Cell and Tissue Research

, Volume 333, Issue 2, pp 207–215 | Cite as

Comparison of mesenchymal tissues-derived stem cells for in vivo chondrogenesis: suitable conditions for cell therapy of cartilage defects in rabbit

  • Hideyuki Koga
  • Takeshi Muneta
  • Tsuyoshi Nagase
  • Akimoto Nimura
  • Young-Jin Ju
  • Tomoyuki Mochizuki
  • Ichiro Sekiya
Regular Article

Abstract

We previously compared mesenchymal stem cells (MSCs) from a variety of mesenchymal tissues and demonstrated that synovium-MSCs had the best expansion and chondrogenic ability in vitro in humans and rats. In this study, we compared the in vivo chondrogenic potential of rabbit MSCs. We also examined other parameters to clarify suitable conditions for in vitro and in vivo cartilage formation. MSCs were isolated from bone marrow, synovium, adipose tissue, and muscle of adult rabbits. Proliferation potential and in vitro chondrogenic potential were compared. Toxicity of the tracer DiI for in vitro chondrogenesis was also examined. MSCs from each tissue were embedded in collagen gel and transplanted into full thickness cartilage defects of rabbits. Cartilage matrix production was compared histologically. The effects of cell density and periosteal patch on the in vivo chondrogenic potential of synovium-MSCs were also examined. Synovium- and muscle-MSCs had a higher proliferation potential than other cells. Pellets from synovium- and bone-marrow-MSCs showed abundant cartilage matrix. DiI had no significant influence on in vitro cartilage formation. After transplantation into cartilage defects, synovium- and bone-marrow-MSCs produced much more cartilage matrix than other cells. When synovium-MSCs were transplanted at a higher cell density and with a periosteal patch, more abundant cartilage matrix was observed. Thus, synovium- and bone-marrow-MSCs had greater in vivo chondrogenic potential than adipose- and muscle-MSCs, but synovium-MSCs had the advantage of a greater proliferation potential. Higher cell density and a periosteum patch were needed to obtain a high production of cartilage matrix by synovium-MSCs.

Keywords

Mesenchymal stem cells Synovium Chondrogenesis Cartilage repair Cell transplantation Rabbit (Japanese White) 

Introduction

Mesenchymal stem cells (MSCs) are an attractive cell source for regenerative medicine because they can be harvested in a minimal invasive manner, and they are easily isolated and expanded, with multipotentiality including chondrogenesis (Prockop 1997; Pittenger et al. 1999; Sekiya et al. 2002). We have previously compared MSCs derived from several kinds of mesenchymal tissues and demonstrated that synovium-derived MSCs have the greatest expansion and chondrogenic ability in vitro than MSCs from other tissues in humans (Sakaguchi et al. 2005) and rats (Yoshimura et al. 2007). This suggests that synovium-derived MSCs are a superior cell source for cartilage regeneration. However, one drawback is that the evaluation of in vitro chondrogenesis may not represent the chondrogenic potential of MSCs transplanted into cartilage defect.

To obtain a suitable outcome of treatment of cell therapy for cartilage injury, clarification of other conditions or parameters and a selection of the cell source is required. Currently, one of the most widely used scaffolds is collagen gel. We have previously demonstrated the suitable conditions for in vitro synovium-derived MSCs/gel composites (Yokoyama et al. 2005); however, appropriate cell density in the gel for in vivo cartilage formation has not been examined. The fluorescent lipophilic tracer, 1,1’-dioctadecyl-3,3,3’,3’- tetramethylindocarbocyanine perchlorate (DiI), is a useful dye for tracing transplanted cells, although leakage to extracellular structures or its toxicity to labeled cells is a concern. Periosteal coverage is a popular method to fix cells or cell/scaffold composites in cartilage defects. However, it would be advantageous if MSCs could be transplanted without a periosteal patch, because the periosteum may unexpectedly be associated with hypertrophy or ossification (Ochi et al. 2002).

In this study, we first compared the proliferation and chondrogenic potential in vitro and in vivo of rabbit MSCs from synovium, bone marrow, adipose, and muscle. Then, we examined the effect of cell density, DiI-labeling, and periosteal coverage for the treatment of cartilage defects with a synovium-derived MSCs/gel composite. These results demonstrated the suitable conditions of MSC therapy for cartilage defects and should advance the clinical application of MSC-based cell therapy for cartilage regeneration.

Materials and methods

Animals

Skeletally mature Japanese White Rabbits weighing about 3.0 kg (range: 2.8–3.5 kg) were used. Animal care was in strict accordance with the guidelines of the animal committee of Tokyo Medical and Dental University. The harvesting of tissues and all operations were performed under anesthesia induced by intramuscular injection of 25 mg/kg ketamine hydrochloride and intravenous injection of 45 mg/kg sodium pentobarbital.

Tissue collection and isolation of cells

Bone marrow aspirate was obtained from the iliac crest with an 18-gauge needle. Synovium with subsynovial tissue was harvested from the knee joint. Adipose tissue was harvested from the perinephric fat tissue intraperitoneally. Muscle was harvested from the anterior tibial muscle. Nucleated cells from the bone marrow were isolated on a density gradient (Ficoll-Paque; Amersham Biosciences, Uppsala, Sweden). Synovium, adipose tissue, and muscle were digested in 3 mg/ml collagenase D (Roche Diagnostics, Mannheim, Germany) in αMEM (Invitrogen, Carlsbad, Calif., USA) at 37°C. After 3 h, digested cells were filtered through a 70-μm nylon filter (Becton Dickinson, Franklin Lakes, N.J., USA), and the remaining tissues were discarded. The digested cells were plated at clonal density in 60-cm2 culture dishes (Nalge Nunc International, Rochester, N.Y., USA) in complete culture medium: αMEM containing 10% fetal bovie serum (FBS; Invitrogen; lot selected for rapid growth of bone-marrow-derived MSCs), 100 U/ml penicillin (Invitrogen), 100 µg/ml streptomycin (Invitrogen), and 250 ng/ml amphotericin B (Invitrogen) and incubated at 37°C with 5% humidified CO2. After 3–4 days, the medium was changed to remove non-adherent cells and then cultured for 14 days as passage 0 without refeeding. The cells were then trypsinized, harvested, and replated as passage 1 at 50 cells/cm2 in 145-cm2 culture dishes (Sekiya et al. 2002). After an additional 14 days of growth, the cells were harvested and cryopreserved. To cryopreserve the cells, they were resuspended at a concentration of 1×106 cells/ml in αMEM with 5% dimethylsulfoxide (Wako, Osaka, Japan) and 20% FBS. Aliquots of 1 ml were slowly frozen and cryopreserved in liquid nitrogen (passage 1). To expand the cells, a frozen vial of the cells was thawed, plated at 50 cells/cm2 in 145-cm2 culture dishes, and incubated for 4 days in the recovery plate. These cells (passage 2) were used for further analyses (Koga et al. 2007).

Culture density and proliferation potential

Passage 1 cells derived from various mesenchymal tissues were plated at 10, 50, and 100 cells/cm2 in 60-cm2 dishes. After 7 and 14 days, cells from three plates from each culture density were harvested and counted with a hemocytometer, and the fold increase was calculated (Mochizuki et al. 2006).

In vitro chondrogenesis assay

Two hundred and fifty thousand cells at passage 2 were placed in a 15-ml polypropylene tube (Becton Dickinson) and centrifuged at 450g for 10 min. The pellets were cultured in chondrogenesis medium consisting of high-glucose Dulbecco’s modified Eagle’s medium (Invitrogen, Carlsbad, Calif., USA) supplemented with 500 ng/ml bone morphogenetic protein-2 (Astellas Pharma, Tokyo, Japan), 10 ng/ml transforming growth factor-β3 (R&D Systems, Minneapolis, Minn., USA), 10–7 M dexamethasone (Sigma-Aldrich, St. Louis, Mo., USA), 50 μg/ml ascorbate-2-phosphate, 40 μg/ml proline, 100 μg/ml pyruvate, and 1:100 diluted ITS + Premix (BD Biosciences, Bedford, Mass., USA; 6.25 µg/ml insulin, 6.25 µg/ml transferrin, 6.25 ng/ml selenious acid, 1.25 mg/ml bovine serum albumin, and 5.35 mg/ml linoleic acid). For microscopy, the pellets were embedded in paraffin, cut into 5-μm sections, and stained with toluidine blue (Sekiya et al. 2001, 2005; Shirasawa et al. 2006). Sections dedicated for fluorescent microscopic visualization of DiI-labeled cells were not stained with toluidine blue. Nuclei were counterstained with 4,6-diamidino-2-phenylindole dihydrochloride (DAPI).

Cell labeling

Passage 2 cells were resuspended at 1×106 cells/ml in αMEM, and the fluorescent lipophilic tracer, DiI (Molecular Probes, Eugene, Ore., USA), was added at 5 μl/ml in αMEM. After incubation for 20 min at 37°C with 5% humidified CO2, the cells were centrifuged at 450g for 5 min, washed twice with phosphate-buffered saline, resuspended, and used for further assay.

Preparation of collagen implants

Passage 2 cells were resuspended in 50 μl αMEM with 20% FBS and mixed with an equal volume of collagen gel (Atelocollagen, 3% type I collagen, Koken, Tokyo, Japan). The mixture was placed into a 6-well plate (Becton Dickinson), and 3 ml αMEM with 20% FBS was added to the plate, which was then incubated at 37°C for 1 day to allow for contraction (Yokoyama et al. 2005).

In vivo transplantation

Skeletally mature Japanese white rabbits were used. The rabbits were anesthetized, the right knee joint was approached through a medial parapatellar incision, and the patella was dislocated laterally. Full thickness osteochondral defects (5×5 mm wide, 3 mm deep) were created in the trochlear groove of the femur, and then the defects were filled with cells/collagen gel composites. The defects were covered with autologous periosteum obtained from the medial proximal tibia, except for a group without a periosteal patch. The periosteum was sutured to each corner of the defect with 6–0 nylon sutures with the cambium layer facing down (Koga et al. 2007). All rabbits were returned to their cages after the operation and were allowed to move freely. Animals were killed with an overdose of sodium pentobarbital at 4 and 12 weeks after the operation. Each sample was examined histologically.

Histological examination

The dissected distal femurs were immediately fixed in a 4% paraformaldehyde solution. Each specimen was decalcified in 4% EDTA solution, dehydrated in a gradient ethanol series, and embedded in paraffin blocks. Sagittal sections of 5 μm thickness were obtained from the center of each defect and stained with toluidine blue. Sections destined for fluorescent microscopic visualization of DiI-labeled cells were not stained with toluidine blue, and nuclei were counterstained by DAPI.

Histological sections of the repaired tissue were analyzed in a blinded manner by two observers who were not informed of the group assignment, and sections were quantified by using the histological grading system for cartilage defects described by Wakitani et al. (1994). This system consisted of five categories (cell morphology, matrix staining, surface regularity, cartilage thickness, and integration of donor with host) scored on a 0– to 14-point scale, in which 0 represented complete regeneration and 14 no regeneration. No significant differences in any of the sections were noted in the scoring between the two observers.

Statistical analysis

To assess differences, a Mann-Whitney U test was used. A value of P < 0.05 was considered significant.

Results

Proliferation potential of rabbit mesenchymal-tissue-derived cells

Synovium- and muscle-derived cells had a higher proliferation potential than bone-marrow- and adipose-derived cells. When plated at lower density, the fold increase was higher in synovium-, adipose-, and muscle-derived cells (Fig. 1).
Fig. 1

Proliferation potential of rabbit mesenchymal stem cells (MSCs) derived from synovium, bone marrow, adipose, and muscle. Cells at passage 1 were plated at 10, 50, and 100 cells/cm2, and the fold increase was calculated after 7 and 14 days (d). Data are expressed as means±SD (n = 3, *P < 0.05; Mann-Whitney U test)

In vitro chondrogenesis

Pellet culture was performed to evaluate chondrogenic potential in vitro. Synovium-derived cells formed larger and heavier pellets than bone-marrow-, adipose-, and muscle-derived cells. In particular, only tiny pellets were obtained in adipose- and muscle-derived cells (Fig. 2a,b). Pellets from synovium- and bone-marrow-derived cells showed a sufficient level of metachromasia, whereas those from adipose- and muscle-derived cells did poorly (Fig. 2c–f).
Fig. 2

In vitro chondrogenic potential of rabbit MSCs derived from synovium, bone marrow, adipose, and muscle. a Macrograph of pellets with a 1-mm scale. b Wet weight of pellets. Data are expressed as means±SD (n = 3, *P < 0.05 in comparison with synovium, **P < 0.05 in comparison with bone marrow; Mann-Whitney U test). c–f Histology of pellets from synovium (c), bone marrow (d), adipose (e) and muscle (f), stained with toluidine blue. Bars 50 μm

Toxicity of DiI

The influence of DiI on the production of cartilage matrix in vitro was examined. Synovium-derived cells were labeled with DiI, and in vitro chondrogenesis potential was evaluated by a pellet culture system. DiI-labeled cell pellets became spherical but were pinkish on gross observation (Fig. 3a). DiI-labeling did not affect histological features, and fluorescence was maintained without leakage of DiI to the extracellular matrix for 21 days (Fig. 3b–g). DiI-labeled cell pellets appeared to be smaller, but no significant difference in the weight of the pellets was noted between the DiI-labeled cells and the control cells (Fig. 3h).
Fig. 3

Influence of DiI on in vitro chondrogenesis. Synovium-derived MSCs with or without DiI-labeling were pelleted and cultured in chondrogenic medium for 21 days. a–g Morphologies of cartilage pellets. a Macrograph with a 1-mm scale. b,c Histological section stained with toluidine blue. Bars 50 μm. d,e Sections examined for DiI. Bars 250 μm. f,g Sections examined for DiI and with DAPI. Bars 25 μm. b,d,f Control pellets. c,e,g DiI-labeled pellets. h Wet weight of pellets. Data are expressed as means±SD (n = 3)

Comparison of in vivo chondrogenic potential

Mesenchymal-tissue-derived cells in each group were suspended in collagen gel and transplanted into the cartilage defects with a periosteal patch to assess in vivo chondrogenic potential. At 4 weeks, cartilage defects transplanted with synovium- and bone-marrow-derived cells were filled with abundant cartilage matrix. On the other hand, cartilage matrix synthesis was poor in the defects with transplanted adipose- and muscle-derived cells (Fig. 4a–d). The histological score was significantly better in transplanted synovium- and bone-marrow-derived cells than in transplanted adipose- and muscle-derived cells (Fig. 4e). For the score of integration category, there were no significant differences among these four groups (Fig. 4f). When synovium-derived cells were transplanted, the border between regenerated cartilage-like tissue and subchondral bone moved upward and closed to the native height at 12 weeks (Fig. 4g). Regenerated cartilage matrix seemed stable and kept its metachromasia. The histological score further improved at 12 weeks (Fig. 4h).
Fig. 4

In vivo chondrogenic potential of rabbit MSCs derived from synovium, bone marrow, adipose, and muscle. a–d Sagittal sections of cartilage defects transplanted with MSCs from synovium (a), bone marrow (b), adipose (c), and muscle (d) at 4 weeks (toluidine blue staining). Bars 1 mm. e Histological scores at 4 weeks. A lower score indicates a better result. Data are expressed as means±SD (n = 3, *P < 0.05 in comparison with synovium, **P < 0.05 in comparison with bone marrow; Mann-Whitney U test). f Histological score for integration of donor with host adjacent cartilage (0 both edges integrated, 2 neither edge integrated. Data are expressed as means±SD (n = 3). g Sagittal sections of cartilage defects transplanted with synovium-derived cells at 12 weeks (toluidine blue staining). Bar 1 mm. h Histological scores of the repair cartilage transplanted with synovium-derived cells at 4 and 12 weeks. A lower score indicates a better result. Data are expressed as means±SD (n = 3, *P < 0.05; Mann-Whitney U test)

Effect of cell density

To assess the effect of cell density on in vivo chondrogenic potential, synovium-derived cells were transplanted at different densities, viz., 5×107 cells/ml and 1×106 cells/ml. At 4 weeks, abundant cartilage matrix was observed in the defect transplanted with a higher density of cells, whereas a small amount of cartilage matrix could be observed in the defect transplanted with a lower density of cells (Fig. 5a,b). The histological score in the higher density group was statistically better than that in the lower density group (Fig. 5c).
Fig. 5

Effect of cell density on in vivo chondrogenic potential of synovium-derived MSCs. a,b Sagittal sections of cartilage defects transplanted with synovium-derived cells at two cell densities at 4 weeks. Synovium-derived cells were embedded in collagen gel at 5×107 cells/ml (high density; a) and at 1×106 cells/ml (low density; b). Bars 1 mm. c Histological scores. A lower score indicates a better result. Data are expressed as means±SD (n = 3, *P < 0.05 by Mann-Whitney U test)

Effect of periosteal patch

Synovium-derived cells/gel composites were transplanted without a periosteal patch to evaluate the effect of the periosteal patch. The cell/gel composites remained in situ, and the cells produced cartilage matrix in the defect (Fig. 6a,b). However, regenerated cartilage was shallower than that of the adjacent native one. Moreover, the surface regularity score in the non-periosteal patch group was significantly worse than that in the periosteal patch group (Fig. 6c).
Fig. 6

Effect of periosteal patch on cartilage repair by synovium-derived MSCs. Synovium-derived cell/collagen gel composites at 5×107 cells/ml were transplanted into cartilage defects without a periosteal patch. a,b Sagittal sections of cartilage defect stained with toluidine blue (a) and for DiI detection (b). Bars 1 mm. c Histological score for surface regularity, in which 0 stands for a smooth surface and 3 for an irregular surface. Data are expressed as means±SD (n = 3, *P < 0.05; Mann-Whitney U test)

Discussion

When considering clinical application of MSC-based cell therapy for cartilage regeneration, suitable conditions for in vivo chondrogenesis of MSCs have to be determined. In this study, we have investigated the conditions suitable for cartilage repair from the standpoint of the cell source, the cell density, and the effect of a periosteal patch.

When rabbit MSCs are compared in vitro, synovium- and bone-marrow-derived cells show much more chondrogenic potential than adipose- and muscle-derived cells. This corresponds to our previous studies in humans (Sakaguchi et al. 2005). The in vivo study has also demonstrated that synovium- and bone-marrow-derived cells have much more chondrogenic potential than adipose- and muscle-derived cells. With regard to proliferation potential, synovium-derived cells have much higher proliferation potential than bone-marrow-derived cells. These findings indicate that synovium-derived cells are a superior cell source for cartilage regeneration.

In the clinical situation, we can observe synovial chondromatosis, an uncommon benign chondrometaplasia (Maurice et al. 1988), osteophytes at the synovium-cartilage junction (Hashimoto et al. 2002), and synovial tissue recruitment to the articular cartilage defect (Hunziker and Rosenberg 1996). These findings confirm the high chondrogenic potential of synovium-derived cells. Although extensive synovial tissue proliferation and inflammation are a concern, no obvious pannus formation and synovitis have been macroscopically or histologically observed in any samples transplanted with synovium-derived cells (data not shown).

The weight of the pellets from synovium-derived cells is significantly heavier than that from bone marrow-derived MSCs, although the difference between the two populations of MSCs is smaller than that between synovium- and adipose-derived cells, and between synovium- and muscle-derived cells. Our results suggest that MSCs with poor chondrogenic potential in vitro cannot differentiate into chondrocytes when the cells are transplanted into a cartilage defect.

Transplanting autologous chondrocytes cultured in collagen gel has been reported for the treatment of full-thickness defects of cartilage (Ochi et al. 2001), the chondrocyte density in the collagen gel applied to the clinical study being 106 cells/ml. Transplantation of bone-marrow-derived and muscle-derived MSCs in collagen gel has also been reported (Adachi et al. 2002; Wakitani et al. 2002), at a cell density of about 6.5×106 and 2×106 cells/ml, respectively. However, these reports do not mention any investigation of the effect of cell density. According to our previous in vitro study, the amount of cartilage matrix is reflected by the initial cell density of human synovium-derived MSCs, and the composites consisting of 5×107 and 108 cells/ml in gel are richer in proteoglycans than those consisting of lower cell densities (Yokoyama et al. 2005). Here, we have transplanted synovial MSCs/gel composites with 5×107 cells/ml successfully, whereas transplantation of synovial MSCs/gel composites containing only 106 cells/ml results in failure. Higher cell density should thus promote the healing of a cartilage defect more effectively, although the preparation of a sufficient number of cells for chondrogenic potential is not always possible. Synovium-derived MSCs are advantageous in terms of their predominant proliferation potential.

We have used DiI, a membrane-bound fluorescent dye, to track the fate of transplanted cells. Previous studies have shown that DiI typically exhibits low cell toxicity and does not compromise cell viability and differentiation potential (Crawford and Braunwald 1991; Ponticiello et al. 2000). DiI also retains its fluorescence for a long time, at least 24 weeks after the transplantation of synovium-derived MSCs for cartilage repair (Koga et al. 2007). Here, we have found that DiI-labeled MSCs maintain their fluorescence during in vitro chondrogenesis, and DiI does not compromise the chondrogenic potential of synovium-derived MSCs, suggesting the utility of DiI in tracking the fate of MSCs transplanted for cartilage repair.

The utility of the periosteum in cartilage regeneration has been reported in various studies. MSCs (Fukumoto et al. 2003) or chondrocyte precursors (Ito et al. 2001) have been isolated from the cambium layer of the periosteum. A successful result in cartilage repair only by periosteum transplantation has been reported (O’Driscoll and Fitzsimmons 2001). We have also reported the partial involvement of periosteum-derived cells in cartilage repair when the MSCs/gel composite was transplanted while covered with periosteum (Koga et al. 2007). It also seems necessary from the standpoint of the risk of leakage of transplanted cells. However, surgical invasiveness to harvest periosteum and complications such as hypertrophy or ossification of the periosteum (Ochi et al. 2002) should be a concern for clinical application.

In this study, we have demonstrated that transplanted MSCs in collagen gel can differentiate into chondrocytes and produce cartilage matrix regardless of the presence or absence of a periosteum. However, transplanted composites cannot extend sufficiently to fill the defects adequately. This is probably attributable to the influence of the collagen gel, which may prevent the production or extension of the cartilage matrix, and the periosteum would be needed to fill the defects to the original height of the joint surface. Development of novel transplantation procedures without periosteum and scaffold might be of value.

With regard to the in vivo chondrogenic assay, we cannot rule out that some synovium-derived MSCs shift to a more fibroblast phenotype and produce tissue of inferior quality when transplanted into cartilage defects. We have collected colony-forming cells as MSCs, and the population is heterogeneous. If only MSCs with a high chondrogenic potential can be selected, fibroblastic cells will be avoided when the population is transplanted into cartilage defects. Some markers have been investigated to select MSCs with a high chondrogenic potential (Simmons and Torok-Storb 1991; Arai et al. 2002; Fickert et al. 2003); however, no conclusive markers confirmed by in vivo experiments have been reported so far. Our in vivo chondrogenic assay suggests that the fraction with high chondrogenic potential is higher in synovium- and bone-marrow-derived cells than in adipose- and muscle-derived cells. This finding will be useful to identify markers to distinguish subpopulation with high chondrogenic potential.

During the process of cartilage repair, dedifferentiation, degeneration, or ossification of hyaline cartilage, some of whose markers are type I and X collagen (Kamekura et al. 2006), are a concern. We have previously reported that type I or type X collagen is not detected, whereas the repaired cartilage still keeps its metachromasia up to 24 weeks after the transplantation of synovium-derived MSCs by a similar method (Koga et al. 2007), indicating the stable hyaline cartilage phenotype of the repaired cartilage.

In this paper, we have only performed a comparative evaluation, not an absolute assessment, and do not claim that the quality of the best regenerated cartilage that we have obtained is comparable with neighboring cartilage. Indeed, cartilage matrix production is just one of the parameters that could be used to evaluate cartilage quality, and mechanical testing will be required to strengthen our results. However, histology often reflects biomechanical property (Kuroki et al. 2007). We consider that this paper provides valuable findings even without additional mechanical testing.

Conclusion

We have demonstrated the conditions suitable for in vivo chondrogenic differentiation in MSCs; the cell source is the synovium, the cell density is high, and the periosteum is needed when the cells are embedded in collagen gel. These findings should advance the clinical application of MSC-based cell therapy for cartilage regeneration.

Notes

Acknowledgements

We thank Dr. Kenichi Shinomiya for continuous support, and Miyoko Ojima for expert help with the histology.

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Copyright information

© Springer-Verlag 2008

Authors and Affiliations

  • Hideyuki Koga
    • 1
  • Takeshi Muneta
    • 1
    • 2
  • Tsuyoshi Nagase
    • 1
  • Akimoto Nimura
    • 1
  • Young-Jin Ju
    • 1
  • Tomoyuki Mochizuki
    • 3
  • Ichiro Sekiya
    • 3
  1. 1.Section of Orthopedic Surgery, Graduate SchoolTokyo Medical and Dental UniversityTokyoJapan
  2. 2.Center of Excellence Program for Frontier Research on Molecular Destruction and Reconstruction of Tooth and BoneTokyo Medical and Dental UniversityTokyoJapan
  3. 3.Section of Cartilage Regeneration, Graduate SchoolTokyo Medical and Dental UniversityTokyoJapan

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