Evaluation of a Collagen Scaffold for Cell-Based Bone Repair Ingeborg J. De Kok, DDS, MS1/Deepali Jere, DDS, MS, MPH2/ Ricardo J. Padilla, DDS3/Lyndon F. Cooper, DDS, PhD4
Purpose: To determine whether a collagen scaffold could provide an environment for mesenchymal stem cell (MSC)–related bone repair of critical-size bone defects in rat calvaria. Materials and Methods: Craniotomy defects were created in 28 adult Sprague-Dawley rats. Two additional rats were used as MSC donors by means of femoral bone marrow lavage and culture. The rats were randomly divided into four groups: (1) empty/no graft; (2) collagen scaffold (matrix) + saline; (3) matrix + MSCs; (4) matrix + bone morphogenetic protein. The animals were euthanized 28 days after surgery. Microcomputed tomographic reconstructions were obtained to measure bone fill. The specimens were processed for histologic examination, and the total defect and bone fill areas were measured. Results: Mean bone fill (± standard deviation) of 9.25% ± 10.82%, 19.07% ± 17.38%, 44.21% ± 3.93%, and 66.06% ± 15.08%, respectively, was observed for the four groups; the differences were statistically significant. Bone repair was statistically significant for groups 3 and 4. No significant difference was seen for bone repair between groups 1 and 2 or between groups 3 and 4. Bone formation differed significantly across the four groups. Statistically significant changes in radiodensity were observed between groups 1 and 3, groups 1 and 4, and groups 2 and 4. Significant differences were not observed between groups 1 and 2, groups 2 and 3, or groups 3 and 4. Conclusion: After grafting of adult MSCs adherent within a collagen matrix, repair of bone was significant. Expanded three-dimensional collagen represents a radiolucent, resorbable, biocompatible scaffold that is capable of supporting MSC repair of bone. Int J Oral Maxillofac Implants 2014;29:e122–e129. doi: 10.11607/jomi.te51 Key words: adult mesenchymal stem cell, bone repair, collagen matrix, osteogenesis, rat calvaria model, tissue engineering
W
ithin an aging population, bone repair and skeletally assisted cosmesis needs are constantly increasing. Among the diverse
1Assistant
Professor, Department of Prosthodontics, School of Dentistry, University of North Carolina, Chapel Hill, North Carolina, USA. 2G raduate Resident, Department of Prosthodontics, School of Dentistry, University of North Carolina, Chapel Hill, North Carolina, USA. 3Clinical Assistant Professor, Department of Diagnostic Sciences and General Dentistry, School of Dentistry, University of North Carolina, Chapel Hill, North Carolina, USA. 4Stallings Distinguished Professor, Department of Prosthodontics, School of Dentistry, University of North Carolina, Chapel Hill, North Carolina, USA. Correspondence to: Dr Lyndon F. Cooper, UNC School of Dentistry, 330 Brauer Hall, Chapel Hill, NC 27599, USA. Fax: +919-966-3821. Email: [email protected] ©2014 by Quintessence Publishing Co Inc.
array of different materials and techniques used to improve bone repair or aid in bone augmentation procedures, autogenous bone is regarded as “gold standard” bone graft material. The advantages of an autogenous material include bone-inductive and bone-conductive properties, as well as a malleable and modestly robust mechanical nature, but autogenous bone grafts are also associated with nonunion failure rates ranging from 5% to 35%,1,2 high costs related to hospitalization, and morbidity and pain at the donor site, which is reported in up to 25% of cases after 2 years. The most common alternative to autografts is the allograft, which, despite its frequent application, is associated with host rejection, excessive resorption, and bone revascularization.3,4 Xenografts are less common alternatives because of concerns related to immunogenicity and disease transmission,5 although in dental therapeutics, xenograft materials for augmentation procedures are a relatively common alterative to autogenous bone.6
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Bone tissue engineering primarily involves osteoinductive signaling molecules, host cells/stem cells responding to the signal, and suitable scaffolding for the stimulated cells. Several growth factors and hormones act as signaling molecules, and they play an important role in osteogenesis by stimulation, differentiation, and protein synthesis in osteoblastic cell culture.7 Molecules that effectively signal bone formation include growth factors and morphogens. Wellknown examples include the bone morphogenetic proteins (BMPs), transforming growth factor beta (TGF-β), insulinlike growth factors I and II, plateletderived growth factor, and basic and acidic fibroblast growth factor. Many of these have been used clinically with success. Presently, BMP-2 is available for orthopedic and craniofacial/dental bone repair and augmentation procedures. BMPs were first identified as the osteo inductive component of bone matrix, and they are members of the TGF-β superfamily, which are present in very low but effective concentrations in the bone matrix.8 At least 20 types of BMPs have been identified in humans to date.9 Several investigators have acknowledged the bone-inductive capacity of recombinant human BMP-2 and BMP-7 in healing criticalsize defects in various animal models such as sheep, rats, and dogs10–13; as such, these represent a benchmark for osteoinductive bone repair in animal models. Cell culture studies have demonstrated that BMP-2 signals bone-specific gene expression associated with both osteoinduction and osteogenesis. This direct action involves receptor activation of kinase activity,14,15 which activates Smad (R-Smad) osteoinductive transcription factor function and the downstream induction of other osteoinductive transcription factors such as Runt-related gene 2, Osterix 2, Distal-less homeobox 5, and Msh-like homeobox 2. The stimulation of progenitor cells in the tissues by engrafted BMP-2 contributes to subsequent osteogenesis.7,9,16 Tissue repair is affected by resident or recruited cells. These progenitor cells may exist within the bone as bone-lining cells, as pericytes, or as recruited mesenchymal stem cells (MSCs). The identification of MSCs of bone marrow as osteoprogenitors or colony-forming-unit osteoblasts (CFU-OB)17 suggested their therapeutic potential, and early experiments using multipotent bone marrow stromal cells18 demonstrated that these MSCs of bone marrow could be readily isolated, expanded, and used to engineer bone repair. Several animal studies have demonstrated the effectiveness of mitotically expanded MSCs in supporting bone regeneration, either alone or together with growth factors.19–21 The multipotent adult stem cell can proliferate and differentiate into various cell types to repair different
tissue types.22 This directional and purposeful differentiation is dependent on many factors, including the scaffold to which the cells adhere for engraftment.23 The majority of investigations regarding MSC-based bone repair have utilized ectopic models of bone repair that have included slowly resorbing or nonresorbable hydroxyapatite or hydroxyapatite/tricalcium phosphate scaffolds to achieve osteogenesis.24,25 They are either granular or preformed in shape, they are not readily resorbed, and they are radiopaque. Other scaffolds include novel ceramics, titanium or tantalum foams, and synthetic and naturally occurring polymeric materials.23 Included are chitosans, polyglycolic (and/or lactic) acids, hydrogels,24 and collagens. One currently available collagen scaffold with clinical uses unrelated to bone repair is an expanded tendon-derived collagen, DuraGen (Integra Lifesciences). It is made from bovine tendon, which is a source of type I collagen.26 Type I collagen is currently used in the manufacture of artificial skin,27 absorbable sponges,28 and wound dressings.29 It conforms to complex surfaces of any shape and size and is fully absorbed after complete tissue closure of the dural defect. Unlike some hydrogel alternatives, the scaffold retains its shape upon hydration and cell loading. Its open architecture may also promote repair by supporting neovascularization and the growth and differentiation of cells. Signals contained within the extracellular matrix (ECM) and released during ECM remodeling bind to cells through receptors such as integrins, and they modulate gene expression, tissue differentiation, and the survival of osteoblasts and fibroblasts.30 The aim of the present project was to determine whether a collagen scaffold would provide an environment for bone marrow–derived stem cell (MSC)–related bone repair of critical-size bone defects in rat calvaria. MSCloaded type I collagen scaffolds were implanted in critical-size rat calvaria defects, and the ensuing osteogenesis was compared with bone repair mediated by recombinant human BMP-2 (rhBMP-2).
MATERIALS AND METHODS Animals and Surgical Protocols This investigation of bone repair employed the rat calvaria critical-size defect model.31 The study used a collagen scaffold (DuraGen) as a common matrix for implanting either rhBMP-2 or MSCs. Saline-infused scaffolds served as one control, and empty defects were used for control of the matrix. Thirty Sprague-Dawley rats (each 5 months old and weighing 250 to 300 g each) were used for this experiment. All procedures were performed according The International Journal of Oral & Maxillofacial Implants e123
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to a protocol approved by the University of North Carolina Institutional Animal Care and Use Committee (#06-082) in Division of Laboratory Animal Medicine– approved and –inspected animal surgery and housing facilities. U.S. National Institutes of Health guidelines for the care and use of laboratory animals (NIH Publication #85-23, rev. 1985) were followed. Four different treatment groups of six to eight rats each were compared; animals were randomly allocated to these four treatment groups. • Group 1: Six animals; defects left empty (negative control) • Group 2: Six animals; defects were grafted with the type I collagen scaffold, which had been treated with 0.5 mL of sterile 0.9% saline (control) • Group 3: Eight animals; defects were grafted with the type I collagen scaffold loaded with rat MSCs expanded from bone marrow (2 hours of adhesion using 5 × 106 cells/mL of rat MSCs in culture media at 37°C in 5% carbon dioxide (CO2) • Group 4: Eight animals; defects were grafted with type I collagen, which had been pretreated for 15 minutes at 37°C with rhBMP-2 (positive control) The remaining two rats served as bone marrow stromal cell donors (syngeneic MSCs)32 by means of femoral bone marrow lavage. The bone marrow adhesion selection method was used to isolate and culture the MSCs.33 The donor rats were sacrificed by CO2 inhalation and the femurs were dissected aseptically. Bone marrow was flushed from the cavities with phosphate-buffered saline (PBS), and the cell suspension was pelleted by centrifugation. Cell pellets were rinsed with serum-free lower glucose– Dulbecco modified Eagle medium (LG-DMEM) (Hyclone), reprecipitated by centrifugation, resuspended in complete LG-DMEM supplemented with penicillin/streptomycin and 10% fetal bovine serum, and transferred into culture flasks. At 80% confluence in 100-mm dishes, cells were detached by treatment with trypsin-ethylenediaminetetraacetic acid (EDTA).34 Cells were then plated in the matrix for 4 hours. The rats were anesthetized presurgically by intraperitoneal injection of ketamine/xylazine (40 to 80 mg/kg and 2 to 10 mg/kg). Immediately after surgery, each animal was given a subcutaneous injection of buprenorphine (0.15 mg/kg; Henry Schein) for postoperative analgesia and an intraperitoneal injection of 3 mL of normal saline (0.9% sodium chloride; Henry Schein). After the skin was opened and a periosteal flap was elevated, a critical-size defect involving the sagittal suture was created without violating the dura. A trephine bur with an external diameter of 8.9 mm was
used (ACE Surgical Supply). Tissue engineering constructs were prepared as described and placed into the surgical defects in groups 2 to 4. The surgical wound was closed in layers with resorbable 5-0 Vicryl sutures (Ethicon). All animals were observed postoperatively until ambulation was noted. Approximately 8 hours after surgery (following recovery from anesthesia), each animal was given another subcutaneous injection of buprenorphine (0.15 mg/kg). Twenty-four hours after surgery, each animal was also given a subcutaneous injection of ketoprofen (5 mg/kg; Henry Schein). Twenty-eight days after surgery, animals were euthanized by CO2 inhalation per standard protocol.
Sample Preparation and Examination Perfusion fixation was accomplished after euthanasia, and the calvaria were recovered and placed in 4% paraformaldehyde solution, which was changed daily. After 2 days of fixation, the specimens were washed with PBS and trimmed, and microcomputed tomographic (microCT) scanning was completed with the Skyscan 1074 microCT scanner (Micro Photonics Inc) to demonstrate the defect size and to quantify the amount of mineralized bone formed within each defect. The specimens were positioned on a holder in the digital image receptor at a right angle to the source of the beam and exposed to 66 kVp, 8 mA, for 0.06 seconds. The images were reconstructed in three dimensions, and densitometric tracing of the CT scans was used to estimate bone fill. A mean grey value (MGV) was calculated for each radiograph using ImageJ 1.37 for Windows, and the total area of newly formed bone in the critical-size defect was determined by MGV measurements for the four treatment groups (range, 0 to 255; higher numbers indicate less bone formation). Three-dimensional images were reconstructed from the CT scans, and an 8-mm-diameter circle was measured centrally within each 8.9-mm defect to define the region of interest. Following completion of the microCT study, the specimens were each placed in 45 mL of 0.5 mol/L EDTA, pH 8.0, for decalcification under constant oscillation at room temperature. Every 3 days, the EDTA buffer was drained and fresh buffer (45 mL) was added. After 30 days of EDTA decalcification, the specimens were bisected along a line perpendicular to the sagittal suture in the center of the defect. At least six 5-µm sections were evaluated across the 8.9 mm of each defect and stained with hematoxylin and eosin and Masson trichrome to enable accurate comparison of the groups. The total areas of the defect and new bone were measured using ImageJ 1.37 software in two randomly selected histologic sections of each animal. The bone fill estimates were calculated as a percentage of the total defect area. Masson trichrome staining was
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Fig 1 Radiomorphometric analysis of newly formed bone in the critical-size defect at 28 days. The total area was determined by MGV measurements in each treatment group.
250
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Fig 2a MicroCT image of empty/negative control site (group 1) at 4 weeks.
Fig 2b MicroCT image of collagen matrix + saline (group 2) at 4 weeks.
also performed on all the slides to identify areas of osteoid production within the critical-size defects. Group means and standard deviations (SDs) were calculated for the percentage of defect area occupied by bone in the microradiographs and the histologic sections. Two evaluators were calibrated for measurement of bone fill using both microCT and histology.
Statistical Analysis Statistical analysis was performed using GraphPad Prism (version 5.04 for Windows, GraphPad Software). Descriptive statistics were obtained using SPSS software (version 15.0, SPSS Inc). Both histomorphometric and microCT data were used to compare new bone formation using analysis of variance (ANOVA). A Tukey test (honestly significant difference) was performed to compare new bone formation in each group. The threshold for statistical significance was set at P < .05.
RESULTS MicroCT Findings Bone formation was evaluated after 28 days of healing for the four groups. The animals in group 1 and 2 had significantly less dense bone in comparison to groups 3 and 4 (Fig 1). Specimens from group 1
Fig 2c MicroCT image of matrix + MSCs (group 3) at 4 weeks.
Fig 2d MicroCT image of matrix + rhBMP-2 (positive control, group 4) at 4 weeks.
(Fig 2a) showed minimal evidence of bone formation at 4 weeks (MGV = 216.96; SD = 13.56). Group 2 specimens (Fig 2b) showed modest bone formation (MGV = 192.97; SD = 32.64), especially around the periphery of the circular defect. Specimens from group 3 had significantly more bone formation (MGV = 174.82; SD = 11.42), both peripherally and centrally, in the cranial critical-size defects (Fig 2c). The amount and density of bone formation were comparable to those seen in group 4 (Fig 2d). Animals in group 4 represented the positive control and had the highest density of new bone (MGV = 149.19; SD = 20.96). Reconstructions of microCT images of specimens from group 3 (Fig 2c) showed significant osteoconduction of the defect by the graft. Reconstruction slices of specimens from group 4 showed extensive osteoconduction and healing of the critical-size defect. One-way ANOVA was used to evaluate the differences in bone fill among the four treatment groups. Bone formation differed significantly across the four groups (F[3,20] = 14.13, P < .0001). The Tukey multiple comparison test to compare the four groups indicated that there was no significant difference between groups 1 and 2 (mean difference, 23.99; 95% confidence interval [CI] –11.79, 59.77), but there were significant differences (P < .05) between groups 1 and 3 (mean difference, 42.14; 95% CI 10.14, 74.14) and between groups 1 and 4 (mean difference, 67.77; The International Journal of Oral & Maxillofacial Implants e125
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Fig 3 Histologic measurements of percentage of bone fill of the critical-size defect at 28 days in the different groups.
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% Bone fill
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Fig 4a Histologic image of Fig 4b Histologic image of col- Fig 4c Histologic image of empty/negative control (group 1) lagen matrix + saline (group 2) at matrix + MSCs (group 3) at 4 at 4 weeks (magnification ×4). 4 weeks (magnification ×4). weeks (magnification ×4).
95% CI 37.83, 97.70). There were no statistically significant differences between groups 2 and 3 (mean difference, 18.15; 95% CI –17.63, 53.92) or between groups 3 and 4 (mean difference, 25.63; 95% CI –4.304, 55.56), but groups 2 and 4 (mean difference = 43.78; 95% CI 9.834, 77.72) were significantly different.
Histomorphometry The quantitative histomorphometric results confirmed the radiographic assessments. After 28 days of healing, measurements were calculated by percent of bone fill of the critical-size defect (Fig 3). Group 1 showed the least amount of bone formation (mean, 9.25%; SD = 10.82%), followed by group 2 (19.07%; SD = 17.38%). Bridging osteogenesis was noted in the healing of groups 3 and 4 sites. Within the treated defects of these groups, new bone formation with rich vasculature and abundant osteoblasts producing osteoid were revealed. Focal lamellar architecture of the new bone was seen (Fig 4). The results showed that mean bone fill in group 3 (44.21%; SD = 3.93) was not significantly different from that in group 4 (60.06%; SD = 15.08).
Fig 4d Histologic image of matrix + rhBMP-2 (group 4) at 4 weeks (magnification ×4).
A one-way ANOVA was used to compare the differences in bone fill percentage in the four treatment groups; significant differences were seen among all groups (F[3,27] = 22.08; P < .001). The Tukey multiple comparison test indicated that there was no significant difference between groups 1 and 2 (mean difference, –9.821; 95% CI –32.53, 12.89), but there were statistically significant differences (P < .05) between groups 1 and 3 (mean difference, –34.96; 95% CI –55.27 to –14.65) and between groups 1 and 4 (mean difference, 50.81; 95% CI –69.81, –31.81). While there was general agreement between the findings of microCT and histomorphometry, the microCT results revealed differences between groups 2 and 3 (mean difference, –25.14; 95% CI –47.85, –2.431) and between groups 2 and 4 (mean difference, –40.99; 95% CI –62.53, –19.45). Comparisons between groups 3 and 4 were not statistically significant at P < .05 (mean difference, –15.85; 95% CI –34.85 to 3.148). Masson trichrome staining was applied to detect osteoid tissue (Fig 5). Osteoid formation was observed in all the animals from groups 3 and 4, but it was observed only occasionally in groups 1 and 2.
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Fig 5 Histomorphometric assessment of cell-based bone repair from group 4 animal reveals the edge of the defect on the lower left corner and the new bone within the defect to the top right corner. The original bone shows a lamellar pattern of ossification, while the new bone within the defect appears woven (Masson trichrome).
DISCUSSION The rat calvaria critical-size defect model is a commonly used and convenient wasy to screen and evaluate bone regeneration materials. The rat cranium is a suitable site for grafting studies because the graft is protected from relative motion, which eliminates the need for fixation.35 This orthotopic model differs from the ectopic model of osteoinduction per se, but it represents an endosseous situation that is typical of many clinical situations. A type I collagen (DuraGen) was used as a scaffold for both MSCs and rhBMP-2. The scaffolds were prepared from a commercially available material as 8-mm disks approximately 1.0 mm thick. This collagen matrix was shown to be successful in accelerating dural healing and containing cerebrospinal fluid in cases studied by Narotam and colleagues in 2004.36 Its biocompatibility is further suggested by its support of the survival of cerebral cortical neurons in vitro.37 Danish et al compared the performance and complications of DuraGen and AlloDerm for duraplasty and concluded that both materials could be safe alternatives.38 Preliminary experiments using vital dye staining of MSCs showed that DuraGen was readily loaded with human MSCs applied over a 2- to 4-hour period from densely populated culture media (5 × 106 cells/100 µL) (data not shown). It was assumed that the rhBMP-2 was adherent to the engrafted DuraGen material using the 15-minute incubation procedure required for another type I collagen scaffold. Several studies have observed the binding capacity of demineralized bone matrix to osteoinductive growth factors of the TGF family such as BMPs.39 It is not surprising that the collagen scaffold alone failed to promote remarkable bone repair. Neither did it result in inflammatory reactions. The selected scaffold may be a suitable scaffold for osteoinductive agents,
but it was not capable of promoting critical-size defect repair alone. In contrast, Sweeney and colleagues found that animals treated with type I collagen gels alone showed bone repair of 92.5% at 20 weeks.40 Saadeh et al also demonstrated the ability of type I collagen to heal critical-sized bony defects in the rat mandible. He suggested that type I collagen would make a suitable carrier matrix for improved approaches to bone tissue engineering.41 In the present study, a much shorter 4-week healing period was evaluated. Further, these defects were large (8.9 mm) and were performed in adult animals. The present results suggest that the use of type I collagen scaffold for MSCbased bone repair may be effective in promoting early repair of large defects. Here, rhBMP-2 was utilized as a positive control for bone repair in the critical-size defect. In this case, the same type I collagen scaffold was utilized for both rhBMP-2 and MSCs. Other investigations have clearly demonstrated that rhBMP-2 is capable of affecting bone repair of critical-size calvaria defects. The 5-µg/mL solution used for adsorption of rhBMP-2 onto the collagen scaffold does not represent a remarkably high concentration of morphogen, in comparison to many other investigations. Regardless, the present results with this type I collagen scaffold confirm earlier reports of Chen and collaborators, who tested a collagen-based BMP-2–targeting bone repair system in a rabbit mandibular critical-size defect model and concluded that the improved bone formation could be attributed to the retention of BMP-2 by collagen fibrils.42 The present results further support the exploration of collagen as a basis for rhBMP-2 scaffolds. The engrafted MSCs successfully repaired the calvarial defects without additional osteoinductive agents. It is inferred that the type I scaffold enabled engraftment of sufficient cells and maintained their viability The International Journal of Oral & Maxillofacial Implants e127
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and osteoinductive nature for long enough to engender defect repair. Others have suggested that this type of scaffold supports the differentiation of MSCs into osteoblasts.43 In an in vitro study, a compatible commercial collagen scaffold possessed the ability to support rat MSC osteogenic differentiation as indicated by alkaline phosphatase, osteopontin, and osteocalcin expression.44 In this preliminary study, little effort was made to optimize cell attachment or cell density to achieve bone repair. It is notable that there was great consistency in the bone repair that occurred within 28 days for the cell-engrafted samples. MSC adhesion to type I collagen is well characterized,45 and, while the efficacy of loading these scaffolds has not yet been examined in detail, the results indicate that the scaffold supported delivery of the MSCs to the surgical wounds to evoke direct or indirect effects to encourage bone repair. It is further inferred from the osteogenesis result that the type I collagen scaffold provided a suitable adhesive substrate for MSC differentiation along the osteoblastic pathway. Cell culture studies confirm that type I collagen adhesion of MSCs supports osteoinduction.46 Further experimental application of this collagen scaffold may permit assessment of the minimal cell density/dose required for defined volumetric repair of bone defects. Radiographic assessment of bone formation and three-dimensional reconstruction allowed the authors to visualize the cranial defect from the axial view, which was not possible in histologic sections. A further advantage of radiomorphometry is that it can be performed rapidly after specimen retrieval and can serve as a screening assay. Tabletop systems can inherently induce a discrepancy in the observed MGVs and local bone density, which has been related to the effect of beam hardening. However, this error is assumed to be similar and mutually neutralized for all specimens, since they were comparably of the same size and composition.47 Based on the present data, the radiomorphometric data correlate well with the histomorphometric data. Radiomorphometry was less sensitive than histomorphometry, as indicated by the lack of significant difference seen between groups 2 and 3 in the microCT data. Many authors have validated the use of microCT as a tool for three-dimensional assessment of bone structure for preclinical trials, yet they recommend the use of histology for more conclusive results47,48 and still consider histomorphometry the “gold standard.” The use of microCT for rapid evaluation of bone repair using radiolucent scaffolds such as collagen-based hydrogels is supported by the present data. Resorption of the collagen scaffold prior to completion of bone formation may lead to incomplete bone
fill. Although this may not be a problem with rat MSCs because of their ability to quickly regenerate bone, this could be a potential problem to consider when using such materials in clinical studies with adult human MSCs.
CONCLUSIONS In a rat calvaria critical-size defect model, rat mesenchymal stem cells implanted in a type I collagen matrix promoted bone repair comparable to that induced by recombinant human bone morphogenetic protein 2 in the same matrix material. The use of type I collagen scaffolds for engraftment of mesenchymal stem cells to repair orthotopic defects may be considered in further translational investigations and in clinical practice.
ACKNOWLEDGMENTS This research project was supported by the 2006 Tylman Research Grant from the American Academy of Fixed Prosthodontics. The authors thank Ms Sara Valencia, dental student, and Dr Gustavo Mendonca, Clinical Assistant Professor, Department of Prostho dontics, University of North Carolina School of Dentistry, for their immense help in working with the stem cells and statistical analysis. The authors reported no conflicts of interest related to this study.e
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11. Gerhart TN, Kirker Head CA, Kriz MJ, et al. Healing segmental femoral defects in sheep using recombinant human bone morphogenetic protein. Clin Orthop Relat Res 1993; Aug(293):317–326. 12. Stevenson S, Cunningham N, Toth J, Davy D, Reddi AH. The effect of osteogenin (a bone morphogenetic protein) on the formation of bone in orthotopic segmental defects in rats. J Bone Joint Surg Am 1994;76:1676–1687. 13. Itoh T, Mochizuki M, Fuda K, et al. Femoral nonunion fracture treated with recombinant human bone morphogenetic protein-2 in a dog. J Vet Med Sci 1998;60:535–538. 14. Heldin CH, Miyazono K, ten Dijke P. TGF-beta signalling from cell membrane to nucleus through SMAD proteins. Nature 1997;390:465–471. 15. Ryoo HM, Lee MH, Kim YJ. Critical molecular switches involved in BMP-2-induced osteogenic differentiation of mesenchymal cells. Gene 2006;366:51–57. 16. Solheim E. Growth factors in bone. Int Orthop 1998;22: 410–416. 17. Friedenstein AJ. Stromal mechanisms of bone marrow: Cloning in vitro and retransplantation in vivo. Haematol Blood Transfus 1980;25:19–29. 18. Friedenstein AJ, Piatetzky S II, Petrakova KV. Osteogenesis in transplants of bone marrow cells. J Embryol Exp Morphol 1966;16:381–390. 19. Krebsbach PH, Mankani MH, Satomura K, Kuznetsov SA, Robey PG. Repair of craniotomy defects using bone marrow stromal cells. Transplantation 1998;66:1272–1278. 20. Petite H, Viateau V, Bensaid W, et al. Tissue-engineered bone regeneration. Nat Biotechnol 2000;18:959–963. 21. De Kok IJ, Peter SJ, Archambault M, et al. Investigation of allogeneic mesenchymal stem cell-based alveolar bone formation: Preliminary findings. Clin Oral Implants Res 2003; 14:481–489. 22. Bianco P, Riminucci M, Gronthos S, Robey PG. Bone marrow stromal stem cells: Nature, biology, and potential applications. Stem Cells 2001;19:180–192. 23. Caplan AI, Bruder SP. Mesenchymal stem cells: Building blocks for molecular medicine in the 21st century. Trends Mol Med 2001;7:259–264. 24. Harris CT, Cooper LF. Comparison of bone graft matrices for human mesenchymal stem cell-directed osteogenesis. J Biomed Mater Res A 2004;68:747–755. 25. De Kok IJ, Hicok KC, Padilla RJ, Young RG, Cooper LF. Effect of vitamin D pretreatment of human mesenchymal stem cells on ectopic bone formation. J Oral Implantol 2006;32: 103–109. 26. Beghe F, Menicagli C, Neggiani P, et al. Lyophilized nondenatured type-I collagen (Condress) extracted from bovine Achilles’ tendon and suitable for clinical use. Int J Tissue React 1992;14(suppl):11–19. 27. Wanitphakdeedecha R, Chen TM, Nguyen TH. The use of acellular, fetal bovine dermal matrix for acute, full-thickness wounds. J Drugs Dermatol 2008;7:781–784. 28. Chen YH, Dong WR, Xiao YQ, Zhao BL, Hu GD, An LB. Preparation and bioactivity of human hair keratin-collagen sponge, a new type of dermal analogue. Nan Fang Yi Ke Da Xue Xue Bao 2006;26:131–138. 29. Leipziger LS, Glushko V, DiBernardo B, et al. Dermal wound repair: Role of collagen matrix implants and synthetic polymer dressings. J Am Acad Dermatol 1985;12(2 Pt 2):409–419. 30. Damsky CH. Extracellular matrix-integrin interactions in osteoblast function and tissue remodeling. Bone 1999;25: 95–96.
31. Schmitz JP, Hollinger JO. The critical size defect as an experimental model for craniomandibulofacial nonunions. Clin Orthop Relat Res 1986;:299–308. 32. Kotobuki N, Katsube Y, Katou Y, Tadokoro M, Hirose M, Ohgushi H. In vivo survival and osteogenic differentiation of allogeneic rat bone marrow mesenchymal stem cells (MSCs). Cell Transplant 2008;17:705–712. 33. Sekiya I, Larson BL, Smith JR, Pochampally R, Cuii JG, Prockop DJ. Expansion of human adult stem cells from bone marrow stroma: Conditions that maximize the yields of early progenitors and evaluate their quality. Stem Cells 2002;20: 530–541. 34. Mendonca G, Mendonca DB, Simoes LG, et al. Nanostructured alumina-coated implant surface: Effect on osteoblastrelated gene expression and bone-to-implant contact in vivo. Int J Oral Maxillofac Implants 2009;24:205–215. 35. Wang J, Glimcher MJ. Characterization of matrix-induced osteogenesis in rat calvarial bone defects: II. Origins of bone-forming cells. Calcif Tissue Int 1999;65:486–493. 36. Narotam PK, José S, Nathoo N, Taylon C, Vora Y. Collagen matrix (DuraGen) in dural repair: Analysis of a new modified technique. Spine (Phila Pa 1976) 2004;29:2861–2867. 37. Rabinowitz L, Monnerie H, Shashidhara S, Le Roux PD. Growth of rat cortical neurons on DuraGen, a collagen-based dural graft matrix. Neurol Res 2005;27:887–894. 38. Danish SF, Samdani A, Hanna A, Storm P, Sutton L. Experience with acellular human dura and bovine collagen matrix for duraplasty after posterior fossa decompression for Chiari malformations. J Neurosurg 2006;104(1 suppl):16–20. 39. Zhao Y, Chen B, Lin H, et al. The bone-derived collagen containing mineralized matrix for the loading of collagenbinding bone morphogenetic protein-2. J Biomed Mater Res A 2009;88:725–734. 40. Sweeney TM, Opperman LA, Persing JA, Ogle RC. Repair of critical size rat calvarial defects using extracellular matrix protein gels. J Neurosurg 1995;83:710–715. 41. Saadeh PB, Khosla RK, Mehrara BJ, et al. Repair of a critical size defect in the rat mandible using allogenic type I collagen. J Craniofac Surg 2001;12:573–579. 42. Chen D, Zhao M, Mundy GR. Bone morphogenetic proteins. Growth Factors 2004;22:233–241. 43. Neuss S, Stainforth R, Salber J, et al. Long-term survival and bipotent terminal differentiation of human mesenchymal stem cells (hMSC) in combination with a commercially available three-dimensional collagen scaffold. Cell Transplant 2008;17: 977–986. 44. Donzelli E, Salvadè A, Mimo P, et al. Mesenchymal stem cells cultured on a collagen scaffold: In vitro osteogenic differentiation. Arch Oral Biol 2007;52:64–73. 45. Wang H, Li Y, Zuo Y, Li J, Ma S, Cheng L. Biocompatibility and osteogenesis of biomimetic nano-hydroxyapatite/ polyamide composite scaffolds for bone tissue engineering. Biomaterials 2007;28:3338–3348. 46. Na K, Kim SW, Sun BK, et al. Osteogenic differentiation of rabbit mesenchymal stem cells in thermo-reversible hydrogel constructs containing hydroxyapatite and bone morphogenic protein-2 (BMP-2). Biomaterials 2007;28:2631–2637. 47. Verna C, Dalstra M, Wikeshö UM, Trombelli L; Carles Bosch. Healing patterns in calvarial bone defects following guided bone regeneration in rats. A micro-CT scan analysis. J Clin Periodontol 2002;29:865–870. 48. Engelke K, Karolczak M, Lutz A, Seibert U, Schaller S, Kalender W. Micro-CT. Technology and application for assessing bone structure [in German]. Radiologie 1999;39: 203–212.
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Purpose: To determine whether a collagen scaffold could provide an environment for mesenchymal stem cell (MSC)–related bone repair of critical-size bone defects in rat calvaria. Materials and Methods: Craniotomy defects were created in 28 adult Sprague-Dawley rats. Two additional rats were used as MSC donors by means of femoral bone marrow lavage and culture. The rats were randomly divided into four groups: (1) empty/no graft; (2) collagen scaffold (matrix) + saline; (3) matrix + MSCs; (4) matrix + bone morphogenetic protein. The animals were euthanized 28 days after surgery. Microcomputed tomographic reconstructions were obtained to measure bone fill. The specimens were processed for histologic examination, and the total defect and bone fill areas were measured. Results: Mean bone fill (± standard deviation) of 9.25% ± 10.82%, 19.07% ± 17.38%, 44.21% ± 3.93%, and 66.06% ± 15.08%, respectively, was observed for the four groups; the differences were statistically significant. Bone repair was statistically significant for groups 3 and 4. No significant difference was seen for bone repair between groups 1 and 2 or between groups 3 and 4. Bone formation differed significantly across the four groups. Statistically significant changes in radiodensity were observed between groups 1 and 3, groups 1 and 4, and groups 2 and 4. Significant differences were not observed between groups 1 and 2, groups 2 and 3, or groups 3 and 4. Conclusion: After grafting of adult MSCs adherent within a collagen matrix, repair of bone was significant. Expanded three-dimensional collagen represents a radiolucent, resorbable, biocompatible scaffold that is capable of supporting MSC repair of bone. Int J Oral Maxillofac Implants 2014;29:e122–e129. doi: 10.11607/jomi.te51 Key words: adult mesenchymal stem cell, bone repair, collagen matrix, osteogenesis, rat calvaria model, tissue engineering
W
ithin an aging population, bone repair and skeletally assisted cosmesis needs are constantly increasing. Among the diverse
1Assistant
Professor, Department of Prosthodontics, School of Dentistry, University of North Carolina, Chapel Hill, North Carolina, USA. 2G raduate Resident, Department of Prosthodontics, School of Dentistry, University of North Carolina, Chapel Hill, North Carolina, USA. 3Clinical Assistant Professor, Department of Diagnostic Sciences and General Dentistry, School of Dentistry, University of North Carolina, Chapel Hill, North Carolina, USA. 4Stallings Distinguished Professor, Department of Prosthodontics, School of Dentistry, University of North Carolina, Chapel Hill, North Carolina, USA. Correspondence to: Dr Lyndon F. Cooper, UNC School of Dentistry, 330 Brauer Hall, Chapel Hill, NC 27599, USA. Fax: +919-966-3821. Email: [email protected] ©2014 by Quintessence Publishing Co Inc.
array of different materials and techniques used to improve bone repair or aid in bone augmentation procedures, autogenous bone is regarded as “gold standard” bone graft material. The advantages of an autogenous material include bone-inductive and bone-conductive properties, as well as a malleable and modestly robust mechanical nature, but autogenous bone grafts are also associated with nonunion failure rates ranging from 5% to 35%,1,2 high costs related to hospitalization, and morbidity and pain at the donor site, which is reported in up to 25% of cases after 2 years. The most common alternative to autografts is the allograft, which, despite its frequent application, is associated with host rejection, excessive resorption, and bone revascularization.3,4 Xenografts are less common alternatives because of concerns related to immunogenicity and disease transmission,5 although in dental therapeutics, xenograft materials for augmentation procedures are a relatively common alterative to autogenous bone.6
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Bone tissue engineering primarily involves osteoinductive signaling molecules, host cells/stem cells responding to the signal, and suitable scaffolding for the stimulated cells. Several growth factors and hormones act as signaling molecules, and they play an important role in osteogenesis by stimulation, differentiation, and protein synthesis in osteoblastic cell culture.7 Molecules that effectively signal bone formation include growth factors and morphogens. Wellknown examples include the bone morphogenetic proteins (BMPs), transforming growth factor beta (TGF-β), insulinlike growth factors I and II, plateletderived growth factor, and basic and acidic fibroblast growth factor. Many of these have been used clinically with success. Presently, BMP-2 is available for orthopedic and craniofacial/dental bone repair and augmentation procedures. BMPs were first identified as the osteo inductive component of bone matrix, and they are members of the TGF-β superfamily, which are present in very low but effective concentrations in the bone matrix.8 At least 20 types of BMPs have been identified in humans to date.9 Several investigators have acknowledged the bone-inductive capacity of recombinant human BMP-2 and BMP-7 in healing criticalsize defects in various animal models such as sheep, rats, and dogs10–13; as such, these represent a benchmark for osteoinductive bone repair in animal models. Cell culture studies have demonstrated that BMP-2 signals bone-specific gene expression associated with both osteoinduction and osteogenesis. This direct action involves receptor activation of kinase activity,14,15 which activates Smad (R-Smad) osteoinductive transcription factor function and the downstream induction of other osteoinductive transcription factors such as Runt-related gene 2, Osterix 2, Distal-less homeobox 5, and Msh-like homeobox 2. The stimulation of progenitor cells in the tissues by engrafted BMP-2 contributes to subsequent osteogenesis.7,9,16 Tissue repair is affected by resident or recruited cells. These progenitor cells may exist within the bone as bone-lining cells, as pericytes, or as recruited mesenchymal stem cells (MSCs). The identification of MSCs of bone marrow as osteoprogenitors or colony-forming-unit osteoblasts (CFU-OB)17 suggested their therapeutic potential, and early experiments using multipotent bone marrow stromal cells18 demonstrated that these MSCs of bone marrow could be readily isolated, expanded, and used to engineer bone repair. Several animal studies have demonstrated the effectiveness of mitotically expanded MSCs in supporting bone regeneration, either alone or together with growth factors.19–21 The multipotent adult stem cell can proliferate and differentiate into various cell types to repair different
tissue types.22 This directional and purposeful differentiation is dependent on many factors, including the scaffold to which the cells adhere for engraftment.23 The majority of investigations regarding MSC-based bone repair have utilized ectopic models of bone repair that have included slowly resorbing or nonresorbable hydroxyapatite or hydroxyapatite/tricalcium phosphate scaffolds to achieve osteogenesis.24,25 They are either granular or preformed in shape, they are not readily resorbed, and they are radiopaque. Other scaffolds include novel ceramics, titanium or tantalum foams, and synthetic and naturally occurring polymeric materials.23 Included are chitosans, polyglycolic (and/or lactic) acids, hydrogels,24 and collagens. One currently available collagen scaffold with clinical uses unrelated to bone repair is an expanded tendon-derived collagen, DuraGen (Integra Lifesciences). It is made from bovine tendon, which is a source of type I collagen.26 Type I collagen is currently used in the manufacture of artificial skin,27 absorbable sponges,28 and wound dressings.29 It conforms to complex surfaces of any shape and size and is fully absorbed after complete tissue closure of the dural defect. Unlike some hydrogel alternatives, the scaffold retains its shape upon hydration and cell loading. Its open architecture may also promote repair by supporting neovascularization and the growth and differentiation of cells. Signals contained within the extracellular matrix (ECM) and released during ECM remodeling bind to cells through receptors such as integrins, and they modulate gene expression, tissue differentiation, and the survival of osteoblasts and fibroblasts.30 The aim of the present project was to determine whether a collagen scaffold would provide an environment for bone marrow–derived stem cell (MSC)–related bone repair of critical-size bone defects in rat calvaria. MSCloaded type I collagen scaffolds were implanted in critical-size rat calvaria defects, and the ensuing osteogenesis was compared with bone repair mediated by recombinant human BMP-2 (rhBMP-2).
MATERIALS AND METHODS Animals and Surgical Protocols This investigation of bone repair employed the rat calvaria critical-size defect model.31 The study used a collagen scaffold (DuraGen) as a common matrix for implanting either rhBMP-2 or MSCs. Saline-infused scaffolds served as one control, and empty defects were used for control of the matrix. Thirty Sprague-Dawley rats (each 5 months old and weighing 250 to 300 g each) were used for this experiment. All procedures were performed according The International Journal of Oral & Maxillofacial Implants e123
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to a protocol approved by the University of North Carolina Institutional Animal Care and Use Committee (#06-082) in Division of Laboratory Animal Medicine– approved and –inspected animal surgery and housing facilities. U.S. National Institutes of Health guidelines for the care and use of laboratory animals (NIH Publication #85-23, rev. 1985) were followed. Four different treatment groups of six to eight rats each were compared; animals were randomly allocated to these four treatment groups. • Group 1: Six animals; defects left empty (negative control) • Group 2: Six animals; defects were grafted with the type I collagen scaffold, which had been treated with 0.5 mL of sterile 0.9% saline (control) • Group 3: Eight animals; defects were grafted with the type I collagen scaffold loaded with rat MSCs expanded from bone marrow (2 hours of adhesion using 5 × 106 cells/mL of rat MSCs in culture media at 37°C in 5% carbon dioxide (CO2) • Group 4: Eight animals; defects were grafted with type I collagen, which had been pretreated for 15 minutes at 37°C with rhBMP-2 (positive control) The remaining two rats served as bone marrow stromal cell donors (syngeneic MSCs)32 by means of femoral bone marrow lavage. The bone marrow adhesion selection method was used to isolate and culture the MSCs.33 The donor rats were sacrificed by CO2 inhalation and the femurs were dissected aseptically. Bone marrow was flushed from the cavities with phosphate-buffered saline (PBS), and the cell suspension was pelleted by centrifugation. Cell pellets were rinsed with serum-free lower glucose– Dulbecco modified Eagle medium (LG-DMEM) (Hyclone), reprecipitated by centrifugation, resuspended in complete LG-DMEM supplemented with penicillin/streptomycin and 10% fetal bovine serum, and transferred into culture flasks. At 80% confluence in 100-mm dishes, cells were detached by treatment with trypsin-ethylenediaminetetraacetic acid (EDTA).34 Cells were then plated in the matrix for 4 hours. The rats were anesthetized presurgically by intraperitoneal injection of ketamine/xylazine (40 to 80 mg/kg and 2 to 10 mg/kg). Immediately after surgery, each animal was given a subcutaneous injection of buprenorphine (0.15 mg/kg; Henry Schein) for postoperative analgesia and an intraperitoneal injection of 3 mL of normal saline (0.9% sodium chloride; Henry Schein). After the skin was opened and a periosteal flap was elevated, a critical-size defect involving the sagittal suture was created without violating the dura. A trephine bur with an external diameter of 8.9 mm was
used (ACE Surgical Supply). Tissue engineering constructs were prepared as described and placed into the surgical defects in groups 2 to 4. The surgical wound was closed in layers with resorbable 5-0 Vicryl sutures (Ethicon). All animals were observed postoperatively until ambulation was noted. Approximately 8 hours after surgery (following recovery from anesthesia), each animal was given another subcutaneous injection of buprenorphine (0.15 mg/kg). Twenty-four hours after surgery, each animal was also given a subcutaneous injection of ketoprofen (5 mg/kg; Henry Schein). Twenty-eight days after surgery, animals were euthanized by CO2 inhalation per standard protocol.
Sample Preparation and Examination Perfusion fixation was accomplished after euthanasia, and the calvaria were recovered and placed in 4% paraformaldehyde solution, which was changed daily. After 2 days of fixation, the specimens were washed with PBS and trimmed, and microcomputed tomographic (microCT) scanning was completed with the Skyscan 1074 microCT scanner (Micro Photonics Inc) to demonstrate the defect size and to quantify the amount of mineralized bone formed within each defect. The specimens were positioned on a holder in the digital image receptor at a right angle to the source of the beam and exposed to 66 kVp, 8 mA, for 0.06 seconds. The images were reconstructed in three dimensions, and densitometric tracing of the CT scans was used to estimate bone fill. A mean grey value (MGV) was calculated for each radiograph using ImageJ 1.37 for Windows, and the total area of newly formed bone in the critical-size defect was determined by MGV measurements for the four treatment groups (range, 0 to 255; higher numbers indicate less bone formation). Three-dimensional images were reconstructed from the CT scans, and an 8-mm-diameter circle was measured centrally within each 8.9-mm defect to define the region of interest. Following completion of the microCT study, the specimens were each placed in 45 mL of 0.5 mol/L EDTA, pH 8.0, for decalcification under constant oscillation at room temperature. Every 3 days, the EDTA buffer was drained and fresh buffer (45 mL) was added. After 30 days of EDTA decalcification, the specimens were bisected along a line perpendicular to the sagittal suture in the center of the defect. At least six 5-µm sections were evaluated across the 8.9 mm of each defect and stained with hematoxylin and eosin and Masson trichrome to enable accurate comparison of the groups. The total areas of the defect and new bone were measured using ImageJ 1.37 software in two randomly selected histologic sections of each animal. The bone fill estimates were calculated as a percentage of the total defect area. Masson trichrome staining was
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De Kok et al
Fig 1 Radiomorphometric analysis of newly formed bone in the critical-size defect at 28 days. The total area was determined by MGV measurements in each treatment group.
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Fig 2a MicroCT image of empty/negative control site (group 1) at 4 weeks.
Fig 2b MicroCT image of collagen matrix + saline (group 2) at 4 weeks.
also performed on all the slides to identify areas of osteoid production within the critical-size defects. Group means and standard deviations (SDs) were calculated for the percentage of defect area occupied by bone in the microradiographs and the histologic sections. Two evaluators were calibrated for measurement of bone fill using both microCT and histology.
Statistical Analysis Statistical analysis was performed using GraphPad Prism (version 5.04 for Windows, GraphPad Software). Descriptive statistics were obtained using SPSS software (version 15.0, SPSS Inc). Both histomorphometric and microCT data were used to compare new bone formation using analysis of variance (ANOVA). A Tukey test (honestly significant difference) was performed to compare new bone formation in each group. The threshold for statistical significance was set at P < .05.
RESULTS MicroCT Findings Bone formation was evaluated after 28 days of healing for the four groups. The animals in group 1 and 2 had significantly less dense bone in comparison to groups 3 and 4 (Fig 1). Specimens from group 1
Fig 2c MicroCT image of matrix + MSCs (group 3) at 4 weeks.
Fig 2d MicroCT image of matrix + rhBMP-2 (positive control, group 4) at 4 weeks.
(Fig 2a) showed minimal evidence of bone formation at 4 weeks (MGV = 216.96; SD = 13.56). Group 2 specimens (Fig 2b) showed modest bone formation (MGV = 192.97; SD = 32.64), especially around the periphery of the circular defect. Specimens from group 3 had significantly more bone formation (MGV = 174.82; SD = 11.42), both peripherally and centrally, in the cranial critical-size defects (Fig 2c). The amount and density of bone formation were comparable to those seen in group 4 (Fig 2d). Animals in group 4 represented the positive control and had the highest density of new bone (MGV = 149.19; SD = 20.96). Reconstructions of microCT images of specimens from group 3 (Fig 2c) showed significant osteoconduction of the defect by the graft. Reconstruction slices of specimens from group 4 showed extensive osteoconduction and healing of the critical-size defect. One-way ANOVA was used to evaluate the differences in bone fill among the four treatment groups. Bone formation differed significantly across the four groups (F[3,20] = 14.13, P < .0001). The Tukey multiple comparison test to compare the four groups indicated that there was no significant difference between groups 1 and 2 (mean difference, 23.99; 95% confidence interval [CI] –11.79, 59.77), but there were significant differences (P < .05) between groups 1 and 3 (mean difference, 42.14; 95% CI 10.14, 74.14) and between groups 1 and 4 (mean difference, 67.77; The International Journal of Oral & Maxillofacial Implants e125
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De Kok et al
Fig 3 Histologic measurements of percentage of bone fill of the critical-size defect at 28 days in the different groups.
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Fig 4a Histologic image of Fig 4b Histologic image of col- Fig 4c Histologic image of empty/negative control (group 1) lagen matrix + saline (group 2) at matrix + MSCs (group 3) at 4 at 4 weeks (magnification ×4). 4 weeks (magnification ×4). weeks (magnification ×4).
95% CI 37.83, 97.70). There were no statistically significant differences between groups 2 and 3 (mean difference, 18.15; 95% CI –17.63, 53.92) or between groups 3 and 4 (mean difference, 25.63; 95% CI –4.304, 55.56), but groups 2 and 4 (mean difference = 43.78; 95% CI 9.834, 77.72) were significantly different.
Histomorphometry The quantitative histomorphometric results confirmed the radiographic assessments. After 28 days of healing, measurements were calculated by percent of bone fill of the critical-size defect (Fig 3). Group 1 showed the least amount of bone formation (mean, 9.25%; SD = 10.82%), followed by group 2 (19.07%; SD = 17.38%). Bridging osteogenesis was noted in the healing of groups 3 and 4 sites. Within the treated defects of these groups, new bone formation with rich vasculature and abundant osteoblasts producing osteoid were revealed. Focal lamellar architecture of the new bone was seen (Fig 4). The results showed that mean bone fill in group 3 (44.21%; SD = 3.93) was not significantly different from that in group 4 (60.06%; SD = 15.08).
Fig 4d Histologic image of matrix + rhBMP-2 (group 4) at 4 weeks (magnification ×4).
A one-way ANOVA was used to compare the differences in bone fill percentage in the four treatment groups; significant differences were seen among all groups (F[3,27] = 22.08; P < .001). The Tukey multiple comparison test indicated that there was no significant difference between groups 1 and 2 (mean difference, –9.821; 95% CI –32.53, 12.89), but there were statistically significant differences (P < .05) between groups 1 and 3 (mean difference, –34.96; 95% CI –55.27 to –14.65) and between groups 1 and 4 (mean difference, 50.81; 95% CI –69.81, –31.81). While there was general agreement between the findings of microCT and histomorphometry, the microCT results revealed differences between groups 2 and 3 (mean difference, –25.14; 95% CI –47.85, –2.431) and between groups 2 and 4 (mean difference, –40.99; 95% CI –62.53, –19.45). Comparisons between groups 3 and 4 were not statistically significant at P < .05 (mean difference, –15.85; 95% CI –34.85 to 3.148). Masson trichrome staining was applied to detect osteoid tissue (Fig 5). Osteoid formation was observed in all the animals from groups 3 and 4, but it was observed only occasionally in groups 1 and 2.
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Fig 5 Histomorphometric assessment of cell-based bone repair from group 4 animal reveals the edge of the defect on the lower left corner and the new bone within the defect to the top right corner. The original bone shows a lamellar pattern of ossification, while the new bone within the defect appears woven (Masson trichrome).
DISCUSSION The rat calvaria critical-size defect model is a commonly used and convenient wasy to screen and evaluate bone regeneration materials. The rat cranium is a suitable site for grafting studies because the graft is protected from relative motion, which eliminates the need for fixation.35 This orthotopic model differs from the ectopic model of osteoinduction per se, but it represents an endosseous situation that is typical of many clinical situations. A type I collagen (DuraGen) was used as a scaffold for both MSCs and rhBMP-2. The scaffolds were prepared from a commercially available material as 8-mm disks approximately 1.0 mm thick. This collagen matrix was shown to be successful in accelerating dural healing and containing cerebrospinal fluid in cases studied by Narotam and colleagues in 2004.36 Its biocompatibility is further suggested by its support of the survival of cerebral cortical neurons in vitro.37 Danish et al compared the performance and complications of DuraGen and AlloDerm for duraplasty and concluded that both materials could be safe alternatives.38 Preliminary experiments using vital dye staining of MSCs showed that DuraGen was readily loaded with human MSCs applied over a 2- to 4-hour period from densely populated culture media (5 × 106 cells/100 µL) (data not shown). It was assumed that the rhBMP-2 was adherent to the engrafted DuraGen material using the 15-minute incubation procedure required for another type I collagen scaffold. Several studies have observed the binding capacity of demineralized bone matrix to osteoinductive growth factors of the TGF family such as BMPs.39 It is not surprising that the collagen scaffold alone failed to promote remarkable bone repair. Neither did it result in inflammatory reactions. The selected scaffold may be a suitable scaffold for osteoinductive agents,
but it was not capable of promoting critical-size defect repair alone. In contrast, Sweeney and colleagues found that animals treated with type I collagen gels alone showed bone repair of 92.5% at 20 weeks.40 Saadeh et al also demonstrated the ability of type I collagen to heal critical-sized bony defects in the rat mandible. He suggested that type I collagen would make a suitable carrier matrix for improved approaches to bone tissue engineering.41 In the present study, a much shorter 4-week healing period was evaluated. Further, these defects were large (8.9 mm) and were performed in adult animals. The present results suggest that the use of type I collagen scaffold for MSCbased bone repair may be effective in promoting early repair of large defects. Here, rhBMP-2 was utilized as a positive control for bone repair in the critical-size defect. In this case, the same type I collagen scaffold was utilized for both rhBMP-2 and MSCs. Other investigations have clearly demonstrated that rhBMP-2 is capable of affecting bone repair of critical-size calvaria defects. The 5-µg/mL solution used for adsorption of rhBMP-2 onto the collagen scaffold does not represent a remarkably high concentration of morphogen, in comparison to many other investigations. Regardless, the present results with this type I collagen scaffold confirm earlier reports of Chen and collaborators, who tested a collagen-based BMP-2–targeting bone repair system in a rabbit mandibular critical-size defect model and concluded that the improved bone formation could be attributed to the retention of BMP-2 by collagen fibrils.42 The present results further support the exploration of collagen as a basis for rhBMP-2 scaffolds. The engrafted MSCs successfully repaired the calvarial defects without additional osteoinductive agents. It is inferred that the type I scaffold enabled engraftment of sufficient cells and maintained their viability The International Journal of Oral & Maxillofacial Implants e127
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De Kok et al
and osteoinductive nature for long enough to engender defect repair. Others have suggested that this type of scaffold supports the differentiation of MSCs into osteoblasts.43 In an in vitro study, a compatible commercial collagen scaffold possessed the ability to support rat MSC osteogenic differentiation as indicated by alkaline phosphatase, osteopontin, and osteocalcin expression.44 In this preliminary study, little effort was made to optimize cell attachment or cell density to achieve bone repair. It is notable that there was great consistency in the bone repair that occurred within 28 days for the cell-engrafted samples. MSC adhesion to type I collagen is well characterized,45 and, while the efficacy of loading these scaffolds has not yet been examined in detail, the results indicate that the scaffold supported delivery of the MSCs to the surgical wounds to evoke direct or indirect effects to encourage bone repair. It is further inferred from the osteogenesis result that the type I collagen scaffold provided a suitable adhesive substrate for MSC differentiation along the osteoblastic pathway. Cell culture studies confirm that type I collagen adhesion of MSCs supports osteoinduction.46 Further experimental application of this collagen scaffold may permit assessment of the minimal cell density/dose required for defined volumetric repair of bone defects. Radiographic assessment of bone formation and three-dimensional reconstruction allowed the authors to visualize the cranial defect from the axial view, which was not possible in histologic sections. A further advantage of radiomorphometry is that it can be performed rapidly after specimen retrieval and can serve as a screening assay. Tabletop systems can inherently induce a discrepancy in the observed MGVs and local bone density, which has been related to the effect of beam hardening. However, this error is assumed to be similar and mutually neutralized for all specimens, since they were comparably of the same size and composition.47 Based on the present data, the radiomorphometric data correlate well with the histomorphometric data. Radiomorphometry was less sensitive than histomorphometry, as indicated by the lack of significant difference seen between groups 2 and 3 in the microCT data. Many authors have validated the use of microCT as a tool for three-dimensional assessment of bone structure for preclinical trials, yet they recommend the use of histology for more conclusive results47,48 and still consider histomorphometry the “gold standard.” The use of microCT for rapid evaluation of bone repair using radiolucent scaffolds such as collagen-based hydrogels is supported by the present data. Resorption of the collagen scaffold prior to completion of bone formation may lead to incomplete bone
fill. Although this may not be a problem with rat MSCs because of their ability to quickly regenerate bone, this could be a potential problem to consider when using such materials in clinical studies with adult human MSCs.
CONCLUSIONS In a rat calvaria critical-size defect model, rat mesenchymal stem cells implanted in a type I collagen matrix promoted bone repair comparable to that induced by recombinant human bone morphogenetic protein 2 in the same matrix material. The use of type I collagen scaffolds for engraftment of mesenchymal stem cells to repair orthotopic defects may be considered in further translational investigations and in clinical practice.
ACKNOWLEDGMENTS This research project was supported by the 2006 Tylman Research Grant from the American Academy of Fixed Prosthodontics. The authors thank Ms Sara Valencia, dental student, and Dr Gustavo Mendonca, Clinical Assistant Professor, Department of Prostho dontics, University of North Carolina School of Dentistry, for their immense help in working with the stem cells and statistical analysis. The authors reported no conflicts of interest related to this study.e
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