TRANSLATIONAL AND CLINICAL RESEARCH Acceleration of Skeletal Muscle Regeneration in a Rat Skeletal Muscle Injury Model by Local Injection of Human Peripheral Blood-Derived CD133-Positive Cells MING SHI,a MASAKAZU ISHIKAWA,a NAOSUKE KAMEI,a,b TOMOYUKI NAKASA,a NOBUO ADACHI,a MASATAKA DEIE,a TAKAYUKI ASAHARA,c MITSUO OCHIa a
Department of Orthopedic Surgery, Graduate School of Biomedical Sciences, Hiroshima University, Hiroshima, Japan; bKobe Institute of Biomedical Research and Innovation/RIKEN Center for Developmental Biology, Kobe, Japan; cDepartment of Regenerative Medicine Science, Tokai University School of Medicine, Kanagawa, Japan Key Words. Peripheral blood • CD1331 cells • Muscle regeneration • Endothelial progenitor cell • Vasculogenesis
ABSTRACT Muscle injuries in sport activities can pose challenging problems in traumatology and sports medicine. The best treatment for muscle injury has not been clearly established except for the conservative treatment that is routinely performed. We investigated the potential of human adult CD1331 cells to contribute to skeletal muscle regeneration in an athymic rat model. We tested whether CD1331 cells locally transplanted to the skeletal muscle lacerated models could (a) induce vasculogenesis/angiogenesis, (b) differentiate into endothelial and myogenic lineages, and (c) ﬁnally promote histological and functional skeletal myogenesis. Granulocyte colony stimulating factor-mobilized peripheral blood (PB) CD1331 cells, PB mononuclear cells, or phosphate-buffered saline was locally injected after creating a muscle laceration in the tibialis anterior muscle in athymic rats. After treatment, histological and functional skeletal myogenesis was
observed signiﬁcantly in the CD1331 group. The injected CD1331 cells differentiated into endothelial and myogenic lineages. Using real-time polymerase chain reaction analysis, we found that the gene expressions related to microenvironment conduction for host angiogenesis, ﬁbrosis, and myogenesis were ideally up/downregulated. Our results show that CD1331 cells have the potential to enhance the histological and functional recovery from skeletal muscle injury rather via indirect contribution to environment conduction for muscular regeneration. It would be relatively easy to purify this cell fraction from PB, which could be a feasible and attractive autologous candidate for skeletal muscle injuries in a clinical setting. These advantages could accelerate the progression of cell-based therapies for skeletal muscle injuries from laboratory to clinical implementation. STEM CELLS 2009;27:949–960
Disclosure of potential conﬂicts of interest is found at the end of this article.
INTRODUCTION Muscle injuries in sport activities can pose challenging problems in traumatology and sports medicine. The best treatment for muscle injury has not yet been clearly deﬁned, and the recommended treatment regimens are mostly conservative and routine. To establish intensive treatments, many investigators have focused on antiﬁbrosis after muscle injury using drugs or cytokine inhibitors with antiﬁbrotic properties to obtain muscle recovery [1, 2].
Recently, an attractive agent for regenerative medicine, endothelial progenitor cells (EPCs), has emerged. Vasculogenesis by EPCs, which is involved in the development of the blood vessel system during embryogenesis , was not identiﬁed as a mechanism of postnatal endothelial regeneration until the discovery of bone marrow-derived and circulating EPCs in adults [4–6]. After this discovery, there were many reports that investigated the potential of EPCs for therapeutic neovascularization and demonstrated their efﬁcacy for limb ischemia, stroke, diabetic wounds, fracture healing, ligament injury, and axon growth in the in vitro and in vivo models
Author contributions: M.S.: collection and/or assembly of data, data analysis and interpretation, manuscript writing; M.I.: conception and design, provision of study material or patients, collection and/or assembly of data, data analysis and interpretation, manuscript writing, equal contribution to ﬁrst author; N.K.: collection and/or assembly of data, data analysis and interpretation; T.N.: collection and/or assembly of data, data analysis and interpretation; N.A.: collection and/or assembly of data, data analysis and interpretation; M.D.: collection and/or assembly of data, data analysis and interpretation; T.A.: collection and/or assembly of data, data analysis and interpretation; M.O.: ﬁnancial support, data analysis and interpretation, ﬁnal approval of manuscript. Correspondence: Masakazu Ishikawa, M.D., Ph.D., Department of Orthopedic Surgery, Graduate School of Biomedical Sciences, Hiroshima University, 1-2-3 Kasumi, Minami-ku, Hiroshima 734-8551, Japan. Telephone: þ81-82-257-5233; Fax: þ81-82-257-5234; e-mail: [email protected]
Received July 7, 2008; accepted for publication December 17, 2008; ﬁrst published online in STEM C AlphaMed Press 1066-5099/2009/$30.00/0 doi: 10.1002/stem.4 CELLS EXPRESS January 15, 2009. V
STEM CELLS 2009;27:949–960 www.StemCells.com
CD133þ Cells Accelerate Muscle Regeneration
[7–13]. In particular, EPC transplantation for cardiac repair has recently emerged as a promising new approach for cardiomyogenesis through vasculogenesis . Inspired by these data, we focused on EPCs as a candidate for skeletal myogenesis. We used peripheral blood (PB)-derived CD133þ cells, which are a subpopulation of CD34þ cells, a well-characterized hematopoietic stem cell/EPC fraction [15, 16]. The CD133 epitope is a 120-kDa glycosylated polypeptide expressed by a population of circulating human hematopoietic/endothelial progenitors [17–19]. This cell fraction is also recognized as highly proliferating and an early committed stem/progenitor cells, compared with CD34þ cells . A recent article reviewed that CD133 is the most promising stem cell marker and that CD133þ cells could be isolated from human PB, bone marrow, cord blood, fetal liver, adult kidney, neonatal foreskin, and pancreas. The isolated CD133þ cell fraction was capable of both self-renewal and multilineage differentiation in vitro and in vivo and could contribute to many tissue regenerations, thus representing a promising treatment in clinical situations . We demonstrated in a recent report that circulating CD133þ cells accelerated axon growth in a rat organ culture system via upregulation of vascular endothelial growth factor (VEGF) . It was also revealed that CD133þ cells have the potential to conduct the host’s environment to regeneration via modiﬁcation with vasculogenic/angiogenic factor. Accordingly, we hypothesized whether locally transplanted CD133þ cells in animal skeletal muscle lacerated models could (a) induce vasculogenesis/angiogenesis, (b) differentiate into endothelial and skeletal myogenic lineages, and (c) ﬁnally promote functional skeletal myogenesis. Here, we demonstrate an acceleration of skeletal muscle regeneration induced by the transplantation of human PB-derived CD133þ cells into a rat skeletal muscle injury model. We speculate that transplantation of circulating CD133þ cells could represent a future treatment for skeletal muscle injury.
The Institutional Animal Care and Use Committees of the University of Hiroshima approved all animal procedures, including human cell transplantation.
Preparation and Characterization of Cells for Transplantation Human PB-derived mononuclear cells (MNCs) were isolated by density gradient centrifugation from healthy male volunteers (2836 years, n ¼ 3). Granulocyte colony stimulating factor (G-CSF)mobilized (GM) human PB-derived CD133þ cells were purchased from Cambrex (Walkersville, MD, http://www.cambrex.com; lot nos. 030865-5, 061069A, 061308A). For transplantation, frozen cells were thawed according to the manufacturer’s protocol and resuspended aseptically in phosphate-buffered saline (PBS). To conﬁrm the purity and characterize MNCs and CD133þ cells, ﬂuorescence-activated cell sorting (FACS) analysis (Becton, Dickinson and Company, Franklin Lakes, NJ, http://www.bd.com) was performed with a FACS Calibur Analyzer and CellQuest Pro Software (Becton Dickinson Immunocytometry Systems, Mountain View, CA, http://www.bd.com). Dead cells were excluded from the plots on the basis of propidium iodide (PI) staining (Sigma-Aldrich, St. Louis, MO, http://www.sigmaaldrich.com). Cells were stained with monoclonal antibodies and analyzed. The monoclonal antibodies CD133-APC (clone 293C3; Miltenyi Biotec, Bergisch Gladbach, Germany, http://www.miltenyibiotec.com), CD34-PE (clone 581; BD Pharmingen, San Diego, CA,
http://www.bdbiosciences.com/index_us.shtml), CD45-FITC (clone HI30; BD Pharmingen), IgG1-PE isotype control (BD Pharmingen), IgG1-FITC (BD Pharmingen), and IgG2b-APC (BD Pharmingen) and PI were used.
Experimental Animals Female F344/NJcl-rnu/rnu athymic nude rats (9 weeks old, n ¼ 40) were used in this study. All rats were obtained from CLEA Japan Inc. (Meguro-ku, Tokyo, Japan, http://www.clea-japan. com). Rats were fed a standard maintenance diet and provided with water continuously.
Rat Model of Skeletal Muscle Injury and Cell Transplantation The rat muscle injury model was based on a previous report from our department . Rats were anesthetized with ketamine and xylazine (60 and 10 mg/kg), and an adequate depth of anesthesia was maintained such that the rats were unresponsive to tactile stimulation. The tibialis anterior muscles were used in all rats. An anterolateral skin incision was made in the left leg, and the tibialis anterior muscle was exposed. The fascia was incised longitudinally and carefully released from the muscle belly. The muscle belly was then lacerated transversely at the midportion, using a scalpel. The defect was wedge shaped, and its size was approximately 6 mm long, 4 mm wide, and 5 mm deep. CD133þ cells were resuspended in 30 ll PBS and transplanted into the muscle defect after closure of the fascia with a suture (transplanted number of cells: 1 105 cells per animal). The same number of MNCs, or the same volume of PBS without cells, was transplanted as controls (each group, n ¼ 13). The skin was closed and no immobilizers were applied. All rats could move freely in their cages.
Macroscopic Assessment of Lacerated Muscle All animals were evaluated macroscopically during healing and regeneration at 1 and 4 weeks after injury under anesthesia with intraperitoneal ketamine (60 mg/kg) injection. The bridging tissue occupying the lacerated sites was observed carefully and captured using a digital camera. In this experiment, no obvious infection was observed in any of the operated rats.
Electromechanical Evaluation To evaluate the functional regeneration of the tibialis anterior muscles after cell transplantation, isometric tensile strength produced by stimulating the common peroneal nerve was measured with transducer load cells (LVS-1KA; Kyowa Electronic Instruments, Tokyo, Japan, http://www.kyowa-ei.co.jp) and recorded with a sensor interface (PCD-300A; Kyowa Electronic Instruments) and software (PCD-30A; Kyowa Electronic Instruments) as described previously . All rats were set in the supine position under general anesthesia with intraperitoneal ketamine (60 mg/kg) injection. An anterolateral skin incision was applied to both legs and the tibialis anterior muscles and common peroneal nerves were exposed. Both legs were ﬁxed tightly to the leg holder, which was made especially for this study. The common peroneal nerves of both legs were stimulated with an electrostimulator (SEN2201; Nihon Koden, Tokyo, Japan, http://www.nihonkohden. co.jp). Frequencies of stimulation were 1 Hz (fast twitch) and 50 Hz (tetanus). To determine the voltage of stimulation, the minimum voltage that visibly contracted the tibialis anterior muscle was measured as a threshold. The nerve was then stimulated by a voltage that was 10-fold higher than that of threshold and the maximum isometric tensile strength produced by the tibialis anterior muscles of both legs was measured. To minimize interanimal variation, the data were normalized to nontreated controls, that is, the strength of the experimental muscle was divided by that of the control noninjured muscle which was measured from the
Shi, Ishikawa, Kamei et al.
contralateral side and showed as a strength ratio. Strength ratios of the lacerated side to the contralateral side were also calculated in fast twitch and tetanus stimulations.
Processing of the Regenerated Muscle Tissue After electromechanical evaluation, the wet weight of the tibialis anterior muscle of both legs was measured and the specimens were analyzed histologically (n ¼ 5 for each time point and group). All rats were euthanized with an overdose of sodium pentobarbital at the end of electromechanical studies and regenerated tissues were harvested and placed in ﬁxative solutions for morphometric evaluations or were quickly embedded in tissue-freezing medium (Triangle Biomedical Sciences, Durham, NC, http:// www.trianglebiomedical.com), snap-frozen in liquid nitrogen, and stored at –80 C for histological, histochemical, and immunoﬂuorescent staining. Pieces of regenerated tissue from each group were also used for RNA isolation.
Morphometric Evaluation of Fibrosis, Myoﬁber, Muscle Regeneration, and Capillary Density For evaluation of the ﬁbrotic area, the myoﬁber diameter, and the number of centronucleated myoﬁber after muscle laceration, Masson trichrome staining was carried out on frozen sections of each sample. The average area of ﬁbrosis in saggital sections and the average diameter of myoﬁbers in axial sections were digitized and images were imported into a computer using Photoshop software (Adobe Systems Inc., San Jose, CA, http://www.adobe.com). Computer analysis of this digitized information was performed using the image analysis software ImageJ (National Institutes of Health, Bethesda, MD, http://www.nih.gov). At 200 magniﬁcation, ﬁve randomly selected ﬁelds per muscle were evaluated for myoﬁber diameter in axial sections. For evaluation of muscle regeneration, the total number of centronucleated myoﬁbers was measured within the injured site in ﬁve random ﬁelds of each sample using a previously described protocol [23, 24]. Histochemical staining with isolectin B4, a marker for rat endothelial cell (Vector Laboratories, Burlingame, CA, http://www.vectorlabs.com), was performed and the capillary density was morphometrically evaluated by histological examination of ﬁve randomly selected ﬁelds around the ﬁbrosis area in axial sections at 200 magniﬁcation. Capillaries were recognized as tubular structures positive for isolectin B4. Three observers, blinded to the experimental groups, performed all of the measurements.
Reverse Transcription-Polymerase Chain Reaction Analysis of RNA Isolated from Transplanted Cells Total RNA was obtained from PB MNCs, CD133þ cells, and regenerated tissues at day 3 (n ¼ 3 in each group) using TRIZOL (Invitrogen, Japan, http://www.invitrogen.co.jp) according to the manufacturer’s procedure. First-strand cDNA was synthesized by using the RNA LA Polymerase Chain Reaction (PCR) Kit, Version 3.0 (Takara, Otsu, Japan, http://www.takara-bio.co.jp), and ampliﬁed by Taq DNA polymerase for reverse transcription (RT)-PCR analysis (AmpliTaq Gold DNA Polymerase; Applied Biosystems, Foster City, CA, http://www.appliedbiosystems.com) using the human-speciﬁc primer sets listed in Table 1. We used cDNA ampliﬁed from human muscle RNAs as the positive control for myogenic and endothelial markers, and the RNAs without reverse transcriptase treatment were used as the negative control in each group. PCR was performed with a thermal cycler (MJ Mini Gradient Thermal Cycler; Bio-Rad, Hercules, CA, http:// www.bio-rad.com) under the following conditions: 35 cycles of 30 seconds of initial denaturation at 94 C, annealing at 56 C for 1 minute, and 30 seconds of extension at 72 C. All procedures were performed according to the manufacturer’s instructions. RTPCR products were electrophoresed in a 1.5% agarose gel containing ethidium bromide in tris-borate-EDTA electrophoresis buffer and visualized by ultraviolet transillumination.
Real-Time PCR Analysis of Intrinsic Factors in the Regenerated Muscles of Rats For real-time RT-PCR, at day 3, the specimens were harvested and analyzed (each group, n ¼ 3). Real-time quantitative RTPCR was performed with a MiniOpticon System (Bio-Rad) using QuantiTect SYBR Green (Qiagen KK, Tokyo, Japan, http:// www.qiagen.co.jp) according to the manufacturer’s protocol, using the rat-speciﬁc primer sets listed in 1. mRNA expression levels were normalized to b-actin. All samples were run in triplicate.
Statistical Analysis All values were expressed as means SEs. Statistical analysis was performed by a one-way analysis of variance and subsequent Fisher’s least signiﬁcant difference test. p values less than .05 were considered as statistically signiﬁcant.
Immunoﬂuorescent Staining Tissues embedded in the freezing medium were serially sectioned at 6 lm, mounted on silane-coated glass slides, and air-dried before being ﬁxed with 4% paraformaldehyde at 4 C for 5 minutes and stained immediately. To evaluate ﬁbrosis and muscle regeneration and to detect transplanted human cells in the regenerated tissues and their differentiation to endothelial and myogenic lineages, immunohistochemistry was performed with the following antibodies: vimentin (Santa Cruz Biotechnology Inc., Santa Cruz, CA, http://www.scbt.com) to detect ﬁbrotic tissue [23–26], desmin and MyoD1 (both from Santa Cruz Biotechnology) to detect rat- and human-derived myotube formation , Pax7 (R&D Systems Inc., Minneapolis, MN, http://www.rndsystems.com) and M-cadherin (Santa Cruz Biotechnology) to detect rat satellite cells in skeletal muscle [28, 29], antibody against human-speciﬁc mitochondria (hMit; Chemicon International, Temecula, CA, http://www.chemicon.com) to detect transplanted human cells, and von Willebrand factor (vWF) protein (Santa Cruz Biotechnology) to detect rat- and human-derived endothelial cells. Secondary antibodies used for each immunostaining were as follows: Alexa Fluor 488- or 568-conjugated goat anti-mouse IgG1 (Molecular Probes Inc., Eugene, OR, http://probes.invitrogen.com) for hMit and Alexa Fluor 488- or 568-conjugated goat anti-rabbit IgG (Molecular Probes) for vimentin, desmin, MyoD1, and vWF protein. 4,6-Diamidino-2-phenylindole solution was applied for 5 minutes for nuclear staining.
RESULTS Characterization of PB-Derived MNCs and GM-PB CD1331 Cells Using FACS analysis, it was demonstrated that MNCs contained only an extremely small number of CD133þ cells (0.04%) and was conﬁrmed that the transplanted CD133þ cell population had a purity of 98.33% (Fig. 1A). RT-PCR analysis revealed that no remarkable gene expression could be observed, such as muscle-related genes or endothelial lineage markers, in MNCs or CD133þ cells (Fig. 1B).
Conﬁrmation of Engraftment of Human MNCs and CD133 Cells The ﬁrst critical point of this experiment was to assess the ability of human MNCs and CD133þ cells to engraft into injured skeletal muscles of athymic rats. At 1 week after injection, we identiﬁed human mitochondria-expressing cells in the skeletal muscles of all rats, although the number of detected cells was higher in the CD133þ group than in the MNC group (Fig. 1C).
CD133þ Cells Accelerate Muscle Regeneration
Table 1. Primers utilized in RT-PCR and real-time PCR Gene
hCD133 hPax7 hMyoD1 hM-cadherin hVEGFR2/KDR hGAPDH
rVEGFA rTGF-b1 rPax7 rMyoD1 rMyogenin rMyf5 rMrf4 rACTB
Human-speciﬁc primer pair sequence for RT-PCR
Fragment length (bp)
F: 50 -CGTGGATGCAGAACTTGACAAC-30 R: 50 -CACACAGTAAGCCCAGGTAGTAAAA-30 F: 50 -TCTCCCGACAGCTGCGTGTCT-30 R: 50 -CTCGTCCAGCCGGTTC-30 F: 50 -GCCGCTCCGTGTTCC-30 R: 50 -GCCTGCGGTATAAACGTACA-30 F: 50 -CGATCAGCGTATCCGAGAA-30 R: 50 -CTCTTAGCCTGAAGCGATCA F: 50 -CAAATGTGAAGCGGTCAACAAAGTC-30 R: 50 -ATGCTTTCCCCAATACTTGTCGTCT-30 F: 50 -CTGATGCCCCCATGTTCGTC-30 R: 50 -CACCCTGTTGCTGTAGCCAAATTCG-30
Rat-speciﬁc primer pair sequence for real-time PCR
F: 50 -TACCTCCACCATGCCAAGTG-30 R: 30 -TCTGCTCCCCTTCTGTCGTG-30 F: 50 -CCACGTGGAAATCAATGGGA-30 R: 50 -GGCCATGAGGAGCAGGAAG-30 F: 50 -GCCCTCAGTGAGTTCGATTAGC-30 R: 50 -TCCTTCCTCATCGTCCTCTTTC-30 F: 50 -GCGACAGCCGATGACTTCTAT-30 R: 50 -GGTCCAGGTCCTCAAAAAAGC-30 F: 50 -GACCCTACAGGTGCCCACAA-30 R: 50 -ACATATCCTCCACCGTGATGCT-30 F: 50 -GGCTGGTCACTGCCTCATGT-30 R: 50 -CTTGCGTCGATCCATGGTAGT-30 F: 50 -GCCCCTTTCCGCCTAATC-30 F-50 -ACTAAGTCTCTTGCCTTTCATAAATTCTG-30 F: 50 -GATCATTGCTCCTCCTGAGCG-30 F: 50 -TGCTGATCCACATCTGCTGGA-30
389 351 213 416 596
Fragment length (bp)
55 91 70 73 70 70 71 92
Abbreviations: bp, base pair; F, forward; hGAPDH, human glyceraldehyde 3 phosphate dehydrogenase; hM-cadherin, human M-cadherin; hVEGFR2, human vascular endothelial growth factor 2; R, reverse; RT-PCR, reverse transcription-polymerase chain reaction; rVEGFA, rat vascular endothelial growth factor A; TGFb1, transforming growth factor beta 1.
Morphological and Functional Regeneration of Skeletal Muscle After CD1331 Cell Injection We initially examined the gross appearance of the muscles and performed histological examinations. Gross observation demonstrated that the lacerated regions were signiﬁcantly occupied with bridging tissue in all rats in the CD133þ group. Conversely, lacerated regions showed signiﬁcant recess in the control groups (Fig. 2A). However, there was no significant difference in the moist weight ratio among the groups (at week 1: PBS group, 0.861 0.028; MNC group, 0.880 0.019; CD133 group, 0.891 0.015, respectively; at week 4: PBS group, 0.794 0.051; MNC group, 0.872 0.019; CD133 group, 0.908 0.018, respectively) (Fig. 2B). We conducted physiological measurements of strength recovery as an amount of functional regeneration on the three groups at 1 and 4 weeks after injury. We compared the fasttwitch and tetanus strength ratios of the CD133þ cell injection group and the control groups. The fast-twitch strength ratio was signiﬁcantly higher in the CD133þ group compared with control groups. However, MNC treatment showed higher improvement of muscle strength compared with the PBS group at both weeks 1 and 4 (at week 1: PBS group, 0.527 0.015; MNC group, 0.614 0.027; CD133 group, 0.729 0.016, respectively [p < .01 for PBS vs. MNC, PBS vs. CD133, and MNC vs. CD133 group]; at week 4: PBS group, 0.629 0.013; MNC group, 0.672 0.057; CD133 group, 0.819 0.005, respectively [p < .01 for PBS vs. CD133 and MNC vs. CD133 group; p < .05 for PBS vs. MNC group]).
The tetanus strength ratio was also signiﬁcantly higher in the CD133þ group compared with control groups (at week 1: PBS group, 0.673 0.084; MNC group, 0.742 0.063; CD133 group, 0.836 0.073, respectively [p < .01 for PBS vs. CD133 and MNC vs. CD133 group; p < .05 for PBS vs. MNC group]; at week 4: PBS group, 0.697 0.036; MNC group, 0.803 0.037; CD133 group, 0.908 0.067, respectively [p < .01 for PBS vs. MNC, PBD vs. CD133, and MNC vs. CD133 group]) (Fig. 2C). Masson trichrome staining for evaluation of ﬁbrosis demonstrated that ﬁbrosis was prevented at week 1 with a small number of inﬂammatory cells in the CD133þ group and there was almost complete healing with a very small area of ﬁbrosis at week 4. In contrast, large defects with abundant inﬂammatory cells were easily observed at week 1 and large ﬁbrosis areas were observed at week 4 in the control groups (at week 1: PBS group, 6,416.143 322.671 lm2; MNC group, 1,537.571 114.473 lm2; CD133 group, 600.429 56.745 lm2, respectively [p < .01 for PBS vs. MNC and PBS vs. CD133 group; p < .05 for MNC vs. CD133]; at week 4: PBS group, 13,669.710 942.549 lm2; MNC group, 5,317.714 446.150 lm2; CD133 group, 481.714 37.503 lm2, respectively [p < .01 for PBS vs. MNC, PBS vs. CD133, and MNC vs. CD133 group]) (Fig. 2D, 2E). The diameter of myoﬁbers near the periﬁbrosis sites of all groups was evaluated. At week 1 after injury, signiﬁcant inﬂammatory cell accumulation was observed in the PBS group and the diameter of myoﬁbers was signiﬁcantly greater in the CD133 group compared with control groups at both 1
Shi, Ishikawa, Kamei et al.
Figure 1. Representative ﬂuorescenceactivated cell sorting analysis of MNCs and granulocyte colony stimulating factor-mobilized peripheral blood (GM-PB) CD133þ cells. (A): Propidium iodide staining excluded dead cells and then the gated MNC population within live cells was assessed. Cells were stained with CD133 and CD34 antibodies. Numbers show the percentage of positive cells in each staining. The purity of GM-PB CD133þ cells was conﬁrmed (98.33%) and only a scarce number of cells were observed in MNCs (0.04%). (B): Reverse transcription-polymerase chain reaction analysis of humanspeciﬁc genes for skeletal myogenic and endothelial lineages in freshly isolated CD133þ cells. There was no expression of skeletal myogenic and endothelial markers in both the MNCs and GM-PB CD133þ cells. (C): Transplanted human cells, MNCs, and GM-PB CD133þ cells were conﬁrmed by staining with hMit. In the interstitial space, hMit-positive cells were observed in the MNC and CD133þ groups at 1 week after transplantation (arrows), whereas the number of human cells was abundant in the CD133þ group and there were no hMit-positive human cells in the PBS group. Scale bar ¼ 100 lm; original magniﬁcation, 400. Abbreviations: APC, allophycocyanin; DAPI, 4,6-diamidino-2phenylindole; FSC, forward scatter; hGAPDH, human glyceraldehyde 3 phosphate dehydrogenase; hM-cadherin, human M-cadherin; hMit, human-speciﬁc mitochondria; hVEGFR2, human vascular endothelial growth factor receptor 2; MNCs, mononuclear cells; PBS, phosphate-buffered saline; PE, phycoerythrin; RT, reverse transcription; SSC, side scatter.
and 4 weeks after cell injection (at week 1: PBS group, 38.070 0.877 lm; MNC group, 42.71 0.686 lm; CD133 group, 48.96 0.589 lm, respectively [p < .01 for PBS vs. MNC, PBS vs. CD133, and MNC vs. CD133 group]; at week 4: PBS group, 79.944 7.159 lm; MNC group, 107.786 1.951 lm; CD133 group, 130.404 2.719 lm, respectively [p < .01 for PBS vs. MNC and PBS vs. CD133 group; p < .05 for MNC vs. CD133 group]) (Fig. 3A, 3B). We observed more centronucleated (regenerating) myoﬁbers in the CD133þ group than in the control groups, although centronucleated myoﬁbers were also observed in the control groups at 1 and 4 weeks after injury (at week 1: PBS group, 15.20 1.35; MNC group, 18.90 1.28; CD133 group, 24.00 1.90, respectively [p < .01 for PBS vs. CD133. p < www.StemCells.com
.05 for MNC vs. CD133 group]; at week 4: PBS group, 13.40 1.07; MNC group, 14.10 0.57; CD133 group, 20.70 1.17, respectively [p < .01 for PBS vs. CD133 group and MNC vs. CD133 group]) (Fig. 3C).
Enhancement of Intrinsic Angiogenesis by Capillary Density Enhanced angiogenesis by recipient cells following cell transplantation was assessed by capillary density at weeks 1 and 4. Vascular staining of isolectin B4 in tissue samples demonstrated enhanced neovascularization around the ﬁbrosis area in the CD133þ group compared with control groups (Fig. 3D, 3E). Neovascularization assessed by capillary density was
CD133þ Cells Accelerate Muscle Regeneration
Figure 2. Macroscopic and functional improvement and prevention of ﬁbrosis in animals after human CD133þ cell injection. Representative macroscopic ﬁndings of the lacerated sites of tibialis anterior muscles (arrows) at weeks 1 and 4. Lacerated sites were ﬁlled with well-formed tissue only in the CD133þ group (A), although there was no significant difference in moist weight ratios among the groups (B). (C): Fast-twitch and tetanus strength ratios were signiﬁcantly higher in the CD133þ group compared with the control groups at weeks 1 and 4. (D): Representative histological evaluation of ﬁbrosis by Masson trichrome staining. Fibrosis (bluestained area) was signiﬁcantly smaller and replaced with normal skeletal muscle tissue in the CD133þ group compared with control groups (original magniﬁcation, 40). (E): Data were calculated as means SEs. *, p < .05; **, p < .01. Abbreviations: MNC, mononuclear cell; PBS, phosphate-buffered saline; W, week.
signiﬁcantly enhanced in the CD133þ group compared with either the PBS or the MNC group (at week 1: PBS group, 35.561 1.847 per square millimeter; MNC group, 45.084 0.990 per square millimeter; CD133 group, 67.845 1.485 per square millimeter, respectively [p < .01 for PBS vs. MNC, PBS vs. CD133, and MNC vs. CD133 group]; at week 4: PBS group, 50.237 2.859 per square millimeter; MNC group, 56.965 3.542 per square millimeter; CD133 group, 76.432 3.376 per square millimeter, respectively [p < .01 for PBS vs. CD133 group; p < .05 for MNC vs. CD133 group]) (Fig. 3D, 3E).
Immunoﬂuorescent Evaluation of Muscle Fibrosis and Regeneration After Cell Injection At week 1, histological staining and immunoﬂuorescence assessment were carried out on all the groups in axial sections
of injured sites. For evaluation of ﬁbrosis, we used vimentin as a correlative indicator of ﬁbrosis scar formation in injured muscle following previous report [22–25]. Injection of CD133þ cells appeared to prevent ﬁbrosis more effectively compared with control groups by immunoﬂuorescence staining, although injection of MNCs tend to show less ﬁbrosis than the PBS group (Fig. 4A). Inhibition of muscle ﬁbrosis among all the groups appeared to be dependent on the injected cells and there was a signiﬁcant difference in ‘‘percentage of ﬁbrosis area’’ among the groups (Fig. 4B). Desmin, a cytoskeletal intermediate ﬁlament, is expressed during skeletal muscle development and myotube formation  and is widely used as a marker for distinguishing individual cell types within a tissue, such as myoblasts from ﬁbroblasts in the regenerating and central zones of muscle injury. Muscles injected with CD133þ cells contained numerous regenerating myoﬁbers, that is, desmin-positive myoﬁbers,
Figure 3. Histological skeletal muscle healing and enhancement of intrinsic angiogenesis in animals receiving CD133þ cells. (A): Representative histological evaluation carried out using Masson trichrome-stained samples. (B, C): In the CD133þ group, the diameter of myoﬁbers and the number of regenerating myoﬁbers, that is, centronucleated myoﬁbers, were signiﬁcantly increased at weeks 1 and 4 compared with the control groups. Abundant inﬂammatory cells were observed at week 1 and a small number of thin diameter myoﬁbers were found at week 4 in the control groups. Scale bars ¼ 100 lm; original magniﬁcation, 200. Data were calculated as means SEs. *, p < .05; **, p < .01. (D): Representative immunostaining for isolectin B4 in each group at weeks 1 and 4. (E): Neovascularization assessed by capillary density was signiﬁcantly enhanced in the CD133þ group compared with control groups. Scale bars ¼ 100 lm; original magniﬁcation, 200. Data were calculated as means SEs. *, p < .05; **, p < .01. Abbreviations: MNC, mononuclear cell; PBS, phosphate-buffered saline; W, week.
CD133þ Cells Accelerate Muscle Regeneration
Figure 4. Immunoﬂuorescent evaluation of muscle ﬁbrosis and regeneration and differentiation of human cells into endothelial and skeletal myogenic lineages. (A, B): Representative immunoﬂuorescent evaluation with vimentin and desmin revealed that the percentage of ﬁbrosis area was smaller and percentage of regenerated area was signiﬁcantly greater after CD133þ cell transplantation. Scale bars ¼ 500 lm; original magniﬁcation, 40. (C): Representative double-immunoﬂuorescence staining for vWF and hMit at day 28 in each group. Human endothelial cells were identiﬁed as double-positive cells for vWF (green) and hMit (red) (arrows). The number of double-positive cells was scarce in the MNC group. There were no double-positive cells in the PBS group. (D): Representative double-immunoﬂuorescence staining for MyoD1 and hMit at day 28 in each group. Human myocytes were identiﬁed as double-positive cells for MyoD1 (red) and hMit (green) (arrows), although there were few double-positive cells in the MNC group. There were no double-positive cells in the PBS group. Scale bars ¼ 100 lm; original magniﬁcation, 400. Abbreviations: DAPI, 4,6-diamidino-2-phenylindole; hMit, human-speciﬁc mitochondria; MNC, mononuclear cell; PBS, phosphatebuffered saline; vWF, von Willebrand factor; W, week.
at the site of injury. These myoﬁbers were spread uniformly throughout the lacerated area in both the superﬁcial and deep regions of the muscle. Injection of CD133þ cells appeared to most enhance muscle regeneration, increasing the desminpositive area compared with PBS and MNC groups. Also, injection of MNCs was more effective than PBS injection but less effective than CD133þ cell injection (Fig. 4A, 4B). All histological and functional data are summarized in Table 2.
Differentiation of Injected Human Cells into Endothelial Cells and Skeletal Myocytes Differentiation of human cells into endothelial cells was conﬁrmed by double-labeling immunohistochemistry with vWF and hMit antibodies in tissue samples obtained at 1 week after cell injection. Differentiated human ECs, derived from the injected MNCs and CD133þ cells, were observed in the interstitial space between muscle ﬁbers. The number of identiﬁed double-positive cells was greater in the CD133þ group than in the MNC group, which showed a scarce number of cells
(Fig. 4C). As expected, there were no double-positive cells in the PBS group. These ﬁndings indicated that injected CD133þ cells contributed more effectively to vasculogenesis in the injured muscle. Human myoﬁbers derived from the injected MNCs and CD133þ cells were mainly identiﬁed in the periﬁbrosis area by double staining for MyoD1 and hMit in tissue samples at 4 weeks after cell injection. Identiﬁed human myoﬁbers were observed at several points around the periﬁbrosis area in the CD133þ group. There were a scarce number of hMit/ MyoD1-positive ﬁbers in the MNC group. In contrast, differentiated human myoﬁbers were not identiﬁed in the PBS group. From these data, CD133þ cells contributed to myogenesis more effectively in the injured muscle (Fig. 4D).
Unique Gene Expressions of Recipient Animals After CD1331 Cell Injection To further investigate vasculogenesis and myogenesis, we performed real-time PCR with rat tibialis anterior muscle rat-speciﬁc primers for VEGFA and transforming growth
Shi, Ishikawa, Kamei et al.
Table 2. Summary of histological and functional evaluations Evaluation headings
Moist weight ratio (Lt/Rt) 1 wk 4 wk Fast-twitch strength ratio (Lt/Rt) 1 wk 4 wk Tetanus strength ratio (Lt/Rt) 1 wk 4 wk Area of ﬁbrosis (lm2) 1 wk 4 wk Diameter of myoﬁbers (lm) 1 wk 4 wk Number of centronucleated myoﬁbers (n) 1 wk 4 wk Capillary density (n per mm2) 1 wk 4 wk %Fibrosis area (%) 1 wk % Regenerated area (%) 1 wk
0.861 0.028 0.794 0.051
0.880 0.019 0.872 0.019
0.891 0.015 0.908 0.018
0.527 0.015 0.629 0.013
0.614 0.027* 0.672 0.057**
0.729 0.016* 0.819 0.005*
0.673 0.084 0.697 0.036
0.742 0.063** 0.803 0.037*
0.836 0.073* 0.908 0.067*
6416.143 322.671 13669.710 942.549
1537.571 114.473* 5317.714 446.150*
600.429 56.745* 481.714 37.503*
38.070 0.877 79.944 7.159
42.710 0.686* 107.786 1.951*
48.960 0.589* 130.404 2.719*
15.20 1.35 13.40 1.07
18.90 1.28 14.10 0.57
24.00 1.90* 20.70 1.17*
35.561 1.847 50.237 2.859
45.084 0.990* 56.965 3.542
67.845 1.485* 76.432 3.376*
Data are means SEs. n ¼ 5 animals per time point and group. Signiﬁcant difference among groups (one-way analysis of variance and Fisher’s least signiﬁcant difference test): *p < .01, **p < .05. Abbreviations: Lt/Rt, Left/Right; PBS, phosphate-buffered saline; MNC, mononuclear cell.
factor beta 1 (TGFb1), markers for vasculogenesis and ﬁbrosis, and for Pax7, MyoD1, and myogenin, markers for myogenesis. From the macroscopic and histological evaluations described above, we assumed that the initial necrosisdegeneration and inﬂammation phase, which occurs within the ﬁrst minutes and up to 1-2 weeks postinjury, is the most critical phase affecting the injected cells. Therefore, we used tissue samples obtained at day 3 after injury for analysis of gene expressions. Real-time PCR analysis revealed that VEGFA expression at day 3 was signiﬁcantly higher in the CD133þ group compared with the PBS group (PBS group, [0.982 0.081]-fold; MNC group, [1.048 0.096]-fold; CD133 group, [1.414 0.173]-fold, respectively; p < .05 for PBS vs. CD133 group). Conversely, gene expression of TGFb1 was signiﬁcantly lower in the CD133þ group than the PBS group (PBS group, [0.869 0.057]-fold; MNC group, [0.733 0.036]-fold; CD133 group, [0.718 0.038]-fold, respectively; p < .05 for PBS vs. CD133 group) (Fig. 5A, upper panel). Interestingly, from the real-time PCR analysis for myogenic markers, expression of Pax7, an early marker of skeletal muscle progenitor cells, was signiﬁcantly higher in the CD133þ group (PBS group, [0.833 0.075]-fold; MNC group, [0.858 0.052]-fold; CD133 group, [1.880 0.501]-fold, respectively; p < .01 for PBS vs. MNC and PBS vs. CD133 group). Expression of MyoD1 and myogenin, lineage-committed markers, was signiﬁcantly lower than that in the control groups, although there was no signiﬁcant difference in the expressions of Myf5 and Mrf4 among the groups (MyoD1: PBS group, [1.033 0.075]fold; MNC group, [1.051 0.048]-fold; CD133 group, [0.590 0.026]-fold, respectively [p < .01 for PBS vs. CD133 and MNC vs. CD133 group]; Myogenin: PBS group, [1.115 0.109]-fold; MNC group, [0.976 0.083]-fold; CD133 group, [0.488 0.155]-fold, respectively [p < .01 www.StemCells.com
for PBS vs. CD133; p < .05 for MNC vs. CD133 group]; Myf5: PBS group, [0.958 0.090]-fold; MNC group, [0.856 0.089]-fold; CD133 group, [1.150 0.165]-fold, respectively [no signiﬁcant difference]; Mrf4: PBS group, [0.871 0.115]-fold; MNC group, [0.910 0.073]-fold; CD133 group, [1.104 0.203]-fold, respectively [no signiﬁcant difference]) (Fig. 5A, lower panel). To conﬁrm these interesting myogenic gene expression patterns, we performed immunoﬂuorescent staining for Pax7 protein expression. In the CD133þ group, Pax7-positive cells were observed signiﬁcantly around the peri-injured sites at week 1 (Fig. 5B).
The data presented in this report demonstrate that PB-derived CD133þ cell injection improves histological and functional muscular regeneration after skeletal muscle laceration, as indicated by enhanced angiogenesis and reduced ﬁbrosis. Muscle tissue of CD133þ cell-treated animals showed a greater number of capillaries and a larger myoﬁber diameter within the early injury phase. This suggests that acceleration of muscle healing by the endothelial progenitor fraction is due to increased vasculogenesis at the site of injury. The ﬁndings presented here are the ﬁrst demonstration of the successful use of human circulating CD133þ cells to treat severe skeletal muscle injury in rats. Moreover, these ﬁndings indicate that skeletal muscle regeneration with autologous PBderived cells can overcome that achieved with conventional conservative treatment in clinical settings. This report is the ﬁrst to apply human circulating CD133þ cells to a mechanically injured skeletal muscle model for the development of a cell therapy for clinical settings.
CD133þ Cells Accelerate Muscle Regeneration
Figure 5. Real-time polymerase chain reaction analysis for intrinsic unique markers at day 3. (A): VEGFA and TGF-b1 gene expressions were evaluated as unique markers for angiogenesis and ﬁbrosis. In the CD133þ group, VEGFA was upregulated and TGF-b1 was downregulated signiﬁcantly ([A], upper panel). *, p < .05. Pax7, a marker for satellite cells, was signiﬁcantly upregulated, whereas lineage-committed markers such as MyoD1 and myogenin were signiﬁcantly downregulated in the CD133þ group ([A], lower panel). *, p < .05; **, p < .01. (B): Upregulation of Pax7 protein was conﬁrmed by immunoﬂuorescent staining of the sample harvested after 1 week. An abundant number of Pax7-positive cells (red; arrows) were observed in the CD133þ group, whereas only scarce in the control groups. Scale bar ¼ 100 lm. Abbreviations: DAPI, 4,6-diamidino-2phenylindole; MNC, mononuclear cell; n.s., nonsigniﬁcant; PBS, phosphate-buffered saline; TGFb1, transforming growth factor beta 1; VEGFA, vascular endothelial growth factor A.
Currently, the therapeutic potential of EPCs has been investigated not only in the cardiovascular ﬁeld but also in various ﬁelds aimed at regenerative medicine for neurogenesis, osteogenesis, and tenogenesis using various animal models [8, 10-12]. These investigations demonstrated that transplanted human CD34þ cells, commonly known as the hematopoietic stem cell/EPC fraction, have a great potential for initiation of the revascularization switch, which is essential for injured, ischemic tissues. Here, it was conﬁrmed that CD133þ cell injection could accelerate functional skeletal myogenesis via enhancement of vasculogenesis in our study. Moreover, we demonstrated that transplanted CD133þ cells contributed to myogenesis and vasculogenesis in differentiating myoﬁbers and endothelial cells in vivo, although the number of differentiated cells was limited. The formation of vascular systems is essential not only for normal growth, differentiation, and organ function but also for injured organs where the vascular bed is specialized to meet the needs of the organ it supplies. Blood vessel formation is highly regulated by the speciﬁc expression and secretion of factors such as VEGF. VEGF is a major regulator of blood
vessel formation during development and in the adult organism . Recent evidence indicates that this factor also plays an important role in sustaining the proliferation and differentiation of different cell types, including progenitor cells of different tissues, for example, bone marrow, bone, and the central nervous system [30, 31]. Many studies have also demonstrated VEGF upregulation in skeletal muscle regeneration, transplantation, and exercise models [32–37]. Moreover, a recent report demonstrated the regulatory mechanisms by which VEGF is upregulated during skeletal muscle differentiation via signal transduction and protein expression . Bone marrow-derived EPCs also have a potential for stimulating host muscle cells to produce angiogenic factors, such as VEGF under ischemic conditions . Moreover, our recent report showed that transplanted CD133þ cells accelerated axon growth in an organotypic coculture system of rat brain and spinal cord via upregulation of VEGF expression . Expanding on these reports, our present data demonstrated that conduction of the skeletal myogenic environment through VEGF upregulation in the host tissue was induced by CD133þ cell injection.
Shi, Ishikawa, Kamei et al.
It is well known that muscle tissue retains its ability to regenerate after injury and the healing process is slow and often incomplete. The complete recovery of injured skeletal muscle appears to be hindered by ﬁbrosis, which begins during the 2nd week after muscle injury. The resultant disorganized scar tissue that often replaces damaged myoﬁbers may be a contributing factor for the tendency of muscle injuries to recur [40–42]. Usually, growth factors are released at the injured site in muscle injuries, which is an important step in the initiation of the healing process. However, some growth factors are not beneﬁcial for myogenesis because of modulation of muscle ﬁbrosis. In particular, TGF-b1 has been considered a key factor in the development of ﬁbrosis in various tissues [43–47]. A previous report demonstrated that suramin, a TGF-b1 inhibitor, injection into a mouse muscle injury model prevented ﬁbrosis in a dosedependent manner . Interestingly, after CD133þ cell injection, TGF-b1 gene expression was signiﬁcantly lower than that of the control groups 3 days after injury and resulted in reduced ﬁbrosis. Although further studies are required, this response in CD133þ cell-injected tissue is the ﬁrst to demonstrate reduced TGF-b1 gene expression, which could be important data to approach mechanisms of CD133þ cell therapeutic effects. Another approach focuses on stem cells that reside in the muscles of the recipient animal, that is, satellite cells. In our study, Pax7 expression was signiﬁcantly upregulated in the CD133þ cell group, both in gene expression and by immunoﬂuorescent staining. Pax7 is a transcription factor that has been implicated in satellite cell speciﬁcation, survival, and selfrenewal [28, 48-52]. Pax7 is required for the normal function of satellite cells and the productive formation of myogenic precursor cells. Conversely, MyoD, myogenin, Myf5, and Mrf4, the basic helix-loop-helix transcription factors, are the myogenic regulatory factor family members and the terminal differentiation markers [53, 54]. The expression pattern of these genes for muscle regulatory factors observed after CD133þ cell injection were very unique. Based on our observations, our present data might demonstrate that the valance of stemness in injured muscle shifted into a self-renewal stage because of CD133þ cellinduced environmental cues. Interestingly, Messina et al. demonstrated that VEGF had skeletal myogenic effect with an augmented number of satellite cells in mice . In our study, VEGF was upregulated after CD133þ cell injection. From these data, we suggest that CD133þ cells efﬁciently produce skeletal muscle regeneration with Pax7 upregulation, which leads to an increase of myogenic precursor cells via VEGF upregulation in host tissue. However, further investigation is required to deﬁne this stemness shift mechanism. This is the ﬁrst data showing the CD133þ cell potential for activating stem/progenitor cells in recipient tissues. Elucidation of these mechanisms is an important approach for the development of safe and effective cell therapies using the CD133þ/CD34þ cell fraction in clinical settings. Although it remains unclear as to which types of cell source would be the optimal candidate for skeletal muscle regeneration clinically, progenitor cell therapy could be a
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potential new treatment option in skeletal muscle injury. To our knowledge, this is the ﬁrst report that describes the application of human circulating CD133þ cells to animal skeletal muscle injury models for the development of clinical cell therapy. However, we should consider the side effects caused during mobilization and leukoapheresis; even if they are minimum, this autologous circulating population could be relatively more accessible and feasible in clinical setting compared to the embryonic stem cells or mesenchymal stem cells having ethical or facility problems. The present study shows that this cell fraction has an ideal potential for skeletal myogenesis, but there is one limitation, that is, only a limited number of available CD133þ cells in PB. To obtain this rare fraction, administration of growth factors such as G-CSF is required in the clinical settings. In our study, we used human GM-PB CD133þ cells. Therefore, the side effects of mobilization as well as the economical issues have to be considered, to obtain a sufﬁcient number of cells for cell therapy. In this regard, ongoing studies in our laboratory are aimed to at expanding the CD133þ cell subset ex vivo, to facilitate the therapeutic use of these cells at the preclinical and, possibly, clinical levels. Additionally, to develop superior treatments for skeletal muscle injuries, we are analyzing skeletal muscle injuries multidirectionally and are applying the data to animal models, including the use of several transcription factors and micro-RNAs relating to myogenesis in vivo. In the near future, we will clarify the feasible roles and efﬁcacy of these approaches for clinical therapy.
CONCLUSION In conclusion, the data presented here could accelerate the progression of cell-based therapies for skeletal muscle injuries from laboratory to clinical implementation. Moreover, this less-invasive cell-based therapy could be good news for the top athletes suffering from skeletal muscle injuries.
ACKNOWLEDGMENTS This work was supported in part by a grant-in-aid for the 21st Century Center of Excellence Program (F21, Radiation Casualty Medical Research Center to Kenji Kamiya) from the Ministry of Education, Culture, Sports, Science and Technology of Japan and a grant from the General Insurance Association of Japan. We thank Professor M. Hide and K. Ishii (Hiroshima University, Hiroshima, Japan) for equipment and technical support.
POTENTIAL CONFLICTS INTEREST
The authors indicate no potential conﬂicts of interest. 4 5 6
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CD133þ Cells Accelerate Muscle Regeneration
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