Functional Recovery of Damaged Skeletal Muscle Through Synchronized Vasculogenesis, Myogenesis, and Neurogenesis by Muscle-Derived Stem Cells
Background— Recent studies have shown that skeletal muscle–derived stem cells (MDSCs) can give rise to several cell lineages after transplantation. However, the potential therapeutic uses of MDSCs, the functional significance of the transplanted tissue, and vasculogenesis, myogenesis, and reconstitution of other tissues have yet to be investigated in detail. In addition, the relationship between MDSCs and mesenchymal bone marrow cells is of interest.
Methods and Results— We developed a severe-damage model of mouse tibialis anterior muscle with a large deficit of nerve fibers, muscle fibers, and blood vessels. We investigated the potential therapeutic use of freshly isolated CD34+/45− (Sk-34) cells. Results showed that, after transplantation, implanted cells give rise to myogenic, vascular (pericytes, vascular smooth muscle cells, and endothelial cells), and neural (Schwann) cells, as well as contributing to the synchronized reconstitution of blood vessels, muscle fibers, and peripheral nerves, with significant recovery of both mass and contractile function after transplantation. Investigation of Sk-34 cell transplantation to the renal capsule (nonmuscle tissue) and fluorescence in situ hybridization analysis for the transplanted muscle detecting the Y chromosome revealed the intrinsic plasticity of the Sk-34 cell population. In addition, there were no donor-derived Sk-34 cells in the muscle of lethally irradiated bone marrow–transplanted animals, indicating that the Sk-34 cells were not derived from bone marrow.
Conclusions— These findings indicate that freshly isolated skeletal muscle–derived Sk-34 cells are potentially useful for reconstitution therapy of the vascular, muscular, and peripheral nervous systems. These results provide new insights into somatic stem and/or progenitor cells with regard to vasculogenesis, myogenesis, and neurogenesis.
Received April 9, 2005; revision received August 4, 2005; accepted August 8, 2005.
There is widespread hope that the multipotency of tissue-derived stem cells will facilitate tissue regeneration and complete reconstitution of the organic components of the circulatory, skeletal, and nervous systems. For this purpose, muscle-derived stem cells (MDSCs) have been identified by use of a variety of isolation methods.1–14 In previous reports, “myo-endothelial progenitor cells” were found to differentiate into myogenic, vasculogenic, and other cells of mesodermal lineage, such as adipogenic cells, in the interstitial spaces of skeletal muscle. Fractionated by FACS on the basis of cell-surface antigens, cells in the CD34+/CD45− (designated Sk-34 cells) fraction exhibited colony formation and the potential to differentiate into mesodermal cells, such as endothelial cells, myogenic cells, and adipocytes, both in vitro and in vivo.9 We also identified cells in the CD34−/45− (double negative: Sk-DN) fraction as a putative cell population that includes further immature stem cells; that is able to form clonal sphere-like colonies in a collagen-based cell culture system with basic fibroblast growth factor and endothelial growth factor (EGF); and that exhibits the potential to differentiate into myogenic and endothelial cells after 10 days of culture.10 These findings suggest that myo-endothelial progenitors, such as Sk-34 and Sk-DN cells, residing in the interstitial spaces of skeletal muscle, can potentially contribute to postnatal myogenesis and vasculogenesis after muscle growth and/or muscle hypertrophy.
Because of their myogenic potency, MDSCs have been injected into the blood circulation of a mouse muscular dystrophy model (mdx), but donor-derived cell incorporation into muscle and the restoration of dystrophin expression in the affected muscle were limited.3,11 Although these therapeutic applications of MDSCs for traumatic, ischemic, degenerative, or infectious diseases seem promising, there are several determinant issues that remain unresolved: (1) the particular cell population to use, (2) the precise cell transplantation methodologies, (3) determining stem cell fate and organization in vivo after transplantation through line-by-line scrutiny, and (4) the functional significance of the reconstituted tissue.
To address these points, we first verified the hypothesis that primary or cultured Sk-34 cells are practical, stem cell–enriched populations for muscle regeneration by investigating the potential for new fiber formation and neovascularization, which would result in significant functional (contractile) recovery. The intrinsic plasticity of Sk-34 cells (not through cell fusion) was also confirmed by fluorescence in situ hybridization (FISH) analysis for the transplanted muscles and transplantation of cells into nonmuscle tissue (under the renal capsule). In addition, the bone marrow derivation of Sk-34 cells was examined by bone marrow transplantation, because myo-endothelial progenitor cells have been also identified in umbilical cord blood.15
Green fluorescent protein transgenic mice (GFP-Tg mice; C57BL/6 TgN[act EGFP]Osb Y01) were provided by Dr M. Okabe (Osaka University, Osaka, Japan) and were used in the cell transplantation studies as donor mice,16 whereas wild-type C57BL/6 mice were used as recipients. All experimental procedures were conducted in accordance with the Japanese Physiological Society Guidelines for the Care and Use of Laboratory Animals, as approved by the Tokai University School of Medicine Committee on Animal Care and Use.
Interstitial cells were extracted from the thigh and lower leg muscles (tibialis anterior [TA], extensor digitorum longus, soleus, plantaris, gastrocnemius, and quadriceps femoris) of 3- to 8-week-old GFP-Tg mice by use of a method for isolating intact, living individual muscle fibers associated with satellite cells, as described previously.9,10 Briefly, whole muscles were treated with 0.1% collagenase type IA (Sigma-Aldrich) in Dulbecco’s modified Eagle’s medium (DMEM) containing 5% fetal calf serum (FCS) with gentle agitation for 2 hours at 37°C. Extracted cells were filtered through 70-, 40-, and 20-μm nylon meshes to remove muscle fibers and debris, followed by washing and resuspension in Iscove’s modified Dulbecco’s medium (IMDM) containing 10% FCS, thus yielding enzymatically extracted cells (EECs). EECs were stained with biotin-conjugated anti-mouse CD34 (RAM34) and streptavidin-allophycocyanin (APC) and phycoerythrin (PE)-conjugated anti-mouse CD45 (30-F11). Dead cells were stained with propidium iodide (PI). CD34+/45− (Sk-34) and CD34−/45− (DN) cells were sorted and counted after PI-positive cells were excluded. All antibodies were purchased from PharMingen. Cell analysis and sorting were performed by use of a triple laser FACS Vantage (Becton Dickinson).
Conditions and Preparation of Cells for Transplantation
Three sets of cell transplantation conditions and 1 control were used: (1) Sk-34 cells transplanted immediately after sorting (Sk-34-0 group); (2) Sk-34 cells transplanted after 1 day of culture (Sk-34-1); (3) Sk-34 cells transplanted after 5 days of culture (Sk-34-5); and (4) DMEM solution injected without cells (control group). The expansion and induction of purified Sk-34 cells were performed on the basis of methods reported previously.9,10 Cells were first cultured in liquid IMDM with 20% FBS for 2 days. The medium was then changed to 0.5% methylcellulose with 5% FBS/IMDM without cytokines, and the cells were cultured for 3 more days.
Severe Muscle Damage Model and Cell Transplantation
To establish a defined range of injury and transplantation sites, so as to allow the careful and precise histological detection of regenerated organic components in situ, we developed a traumatic muscle damage model with large deficits in muscle–blood vessel–nerve units of the TA muscles (Figure 1, A and B). This model has another advantage in stem cell transplantation research, in that the recovery of contractile function of transplanted muscle can be evaluated on the basis of a functional measurement by use of electrical stimulation via the sciatic nerve. Schematic drawings of the preparation of severely damaged TA muscle and cell transplantation are presented in Figure 1A. Surgery and cell transplantation were performed under halothane anesthesia (Fluothane, Takeda Chemical). The left TA muscle of a wild-type C57BL/6 mouse was exposed by skin incision, and its fascia was minimally cut. With forceps, muscle fibers with nerve and blood vessels were then manually torn off the region surrounding the motor point of the TA muscle. The volume removed was weighed, ensuring that the volume removed was similar for each mouse. The right TA muscle was preserved as a control. Sorted and/or cultured Sk-34 cells from GFP-Tg mice were suspended in 2 μL of DMEM. Then, with a micropipette, the cells were injected slowly into the damaged muscle portion, which was then sutured, and a transparent sterile and analgesic plastic dressing (Nobecutan spray; Yoshitomi Chemical) was sprayed over the wound.
Functional Measurement and Macroscopic Observation of Transplanted Muscle In Situ and In Vitro
Four weeks after transplantation, the cell implantation region was observed by fluorescence microscopy, and the tetanic tension outputs of the left and right TA muscles were measured and compared. The in situ tension output of the TA muscle was determined under sodium pentobarbital anesthesia (40 mg/kg IP). Body (rectal) temperature was maintained at 36±1°C with a heating pad throughout the measurement. The TA muscles and sciatic nerves of both sides were carefully exposed. A bipolar silver electrode (interelectrode distance, 2 mm) was placed under the sciatic nerve, and a stainless steel hook was attached to the distal tendon of each TA muscle with a silk ligature. The animal was transferred to a custom-made operating table that allowed stabilization of the head and limbs in a supine position with surgical tape. A stainless steel hook was attached to a force-distance transducer (TB-611T, Nihon Kohden) connected to a carrier amplifier (AP-621G, Nihon Kohden). Care was taken to avoid interference with the normal blood supply of the TA muscle. Isometric twitches were elicited by use of single pulses (0.5 Hz), and electrical stimulation voltage via the sciatic nerve was set at above the threshold for maximum response (1.5 to 3.0 V). Tetanic tension outputs at optimum length (100 to 120 Hz) were recorded. An outline of the functional measurement of TA muscle is presented in Figure 1B. The muscles were then removed, weighed, soaked in PBS, and pinned on a silicone-coated Petri dish at a predetermined in situ length. Further macroscopic analysis in vitro was then performed under a fluorescence dissection microscope (Olympus SZX12).
Immunostaining and Immunoelectron Microscopy
For immunostaining and immunoelectron microscopic analysis, the muscles were fixed with 4% paraformaldehyde/0.1 mol/L phosphate buffer (4% PFA/PB) overnight. Next, the muscles were washed with 0.01 mol/L PBS and quick-frozen in isopentane; several 7-μm cross sections were then obtained. Rat anti-mouse CD31 monoclonal antibody (PharMingen) was used for detection of blood vessels. The localization of the nerve fibers (axons) was detected by use of rabbit anti–microtubule-associated protein 2 (MAP-2) polyclonal antibody (Chemicon). Schwann cells were detected by use of rabbit anti–glial fibrillary acidic protein (GFAP) polyclonal antibody (NeoMarker; Westinghouse). This antibody also weakly identifies nerve axons. Reactions were visualized by use of Alexa Fluor-594 conjugated goat anti-rabbit and anti-rat antibodies (Molecular Probes). Reaction products were observed and detected by use of fluorescent light-microscopy through 2 single-band filters (Olympus U-MGFPHQ, U-MF-2) and 1 triple-band filter (Chroma U-N61000v2). The validity of the GFP signals was also confirmed by immunostaining with anti-GFP monoclonal antibody (JFP-K2, produced by S.C. Fujita and colleagues at the Mitsubishi Institute of Life Sciences, Tokyo, Japan) and spectral analysis by use of a confocal laser scanning microscope (Olympus FV1000). For immunoelectron microscopy, sections were stained with rat anti-GFP monoclonal antibody, and reactions were visualized with DAB after fixation in 1% glutaraldehyde/0.1 mol/L phosphate buffer. Sections were then fixed in 1% osmium tetroxide/0.05 mol/L phosphate buffer and prepared for electron microscopic analysis.
Transplantation to Kidney Capsule
To confirm their intrinsic differentiation potential, Sk-34 cells were transplanted into nonmuscle tissues. Freshly isolated Sk-34 cells from wild-type mice were transplanted between the kidney capsule and the renal cortex by use of a fine-tip glass pipette. Four weeks after transplantation, the kidneys were removed and fixed with 4% PFA/PB overnight. Samples were fixed in 1% glutaraldehyde/0.1 mol/L PB, followed by fixation in 1% osmium tetroxide/0.05 mol/L PB, and then prepared for electron microscopic analysis.
To further confirm the intrinsic plasticity of Sk-34 cells, we performed FISH analysis (based on the protocol of Star-FISH, Cambio). Sk-34 cells were obtained from male GFP mice and transplanted into a female TA injury model. At 4 weeks after transplantation, FISH was performed with the X chromosome (from RPCI-23 202H24 BAC clone and labeled with digoxygenin by nick translation) and the Y chromosome (biotin-conjugated painting probe, Cambio). Hybridization was visualized by use of streptavidin-Texas Red (Y chromosome) and anti–digoxygenin-Cy3 (X chromosome).
Bone Marrow Transplantation
Whole bone marrow cells from a GFP mouse (1×106 IV) were transplanted into 6 lethally irradiated wild-type mice. After 2 to 6 months, blood samples were obtained and analyzed for GFP chimerism. Sk-34 and Sk-DN cell populations were purified from their muscles by use of CD34 and CD45 (see “cell purification”), and the presence of GFP-positive cells in 4 fractions was examined by FACS analysis.
All data are expressed as mean±SEM. ANOVA was used to determine overall differences, and Tukey’s post hoc analysis was used to identify individual group differences (Table). Differences were considered statistically significant at a value of P<0.05.
Validity of Severe Muscle Damage Model
To confirm the functional reduction in the severe-muscle-damage model, TA muscles were electrically stimulated via the sciatic nerve, and the in situ tetanic tension outputs were measured before and after removal of muscle tissues (muscle–blood vessel–nerve units) in a subgroup of 4 mice. Tetanic tension output was substantially diminished after surgical manipulation, with only 3.8±1.4% of the tension output before surgery being recorded, thus demonstrating that this model was suitable for the purposes of this study (see Figure 1, A and B).
Mass and Functional Recovery
To determine the ideal cell population for regenerative tissue plasticity and for therapeutic application in skeletal muscle injury, we performed a series of Sk-34 cell transplantations. The first population that we investigated was composed of freshly sorted Sk-34 cells (Sk-34-0). The second and third populations were composed of Sk-34 cells cultured for 1 and 5 days (Sk-34-1 and Sk-34-5), respectively. The 4 experimental conditions and data regarding mass and functional recovery are summarized in the Table. The initial and final body weights of experimental animals and the mean mass of the removed muscle were similar in all groups. Recovery was evaluated by comparing the ratios of the surgically treated with the contralateral side (right TA muscle). All 3 cell-transplantation groups showed significant mass recovery compared with the control (non–cell transplanted) group, but greater mass recovery (65±3%) was observed in the Sk-34-0 group. Significant functional recovery (54±4%) was observed only in the Sk-34-0 group, and not in the Sk-34-1 (22±4%) and Sk-34-5 (38±5%) groups. The non–cell-transplanted control group showed a recovery of approximately 20% in both mass and function. Typical raw data for the functional measurements are available in Data Supplement I (http://circ.ahajournals.org/cgi/content/full/CIRCULATIONAHA.105.554832/DC1).
Macroscopic Observation of Tissue Regeneration After Cell Transplantation
To determine the plasticity of each population for tissue regeneration, we performed a morphological analysis of the transplanted muscles. In vivo, under a fluorescence dissection microscope, the transplanted TA muscle of the Sk-34-0 group exhibited numerous green muscle fibers throughout its entire length (Figure 2, A and B). When the transplanted muscles were carefully detached and observed under the fluorescence dissection microscope, blood vessels having green walls were observed on the muscle surface (Figure 2C, arrows). These were confirmed as such on the basis of the blood that remained in the vessels (Figure 2D, arrowheads). Surprisingly, peripheral nerve–like structures with a strong green color were also detected in the Sk-34-0–transplanted muscle (Figure 2E). The site of the Sk-34-5–transplanted muscle (Figure 2, F and G) revealed relatively weak green fluorescence in a limited portion of the muscle and a few specific tissue formations, such as muscle fibers, blood vessels, and nerve-like shapes, that were observed in the Sk-34-0–transplanted muscle (Figure 2, A–E). Similar results were observed in the muscles of the Sk-34-1 group. Successful implantation was evaluated on the basis of whether the tissue formations could be seen macroscopically (under the dissection microscope).
Detection of Donor Cell–Derived Muscle Fibers, Blood Vessels, and Nerve Fibers in Histological Sections After Cell Transplantation
On histology, numerous GFP+ muscle fibers (Figure 3A) and blood vessels (Figure 3, C and D, arrows) were observed in the Sk-34-0–transplanted muscles. The formation of muscle fibers and blood vessels was confirmed by immunostaining for skeletal actin (Figure 3B) and CD31 (Figure 3, E–H). All GFP+ muscle fibers in Figure 3A were skeletal actin+ (Fig. 3B), and GFP+ conduit blood vessels (Figure 3, E and F) and capillaries (arrows in Figure 3, G and H) were both positive for CD31.
In addition, nerve fibers (axons) were immunohistochemically identified by use of MAP-2. The stem (I) and branching (II) portions of dendritic anatomy indicated in Figure 3I were clearly observed as organic structures positive for GFP (Figure 3J; cut as indicated by the white line in Figure 3I). The centers of the individual GFP+ circles were MAP-2+ (red reactions, Figure 3, J and K), indicating the presence of axons. These histological results clearly demonstrate that the dendritic anatomy is that of a bundle of nerve fibers and that transplanted green cells formed the myelin sheath. Independent nerve fibers with GFP+ myelin and MAP-2+ axons were also formed around the nerve bundles (Figure 3K, arrowheads). Both GFAP+ and GFP+ cells (probably Schwann cells, Figure 3L, arrow, yellow reactions) were detected near the GFP+ nerve fibers (Figure 3L, arrowheads; weak red reactions were axons), thus confirming formation of the myelin sheath by the transplanted cells. Similarly, GFAP+ cells were also detected adjacent to the muscle fibers, where Schwann cells are normally found in motor nerve terminals (Figure 3N, arrows). These cells were all positive for GFP (Figure 3, M and O, arrows).
This incorporation of transplanted cells into nerves, skeletal muscle, and blood vessels was observed in the Sk-34-0 group but not in the Sk-34-1 and Sk-34-5 groups. The histology of Sk-34-5–transplanted muscle exhibited fibrous and/or fatty tissue–rich reconstitution with several vascular formations, but few muscle fibers and no nerve tissue reconstitution were seen (Figure 3, P–S). Similar and relatively marked results were noted in the Sk-34-1 group, representing nonsignificant functional recovery (see Table). Detailed detection of GFP signals in the transplanted muscles was also confirmed by immunostaining with anti-GFP antibody and by spectral analysis with a confocal laser scanning microscope. These data are available in Data Supplement II (http://circ.ahajournals.org/ cgi/content/full/CIRCULATIONAHA.105.554832/DC1).
Immunoelectron Microscopic Detection of Donor Cell–Derived Vascular and Neural Cells
Donor cell–derived vascular cells, such as pericytes, vascular smooth muscle cells, and endothelial cells, were evident on immunoelectron microscopic examination of blood vessels that were positive for anti-GFP (dark dots in Figure 4A). Figure 4B shows higher magnification of the region outlined in A. Reaction products of DAB (dark dots) were clearly evident in the nuclei of pericytes (PC) and endothelial cells (E) (white arrows in B). Fibroblast-like cells (FB) were also evident adjacent to blood vessels (Figure 4A). Schwann cells around myelinated (MN) and nonmyelinated (N-MN) nerve fibers (Figure 4C) were GFP+ (arrows). The muscle fiber on the lower side in C is GFP+, and that on the upper side is GFP−. Figure 4D shows the motor end plate. The muscle fibers (MF) and Schwann cells (Sw) depicted in D were strongly positive for anti-GFP (dark dots). Dark dots were also evident in the myo-nuclei (MNu).
Transplantation of Sk-34 Cells Into the Kidney Capsule
To confirm the original differentiation potential of Sk-34-0 cells not associated with the cell fusion mechanism, cells were transplanted between the kidney capsule and renal cortex (nonmuscle tissue). Several tissue structures, including muscle fibers, were evident between the kidney capsule and renal cortex at 4 weeks after transplantation (Figure 5A). In addition, on electron microscopy, myelinated nerve fibers with Schwann cells (Figure 5B) and capillaries (Figure 5C) were also evident around the muscle fibers. These results clearly demonstrate that Sk-34 cells can give rise to nerve–blood vessels–muscle, even in nonmuscle tissue microenvironments, according to their original and/or intrinsic differentiation potential. These data also indicate that the independent tissue plasticity is a result of stem cell differentiation rather than cell fusion.
Detection of Donor-Derived Muscle Fibers by FISH
Four weeks after transplantation, female host TA muscles were reconstituted by freshly isolated male donor Sk-34 cells, exhibiting mass recovery (average, 58±4%), as shown in the Table. FISH analysis was performed both in cross sections (Figure 6B) and in longitudinal sections (Figure 6A). Y chromosome–positive donor-derived nuclei were detected in the GFP-positive fibers (arrows in A and B), and X chromosome was observed in GFP-negative fibers (arrowheads in A and B).
Isolation of Sk-34 and Sk-DN Cells After Bone Marrow Transplantation
To examine whether the Sk-34 and Sk-DN cell populations were derived from bone marrow, whole bone marrow cells from a GFP mouse (1×106 cells) were transplanted into 6 lethally irradiated wild-type mice. At 2 to 6 months after bone marrow transplantation (BMT), peripheral blood exhibited 92±6% GFP chimerism. The sorting pattern of the BMT mouse muscle using CD34 and CD45 was similar to that of the normal wild-type mice, and normal Sk-34 and Sk-DN cell fractions were observed (Figure 7A). However, the number of GFP-positive cells was <5% (Figure 7B), whereas the majority of GFP+ cells were CD45+ (Figure 7C). There were no GFP+ cells in the Sk-34 and Sk-DN (CD34−/45−) cell fractions, thus indicating that the Sk-34 and Sk-DN cells are not derived from bone marrow cells.
After transplantation, significant recovery of both muscle mass and function was observed in the Sk-34-0 group (Table). These results clearly indicate that freshly isolated Sk-34 cells are suitable for transplantation. Primary Sk-34 cells were found as putative myo-endothelial progenitor cells in the muscle.9 The present results demonstrate the potential of Sk-34 cells to form muscle–blood vessel–nerve units, thus leading to significant mass and functional recovery in transplanted muscles. Compared with the non–cell-transplantation group, which showed an approximately 20% recovery in both mass and function, the reconstitution of severely damaged muscle (65±3% recovery in mass and 54±4% in function in the Sk-34-0 group) observed within 4 weeks of transplantation is considered to be clinically successful. Therefore, it is possible that freshly isolated Sk-34 cells could be useful for treating large deficits and/or disruption of the nerve–blood vessel–muscle unit caused by traffic or sporting accidents or after the removal of large tumors.
Successful clinical recovery in the Sk-34-0 group was supported by the presence of large numbers of donor cell–derived (GFP+) muscle fibers, blood vessels, and nerve fibers in the transplanted muscles (Figure 3, A–O). Muscle fibers, blood vessels, and nerve fibers are the main components of skeletal muscle, and the observed reconstruction of these components was reflected in the better mass and functional recovery (Table). However, in the Sk-34-5 group, which was one of the cultured Sk-34 cell–transplanted groups, several blood vessels and a large number of fat and fibrous tissues, but few myofibers and no nerve fibers, were observed (Figure 3, P–S). This trend was more prominent in the Sk-34-1 group (data not shown). In addition, these groups exhibited a limited degree of functional recovery (38% and 22%), although significant mass recovery was observed compared with the controls (Table).
These results suggest that, for Sk-34 cells, culture is not suitable to maintain immaturity or to preserve the ability to commit into myogenic or neural lineages. However, culture had little effect on the ability to commit to vascular lineage, because blood vessel formation was seen in the Sk-34-1 and the Sk-34-5 groups. This would suggest that Sk-34 cells are vasculogenic.
With regard to the formation of blood vessels, we also found that Sk-34-0 cells were able to differentiate into not only endothelial cells but also pericytes and smooth muscle cells and that these cells cooperated to form blood vessels (Figure 4A). These results indicate that SK-34-0 cells have the potential to form both small capillaries and large blood vessels, as confirmed by CD31 immunostaining (Figure 3, E–H). Of note, freshly isolated Sk-34 cells were negative for endothelial linage markers such as CD31, FLK-1, and CD144 (VE-cadherin) but were positive for Sca-1 and CD34.9 Taken together, our results show that SK-34-0 cells are able to reconstitute muscle fibers and blood vessels, as well as peripheral nerves, in severely damaged skeletal muscle, thus facilitating significant mass and functional recovery after transplantation.
In the successful SK-34-0 transplantation procedures, nerve system reconstitution after differentiation into Schwann cells was clearly observed, which would imply that neural progenitor cells are present in skeletal muscle and may be useful for nerve reconstitution therapy. A large number of GFP+ myelin-like structures around MAP-2+ axons were evident on immunofluorescent microscopy (Figure 3, J and K), suggesting that transplanted donor cells differentiate into myelin-producing Schwann cells (GFAP+ cells, Figure 3, L–N) and then form the myelin sheath. This was strongly supported by the observation of donor-derived Schwann cells around myelinated and nonmyelinated nerve fibers and the fact that the motor nerve terminal was clearly evident on immunoelectron microscopy (Figure 4, B and C). Therefore, it appears that SK-34-0 cells actively differentiate into Schwann cells, and thus, that skeletal muscle is also a potential source of Schwann progenitor cells.
In the present study, we clearly demonstrated functional improvement after stem cell therapy, and this is the first report to confirm the functional recovery of transplanted skeletal muscle. In this TA-injury model, tetanic tension output induced by electrical stimulation via the sciatic nerve disappeared immediately after removal of peripheral nerves, muscle fibers, and blood vessels. Four weeks after transplantation, tension output recovered by approximately 55% compared with nondamaged muscles in the SK-34-0 group, whereas the non–cell-transplanted control group showed only a 20% recovery. Furthermore, the SK-34-1 and SK-34-5 groups showed significant mass recovery (approximately 45%) but little functional recovery (<38%, Table) after formation of blood vessels and a large amount of fat and fibrous tissue but few myofibers and no nerve fibers (Figure 3, P–S). These results indicate that nerve reconstitution is important for the functional recovery of skeletal muscle and may contribute to almost 50% of the observed total “functional” recovery (55−20=35 versus 55−38=17, and 17/35×100=49%). This further suggests that innervation is essential for functional recovery of severely damaged muscle. Similarly, the formation of blood vessels may contribute to 55% of the observed total “mass” recovery (65−20=45 versus 45−20=25, and 25/45×100=55%), which suggests that new muscle fiber formation originating from transplanted Sk-34 cells may contribute to 45% of the total mass recovery.
In addition to cell plasticity and/or transdifferentiation, it was recently reported that spontaneous cell fusion might be a mechanism by which adult stem cells, particularly bone marrow cells, exhibit various unexpected phenotypes.17,18 There is a possibility that this mechanism contributed to the reconstitution observed in the present study. We therefore confirmed the original and/or intrinsic differentiation potential of SK-34-0 cells by transplantation into nonmuscle tissues. Transplantation of freshly isolated Sk-34 cells between the kidney capsule and renal cortex showed that differentiation into nerve, muscle, and vascular cells occurs even in nonmuscle tissues. This indicates that SK-34-0 cells have an intrinsic potential to differentiate into nerve-muscle-vascular cells that is independent of cell fusion. However, cell fusion is one of the possible mechanisms for muscle fiber regeneration and may have had a role in this study, because programmed cell fusion is the normal physiological mechanism by which myoblasts form multinucleated muscle fibers, although donor cell–derived muscle fibers specifically contributed to total recovery in the present TA injury model, as shown by FISH analysis (Figure 6, A and B). In addition, our BMT experiment clearly demonstrated that Sk-34 cells are not derived from bone marrow, which is consistent with previous reports showing that myogenic stem cells were not of bone marrow origin, and that myogenic engraftment levels after BMT were extremely low and caused primarily by cell fusion.19,20 We therefore consider that the significant reconstitution of muscle–blood vessel–nerve units observed after transplantation was not a result of cell fusion but primarily of the differentiation potential of Sk-34 cells.
The results of our BMT experiment also suggested the existence of tissue-specific endothelial progenitor cells (EPCs) and/or vasculogenic cells. The present results clearly indicate that muscle tissue–specific vasculogenic cells were present and that they were not bone marrow–derived, whereas circulating EPCs were of bone marrow origin.21,22 Recent studies have suggested that the contribution of EPCs to neovessel formation may range from 5% to 25% in response to granulation tissue formation23 or growth factor–induced neovascularization.24 Thus, cooperation between EPCs and “tissue-specific vasculogenic cells” in the process of neovascularization should be studied in the future.
In conclusion, our fractionated, skeletal muscle–derived Sk-34 cells showed the potential for therapeutic use in severely damaged muscle that has substantially diminished function with large deficits in nerve fibers, muscle fibers, and blood vessels. These cells were clearly able to produce a recovery of both muscle mass and function. Our data suggest that there is an extremely low probability that Sk-34 cells are derived from bone marrow.
This work was supported by a Grant-in-Aid for Scientific Research from the Ministry of Education, Science, and Culture of Japan and Tokai University Research aid. We thank Masafumi Koyama, Olympus Co Ltd, for his technical support by performing confocal laser microscopic analysis.
The online-only Data Supplement can be found at http://circ.ahajournals.org/cgi/content/full/CIRCULATIONAHA.105.554832/DC1.
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