Abbreviations: MIDC, mononucleated cells formed by Msx1 induced dedifferentiation of myotubes; PM, Primary Myoblasts. Open in new tabDownload slide Evidence of Msx-1-induced dedifferentiation. (A): Representative pictures of the kinetics of cellularization of Msx1-expressing myotubes. Cellularization begins 24 hours after induction of Msx-1 expression. This continues through day 2 and daughter cell expansion begins. By day 3, cellularization is complete and daughter cell expansion continues, with a doubling time of less than 24 hours and a clone of daughter cells is ready for further expansion by day 5. For real-time observations over 49.5 hours see Supporting Information Movie S5. (B): Western blot showing the expression levels of MyoD, Myf5, Pax7, and Pax3 in control myoblasts (Cont Mb), Msx1-expressing myoblasts (Msx1-Mb), and two MIDC clones (MIDC-C1 and C2). (C): Quantitative Western blot analysis normalized with GAPDH. *, p < .05 and ω, p < .01 versus Cont-Mb Student’s t test. n = 3–4. (D): Myotubes formed by Cont-Mbs and MIDCs stained with adult myosin heavy chain (MyHC) antibody. Scale bar = 100 µm. Abbreviations: MIDC, mononucleated cells formed by Msx1 induced dedifferentiation of myotubes; PM, Primary Myoblasts. Dedifferentiated Mononuclear Cells Reactivate Embryonic Muscle Progenitor Genes Comparison of gene expression profiles of MIDCs with their both their parental myoblasts (where Msx1 expression was turned on but cells had not undergone differentiation/dedifferentiation) and control primary myoblasts demonstrated that the MIDCs exhibit a distinct phenotype. The myogenic determinant factor MyoD was downregulated in MIDCs, whereas genes expressed at high levels in earlier stage myogenic progenitor cells, such as Myf5, Pax3, and Pax7, were upregulated (Fig. 1B, 1C). Interestingly, expression of Msx1 in undifferentiated primary myoblasts also upregulated these genes but to a lesser extent than the levels found in MIDCs (Fig. 1B, 1C). MIDCs also expressed the progenitor cell marker Sox2, which is not expressed in primary myoblasts or myoblasts expressing Msx1 (Supporting Information Fig. S8). Intriguingly, some MIDC clones transiently expressed low levels of pluripotent genes, Nanog, Oct4, and Rex1 but their expression diminished over time with passaging (Supporting Information Fig. S8). With addition of specific growth factors MIDCs could be induced to redifferentiate into cells with adipogenic, chondrogenic, or osteogenic characteristics, while primary myoblasts could not (Supporting Information Fig. S9). Attempts to induce forward differentiation of MIDCs into hepatocytes and neuronal cells failed, suggesting that their differentiation potential is restricted to mesodermal derivatives. Most importantly, when MIDCs were cultured under conditions (low concentration of serum and free of growth factors) that are used to differentiate primary myoblasts, they uniformly differentiated into multinucleated myotubes as indicated by their expression of myosin heavy chain (MyHC) (Fig.1D). MIDCs Only Generate Muscle Tissue In Vivo To determine whether MIDCs are capable of forming de novo muscle tissue in vivo we injected eGFP-labeled MIDCs or control myoblasts subcutaneously in the thigh region of SCID mice (n = 6/group). Using in vivo fluorescence imaging, we observed that the eGFP-signal reached a peak around 4 weeks after mice were injected with MIDCs (Fig. 2A). To prove that the e-GFP+ donor-derived cells were myofibers, 2 months after cell injection, we injected cardiotoxin into the eGFP+ region and followed the change by in vivo fluorescence imaging. After cardiotoxin injection, we observed a rapid decline in the eGFP signal, which was followed by recovery of the fluorescence after 2 weeks (Fig. 2A and Supporting Information Fig. S10). Immunohistochemical analysis confirmed that the injected MIDCs formed striated MyHC+ myofibers within the subcutaneous space, almost all of which had peripherally located myonuclei (Fig. 2C, 2D and Supporting Information Fig. S11). We found no donor-derived cells having migrated into the muscle below nor any donor-derived cells that were immuno-positive for adipogenic, osteogenic or chondrogenic markers, or T-cell factor 4 [20], a gene expressed by mesoderm cells other than muscle during embryonic development (data not shown). In contrast, in mice injected with control myoblasts, the eGFP signal rapidly declined and at 60 days postinjection was completely absent (Fig. 2B). The green fluorescent protein (GFP) signal could not be recovered in response to cardiotoxin injection and post-mortem histological examination revealed no eGFP+ ectopic muscle in these mice. Open in new tabDownload slide Generation of myofibers in an ectopic location. (A, B): Representative in vivo fluorescence images after subcutaneous transplantation of eGFP-labeled mononucleated cells formed by Msx1 induced dedifferentiation of myotubes (MIDCs) (A) or Cont-Mbs (B). (C): Longitudinal section of a MIDC-derived subcutaneous muscle harvested 3 months post-cell-injection. Upper panel, low magnification. Lower panel, high magnification of the framed area. Dual staining with MyHC and DAPI (for the location of myonuclei). (D): Cross-section of 3-month MIDC-derived subcutaneous muscle. The right panels are high magnification of the framed area in left picture. The right most picture demonstrates peripheral myonuclei. Scale bars = 50 µm. Abbreviations: GFP, green fluorescent protein; MyHC, myosin heavy chain. Open in new tabDownload slide Generation of myofibers in an ectopic location. (A, B): Representative in vivo fluorescence images after subcutaneous transplantation of eGFP-labeled mononucleated cells formed by Msx1 induced dedifferentiation of myotubes (MIDCs) (A) or Cont-Mbs (B). (C): Longitudinal section of a MIDC-derived subcutaneous muscle harvested 3 months post-cell-injection. Upper panel, low magnification. Lower panel, high magnification of the framed area. Dual staining with MyHC and DAPI (for the location of myonuclei). (D): Cross-section of 3-month MIDC-derived subcutaneous muscle. The right panels are high magnification of the framed area in left picture. The right most picture demonstrates peripheral myonuclei. Scale bars = 50 µm. Abbreviations: GFP, green fluorescent protein; MyHC, myosin heavy chain. MIDCs Generate Myofibers in a Niche and Time-Dependent Manner To assess whether MIDCs engraft into already formed adult muscle, we injected MIDCs or control myoblasts intramuscularly into preirradiated TA muscles of dystrophin-deficient mdx mice. In muscles harvested at 2, 4, and 8 weeks following injection, we detected an average of 166 ± 12, 1,283 ± 158, and 3,340 ± 304 (n = 5) dystrophin+ myofibers in each MIDC-injected muscle analyzed, whereas only 10 ± 3 (n = 5) dystrophin+ fibers were observed in muscles injected with control myoblasts at 2 weeks and this number did not increase with time (Fig. 3A). The dystrophin+ myofibers in the MIDC-injected group varied widely in size and formed clusters of dystrophin positive fibers. Although some of them had the size of host fibers, the majority of the fibers were small at earlier time points and some of them did not align properly. To further quantify the degree of regeneration by donor-derived fibers, in a separate group, we injected MIDCs into irradiated mdx TA muscles and analyzed dystrophin+ muscle fibers at 4 and 12 weeks postinjection and compared the 12-week samples with cross-sections of age matched mdx control muscles. We measured the total cross-sectional area (CSA) occupied by dystrophin+ fibers and found that the %CSA occupied by dystrophin+ fibers increased significantly in the 12-week animals (44.2% ± 3.7%, n = 3) versus 4-week animals (26.6 ± 2.9, n = 3, p < .02 vs. 12-week group). We also measured and analyzed the CSA of individual dystrophin+ fibers in both groups and found a significant increase in the number of large dystrophin+ fibers and improvement in the alignment of dystrophin+ fibers in the 8- and 12-week groups (Fig. 3B, 3C, Supporting Information Figs. S12, S13). Interestingly, whereas the majority of dystrophin− host fibers (52% ± 15%, 168/337) and dystrophin+ fibers in Primary Myoblast (PM)-injected muscles (92%, 60/65) have centrally located myonuclei, only a small percentage of dystrophin+ fibers in MIDC injected muscles have centrally located nuclei (2.4% ± 1%, 65/2,658, p < .01 vs. both untreated and PM-treated muscles) (Fig. 3A and Supporting Information Fig. S14). Open in new tabDownload slide De novo fiber formation in preirradiated mdx muscle. (A): Merged dystrophin and DAPI staining pictures of MIDC- (top) and Cont-Mb- (bottom) injected muscles harvested at 2, 4, and 8 weeks. (B): Sections showing the size and number of dystrophin+ fibers in MIDC-injected muscles harvested at 4 and 12 weeks post-cell-injection. (C): Quantitative analysis of fiber size distribution in MIDC-injected muscles harvested at 4 and 12 weeks post-cell-injection (n = 3 muscles). Scale bars = 100 µm. Abbreviation: MIDC, mononucleated cells formed by Msx1 induced dedifferentiation of myotubes. Cont Mb, Control Myoblasts. Open in new tabDownload slide De novo fiber formation in preirradiated mdx muscle. (A): Merged dystrophin and DAPI staining pictures of MIDC- (top) and Cont-Mb- (bottom) injected muscles harvested at 2, 4, and 8 weeks. (B): Sections showing the size and number of dystrophin+ fibers in MIDC-injected muscles harvested at 4 and 12 weeks post-cell-injection. (C): Quantitative analysis of fiber size distribution in MIDC-injected muscles harvested at 4 and 12 weeks post-cell-injection (n = 3 muscles). Scale bars = 100 µm. Abbreviation: MIDC, mononucleated cells formed by Msx1 induced dedifferentiation of myotubes. Cont Mb, Control Myoblasts. At 12 weeks post-cell-injection we counted the number of total myofibers from the mid cross-section of each TA muscle harvested. There were 5,345 ± 373 (n = 3) fibers in MIDC-injected TA muscles of which (4,321 ± 491) are dystrophin+, and there were 2,899 ± 273 (n = 3) in control TA muscles (Supporting Information Fig. S15) indicative of new fiber growth in the MIDC injected muscles. When GFP-labeled MIDCs were injected into the muscles of SCID (Fig. 4A, 4B) or nude (Supporting Information Fig. S16) mice that have no degeneration/regeneration signals, the GFP signal quickly reached a plateau and remained stable up to 6 months, the longest time point tested. In addition to eGFP+ myofibers, eGFP+/Pax7+ mononuclear cells could be detected in muscles transplanted with eGFP-labeled MIDCs when measured at 2 months. The majority of these eGFP+/Pax7+ cells (75.8%, 232/306) were located beneath the myofiber basement membrane (the typical location of satellite cells) (Fig. 4C). The remaining cells (24.2%, 74/306) appeared to be in the interstitium encircled by their own basement membrane (Fig. 4D). Open in new tabDownload slide Evidence of environmentally regulated mononucleated cells formed by Msx1 induced dedifferentiation of myotubes (MIDC) regeneration and satellite cell formation. (A): Representative in vivo fluorescence images of eGFP-labeled MIDCs transplanted in SCID tibialis anterior muscles injected with cardiotoxin 3 days prior to cell-transplantation. (B): Graph of fluorescence signal intensity over time. n = 4. Baseline fluorescence prior to injection was null. (C): eGFP+/Pax7+ cells residing underneath basal membrane. (D): An eGFP+/Pax7+ cell residing in the interstitium enclosed by its own basal membrane. Scale bar = 20 µm. Lam; Laminin. Open in new tabDownload slide Evidence of environmentally regulated mononucleated cells formed by Msx1 induced dedifferentiation of myotubes (MIDC) regeneration and satellite cell formation. (A): Representative in vivo fluorescence images of eGFP-labeled MIDCs transplanted in SCID tibialis anterior muscles injected with cardiotoxin 3 days prior to cell-transplantation. (B): Graph of fluorescence signal intensity over time. n = 4. Baseline fluorescence prior to injection was null. (C): eGFP+/Pax7+ cells residing underneath basal membrane. (D): An eGFP+/Pax7+ cell residing in the interstitium enclosed by its own basal membrane. Scale bar = 20 µm. Lam; Laminin. Discussion In this study, by conditionally expressing the transcription factor Msx1 in primary mouse myotubes, we found that they could be induced to re-enter the cell cycle and fragment into mononuclear cells that were lineage-specific proliferating myogenic progenitors. Furthermore, these dedifferentiated mononuclear cells displayed robust, myogenic progenitor-like, highly specialized differentiation potential following in vivo transplantation, indicating that our novel approach may have translational impact toward combating degenerative muscle disorders and volumetric muscle loss. Previous studies have already demonstrated that dedifferentiation can efficiently regenerate impaired organs including fin, heart, and so forth, in zebrafish [1, 21-23] and lost limbs in salamanders [5]. At least in the case of newt myofibers this natural process of regeneration is dependent on the induction of Msx1 expression [6]. In the past few years, several examples of dedifferentiation in mammals have been verified either from in vitro or in vivo experiments [24-26]. A recent study shows that fate-restricted progenitor populations expressing Msx1 are responsible for regenerating the amputated digit tip in vivo in mice [27], which with our results provide evidence that mammals can harness a muscle regeneration strategy if the same Msx1 pathway used by urodele amphibians can be activated. Mammalian embryonic myogenesis and postnatal muscle repair are two distinct and well-defined processes. During myogenesis, the majority of muscles including limb muscles are formed by Pax3+ embryonic myogenic progenitors and Pax3+/Pax7+ fetal myogenic progenitors [28-30]. With rare exceptions, all the myofibers that build up an adult muscle are formed during embryonic myogenesis [31, 32]. Postnatal growth and repair of mammalian muscle mainly rely on mobilization of satellite cells, the adult form of myogenic progenitors [33]. Unlike embryonic myogenic progenitors, satellite cells, although capable of repairing damaged myofibers, appear to have a limited capacity to initiate new fiber formation, partly due to their specific location. Satellite cells are located beneath the basal membrane that wraps the surface of each myofiber [34]. In response to various destructive assaults on myofibers, satellite cells activate, proliferate, and fuse to repair the partially or completely damaged myofiber [35, 36] and, at the same time, attain self-renewal to repopulate a satellite cell pool [37-40]. The process of satellite cell-mediated regeneration is rapid and efficient. Thus, as a collective result of individual fiber replacement, skeletal muscles have an impressive ability to repair themselves after damage. The regeneration of damaged fibers by satellite cells is referred as a tissue level of muscle regeneration by Carlson [41] to distinguish it from the organ regeneration of a partially or entirely lost muscle seen in some urodele amphibians and other lower organisms. Studies have shown that despite the fact that satellite cells of mammalian can migrate beneath the basal membrane of the myofiber with which they are associated [19, 42-44], they can only migrate across the basal membrane to regenerate other fibers during the developmental period when the basal membrane is still fragile, or in some diseases and in extreme injury conditions [45, 46]. Thus, myofiber regeneration occurs by expansion of satellite cell progeny within the original empty endomysial tubes and very few, if any, new fibers are ever formed as a result of de novo regeneration by satellite cells [47-49]. Regeneration of a lost or even partially lost muscle by satellite cells has not been demonstrated in mammals. Strategies to generate artificial muscles using satellite cells and their progeny have had very limited if any success which has posed a major obstacle to achieving that goal [50-57]. Animal experiments show that transplantation of the entire content of a muscle that has been removed, minced, and placed back to its original muscle bed resulted in regeneration of only ∼30% of the original muscle in a somewhat discordant fashion. Due to this limited or lack of ability for de novo fiber formation, mammals respond to growth demand by hypertrophy instead of hyperplasia [58]. Transplantation of primary myoblasts, in both uninjured muscle and muscle injured with cardiotoxin, results in an initial formation of the donor-derived fibers by fusion with each other or with host myofibers. Once this fusion has occurred, however, the number of donor-derived fibers either remains unchanged over time or decreases because of immuno-rejection [59-61]. Although using highly purified subpopulations of satellite cells or freshly isolated satellite cells to engraft muscle have achieved much more robust regeneration compared with cultured myoblasts, the majority of engrafted fibers (98%) are donor-host hybrids and no report has shown an increase in the number of donor-derived myofibers after initial engraftment [62] by isolated satellite cells unlike our findings in this study. In addition, it has been shown that the transplantation of the most primitive satellite cells (Pax7+/Myf5−) results in extensive repopulation of the satellite cell niche without interstitial retention and new fiber formation [37]. Collectively, these experiments demonstrate that isolated satellite cells have a strong tendency to home to and remain in their native niches and thereafter behave no differently after engraftment than native satellite cells. Thus, it is likely that the limited capability of satellite cells to form de novo myofibers is responsible for the inability of mammals to regenerate lost muscle. In contrast to the tissue level of muscle regeneration, in organ level muscle regeneration all the myofibers regenerate de novo [41]. Organ level of muscle regeneration has only been observed in amphibians and other lower organisms such as zebra fish. In organ level muscle regeneration, new muscles are formed in coordination with the regeneration of surrounding structures such as bones, cartilages, tendons, nerves, and blood vessels. Interestingly, it is noticed that myofibers formed during organ level of muscle regeneration have peripheral located nuclei [41] unlike myofibers regenerated by satellite cells which are initially central-nucleated in many species including human [63] and remain central-nucleated in rodents indefinitely [64]. The cell sources of organ level of muscle regeneration have been the subject of intense debate [65]. Studies using different organisms yielded diverse results [23, 66]. A recent study using Cre-loxP-based genetic fate mapping technique has shown that even two closely related salamander species use different cell sources for muscle regeneration and confirmed that in the newt multinuclear muscle cells do revert back to proliferating mononuclear cells after amputation and subsequently contribute to muscle regeneration but in axolotl it is regenerated by resident Pax7+ satellite cells [4]. It has been established by several groups that postmitotic multinuclear mammalian muscle cells can be induced to become proliferating mononuclear cells [9-11]. Whether these dedifferentiated mammalian muscle cells simply regain proliferating property and take one step back revert to the cells that form postmitotic muscles or whether they obtain the myogenic capability of earlier progenitors or the regeneration capability of dedifferentiated newt muscle cells is a critical question for the applicability of dedifferentiation approach to the regenerative medicine. In this study, we provide evidence for the first time showing that at least in the case of Msx1-induced dedifferentiation, dedifferentiated cells reactivated genes expressed by embryonic myogenic progenitors and display a degree of myogenic regenerative capability only observed in embryonic/fetal myogenic progenitors and in dedifferentiated newt muscle cells. Remarkably, in preirradiated mdx muscles, injected MIDCs gradually regenerated 300 times more myofibers over a 3-month-period than were generated in muscles injected with their parental satellite cell-derived myoblasts, and that over 12 weeks time these MIDCs donor-derived fibers made up as much as half of the CSA of the transplanted muscles and increased the total number of fibers per CSA 1.8-fold. Despite the fact that the dystrophin+ fibers occupy only half of the total cross-area they account for 80% of the total number of fiber in the total cross-section. This is because many new fibers still remain smaller than normal. Given the fact that the increase in the total number of myofibers coincides with the decrease in the number of dystrophin− host fibers in the MIDC-injected muscle, from 2,900 fibers in the uninjected muscle to 1,024 fibers/muscle, it is likely that MIDCs have both fused with the existing host fibers and formed de novo fibers (Supporting Information Fig. S11). The unprecedented regenerative power exhibited by MIDCs suggests that if proper cues and niches including supporting structures are provided, these reprogrammed muscle progenitors could potentially reconstruct a fully functional muscle. This offers an attractive therapeutic strategy for treatment of genetic myopathies and volumetric muscle loss due to severe trauma or surgery with ex vivo treated and expanded autologous cells [51, 52, 67, 68]. Conclusions Although why this regenerative pathway is activated by injury in urodele amphibians, but rarely if ever in mammals, remains unknown. Whether our findings can be extended to mammalian tissues other than skeletal muscle is also unclear. Nevertheless, we demonstrate that it is possible to directly reprogram terminally differentiated mammalian cells back into a lineage-specific progenitor state, rather than having to induce them to pass through a pluripotent state or to transit from one adult state to another. Our studies demonstrate that MIDCs are capable of de novo fiber formation when injected in an ectopic location and the increase in dystrophin+ fibers over time in irradiated mdx muscles suggests that de novo fiber formation is also possible in the context of a degenerating muscle although these data do not conclusively prove this. These data shift the current paradigm of mammalian cell reprogramming, and have the potential to provide a shortcut for the regenerative medicine as well. Acknowledgments We thank W. Liu, R. Hirsh, J. Bell, and L. Yang for their technical help. This work was supported by grants from NIH/NIAMS (5KO2AR051181), MDA, and Harvard Stem Cell institute to Y. Wang, NIH/NIAMS (5P01 AR052354) to P.D. Allen. Paul D. Allen is currently affiliated with the Department of Molecular Biosciences, School of Veterinary Medicine, University of California at Davis, Davis, CA. NSF 0958345 to X. Xu. Author Contributions Z.Y. and Y.W.: designed overall experimental strategies, performed experiments and analyzed data, wrote the manuscript, and commented on the manuscript; Q.L.: designed overall experimental strategies, performed experiments and analyzed data, and commented on the manuscript; P.D.A.: designed overall experimental strategies, wrote the manuscript, and commented on the manuscript; R.J.M.: developed strategies for real-time microscopic imaging experiments and performed the experiments, analyzed real-time microscopic imaging results, and commented on the manuscript; D.E.I.: analyzed real-time microscopic imaging results, wrote the manuscript, and commented on the manuscript; X.X.: performed in vivo fluorescence imaging experiments, analyzed the results, and commented on the manuscript; H.L.: performed gene expression experiments, participated in data analysis, and commented on the manuscript; Z.M.: performed myotube dedifferentiation experiments, developed the protocol for analyzing real-time microscopic imaging results, and commented on the manuscript. 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TI - Mononuclear Cells from Dedifferentiation of Mouse Myotubes Display Remarkable Regenerative Capability
JF - Stem Cells
DO - 10.1002/stem.1742
DA - 2014-09-01
UR - https://www.deepdyve.com/lp/oxford-university-press/mononuclear-cells-from-dedifferentiation-of-mouse-myotubes-display-mCFcPltA42
SP - 2492
EP - 2501
VL - 32
IS - 9
DP - DeepDyve
ER -