Although bone repair is a relatively efficient

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1 EXPERIMENTAL Progenitor Cell Mobilization Enhances Bone Healing by Means of Improved Neovascularization and Osteogenesis Xiao Xia Wang, M.D., D.D.S. Robert J. Allen, Jr., M.D. John Paul Tutela, M.D. Alexander Sailon, M.D. Alexander C. Allori, M.D., M.P.H. Edward H. Davidson, M.A., M.B.B.S. Gina K. Paek, M.D. Pierre B. Saadeh, M.D. Joseph G. McCarthy, M.D. Stephen M. Warren, M.D. New York, N.Y.; and Beijing, People s Republic of China Background: Although bone repair is a relatively efficient process, a significant portion of patients fail to heal their fractures. Because adequate blood supply is essential to osteogenesis, the authors hypothesize that augmenting neovascularization by increasing the number of circulating progenitor cells will improve bony healing. Methods: Bilateral full-thickness defects were created in the parietal bones of C57 wild-type mice. Intraperitoneal AMD3100 (n 33) or sterile saline (n 33) was administered daily beginning on postoperative day 3 and continuing through day 18. Circulating progenitor cell number was quantified by fluorescence-activated cell sorting. Bone regeneration was assessed with micro computed tomography. Immunofluorescent CD31 and osteocalcin staining was performed to assess for vascularity and osteoblast density. Results: AMD3100 treatment increased circulating progenitor cell levels and significantly improved bone regeneration. Calvarial defects of AMD3100- treated mice demonstrated increased vascularity and osteoblast density. Conclusions: Improved bone regeneration in this model was associated with elevated circulating progenitor cell number and subsequently improved neovascularization and osteogenesis. These findings highlight the importance of circulating progenitor cells in bone healing and may provide a novel therapy for bone regeneration. (Plast. Reconstr. Surg. 128: 395, 2011.) From the Institute of Reconstructive Plastic Surgery, New York University Langone Medical Center, and the Department of Oral and Maxillofacial Surgery, Peking University School and Hospital of Stomatology. Received for publication August 6, 2010; accepted February 8, Presented in part at the Tissue Engineering and Regenerative Medicine International Society North America 2008 Annual Conference and Exposition, in San Diego, California, December 7 through 10, 2008, and at the 54th Annual Meeting of the Plastic Surgery Research Council, in Pittsburgh, Pennsylvania, May 27 through 30, Copyright 2011 by the American Society of Plastic Surgeons DOI: /PRS.0b013e31821e6e10 Although bone repair is a relatively efficient process, fracture healing in healthy humans still takes 8 to 12 weeks. Furthermore, in certain types of injury (e.g., scaphoid fracture after a fall on an outstretched hand, or comminuted tibial fracture after a motor vehicle accident), a substantial proportion of bone defects do not heal. 1,2 Current therapeutic options for bone defect reconstruction include the use of autogenous and allogeneic grafts, demineralized bone matrices, synthetic materials, semisynthetic scaffolds, various alloplastic materials, and growth hormones. Despite their clinical success, limitations remain: donor-site morbidity, an obligatory graft resorption phase, contour irregularities, insufficient autogenous resources, disease transmission, graft-versus-host disease, immunosuppression, structural failure, stress shielding, and infection of foreign material. 3 9 To avoid these complications and improve the bone healing process, this study investigates a new method of endogenous vascularized bone engineering. In the bone healing process, an adequate blood supply is critical for successful bone regeneration. Recent studies have found that angiogenesis is required for bone formation during the processes of distraction osteogenesis and fracture healing Lee et al. showed that signals from the bone regeneration site can mobilize bone marrow endothelial progenitor cells into the peripheral circulation and recruit these endothelial progenitor cells to the ischemic fracture site, where they Disclosure: The authors received Plerixafor for experimental use through a nonfinancial relationship with Genzyme. They have no financial interests to declare

2 Plastic and Reconstructive Surgery August 2011 contribute to neovascularization and new bone formation. 14 Similarly, a study by Laing et al. found that endothelial progenitor cell levels in the peripheral blood and bone marrow of humans and mice, respectively, were increased 2 to 3 days after fracture. Their findings suggest that a systemic vasculogenic response is initiated in response to musculoskeletal trauma. 15,16 Matsumoto et al. showed that local or intravenous delivery of granulocyte colony-stimulating factor stimulated CD34 cells (endothelial progenitor cell enriched cells) increases vasculogenesis and osteogenesis and improves fracture healing. 12 Their study also demonstrated that transplanted CD34 cells can differentiate into not only endothelial cells but also osteoblastic cells at the injury site. 12,17 19 Although most of the previously mentioned data have been collected in long bone fracture models, endothelial progenitor cells have been shown to be mobilized following various types of ischemic injuries (e.g., bone and soft tissue), and appropriate vasculogenesis is important for bony healing, regardless of the type of bone (e.g., trabecular or cortical bone). Moreover, endothelial progenitor cells are capable of differentiating into osteoblastic cells. Because vasculogenesis is essential for osteogenesis, and adoptive cellular transfer of vasculogenic CD34 cells enhances fracture healing, we hypothesize that bone healing can be accelerated by increasing the number of circulating progenitor cells with the CXCR4 antagonist AMD3100 (Genzyme, Cambridge, Mass.). AMD3100 is a hematopoietic stem cell mobilizer currently used to prepare blood for autologous stem cell transplantation in patients with non-hodgkin s lymphoma or multiple myeloma. It is a CXCR4 chemokine receptor antagonist that augments the mobilization of endothelial progenitor cells from the bone marrow to the ischemic site of injury. MATERIALS AND METHODS Animals All experimental procedures were approved by the New York University Medical Center Institutional Animal Care and Use Committee (no ). Ten to 12-week-old male C57BLKS/J (no. 662) wild-type mice were purchased from the Jackson Laboratories (Bar Harbor, Me.). Animals were housed in a light- and temperature-controlled environment, and food and water were given ad libitum. Calvarial Defect Model All animals (n 60) were anesthetized with an intraperitoneal injection of a standard anesthesia cocktail consisting of ketamine (100 mg/kg), xylazine (20 mg/kg), and acepromazine (3 mg/kg). After appropriate sedation was obtained, the scalp was shaved and prepared aseptically with povidone-iodine and 70% ethanol and allowed to dry. A no. 15 scalpel blade was used to open the scalp at the midline, exposing the underlying calvaria. Bilateral calvarial defects were made in a non suture-associated area of the parietal bone using a 3-mm-diameter round dental burr under constant saline irrigation, and extreme care was taken to prevent injury to the underlying dura mater (Fig. 1, right). After removing the 3-mm-diameter portion of calvaria, the overlying soft-tissue wound was Fig. 1. Mouse calvarial defect model. (Right) Bilateral 3-mm calvarial defects were created in the parietal bones of wild-type mice using a 3-mm-diameter round dental burr, and the underlying dura mater was kept intact. (Left) A three-dimensional micro computed tomographic image of a calvarial defect immediately following wounding. 396

3 Volume 128, Number 2 AMD3100 and Calvarial Defect Healing closed immediately with interrupted 6-0 nylon sutures (Ethicon, Inc., Somerville, N.J.). AMD3100 Application All animals (n 60) were divided randomly into two groups. One group (n 30) of mice (A ) was injected intraperitoneally with AMD3100 (10 mg/kg) beginning on postoperative day 3 and continuing daily for 15 days. A second group (n 30) of mice (control) was injected with an equivalent volume of normal saline. Micro Computed Tomographic Evaluation of Calvarial Defect Healing Animals were euthanized at postoperative weeks 1, 2, 4, 8, and 12 (n 6 per time point per group) and calvariae harvested at the 4-, 8-, and 12-week time points were analyzed for defect closure. Specimens were fixed in 4% paraformaldehyde for 48 hours, positioned vertically, and scanned horizontally at 16- m resolution using a SCANCO Micro-CT 40 device (SCANCO Medical AG, Brüttisellen, Switzerland). Serial axial images of the calvarial defects were used to generate three-dimensional reconstructions. The calvarial defect area was calculated using Adobe Photoshop 7.0 (Adobe Systems, Inc., San Jose, Calif.). The defect area was calibrated against the internal calibrator of the three-dimensional images to correct for any subtle magnification differences (Fig. 1, left). Percentage defect healing was defined as follows: (1 [current defect area]/[original defect area]) 100 percent. Circulating Progenitor Cell Measurement To investigate the effects of AMD3100 on circulating progenitor cell number, peripheral blood was harvested from both saline-treated and A mice at postoperative days 7 and 14 (n 6 per time point per group). To obtain a baseline level of circulating progenitor cells, an additional six animals (not included in the n 60 experimental population) that had not undergone surgical intervention were used and considered to be day-0 samples. The peripheral blood was harvested by means of intracardiac puncture using a 30-gauge needle and a 1-ml heparinized syringe. Peripheral blood derived mononuclear cells were separated by density gradient centrifugation with Histopaque 1083 (Sigma-Aldrich, St. Louis, Mo.). Peripheral blood mononuclear cells were then stained with rat anti-mouse antibodies to further quantify the progenitor cell population [fluorescein isothiocyanate conjugated Sca-1, phycoerythrin-conjugated vascular endothelial growth factor (VEGF) receptor 2, allophycocyanin-conjugated c-kit, biotin-conjugated lineage depletion cocktail, and streptavidin-phycoerythrin conjugated-cy7] (BD Biosciences, San Jose, Calif.). This triple staining marks Sca-1 /c-kit /lineage stem cell enriched populations (e.g., stromal cell derived factor-1 and CXCR4-mediated stem cells). Samples were collected using a BD FACSCaliber flow cytometer (BD Biosciences), and analysis was performed with FlowJo 8.0 software (Tree Star, Inc., Ashland, Ore.) to quantify lineage /Sca-1, lineage /c-kit, lineage /Sca-1 /c-kit, and lineage /Sca-1 /flk populations. Tissue Harvest and Processing Animals were euthanized by carbon dioxide narcosis and cervical dislocation at postoperative weeks 1, 2, 4, 8, and 12 (n 6 per group per time point). After being euthanized, the mice were decapitated; the skin, other underlying superficial soft tissues, and mandible were removed; and the calvariae were harvested. Half of the samples harvested at 1, 2, and 4 weeks (n 3 per group per time point) were immediately embedded in optimal cutting temperature compound (Tissue-Tek OCT; Sakura Finetek, Torrance, Calif.), snap-frozen in liquid nitrogen, and stored at 80 C for CD31 immunofluorescent staining (see below). The rest of the samples harvested at 1, 2, and 4 weeks (n 3 per group per time point) were fixed in a 4% paraformaldehyde/10% sucrose solution at 4 C for 48 hours and decalcified in Immunocal formic acid decalcifier solution (Decal Chemical Corp., Tallman, N.Y.) at 4 C for 5 days. After decalcification, the calvaria were embedded in Tissue-Tek optimal cutting temperature compound and stored as mentioned above for immunofluorescent osteocalcin staining (see below). Samples harvested at 8 and 12 weeks (n 6/group) were fixed in 10% formalin, decalcified, and embedded in paraffin for permanent sections and stained with hematoxylin and eosin and Gomori trichrome. Immunofluorescent Staining and Histology To analyze the tissue samples for vascularity and new bone formation, tissue samples were stained with CD31 or osteocalcin, respectively. For all the staining performed on frozen sections, 6- m serial sections were made using a Leica CM1900 cryostat (Leica Microsystems, Bannockburn, Ill.). Sections to be stained with CD31 were fixed with acetone and stained overnight with monoclonal rat anti-mouse CD31 primary anti- 397

4 Plastic and Reconstructive Surgery August 2011 Fig. 2. AMD3100 improves calvarial defect healing. (Above) Micro-computed tomographic images of defects at weeks 4, 8, and 12 demonstrate that AMD3100-treated mice had improved 398

5 Volume 128, Number 2 AMD3100 and Calvarial Defect Healing body (1:100 dilution; BD Biosciences) followed by Alexa Fluor 594 goat anti-rat immunoglobulin G secondary antibody staining (1:100 dilution; Molecular Probes, Eugene, Ore.). Slides were mounted with Vectashield with 4=,6-diamidino-2- phenylindole (Vector Laboratories, Burlingame, Calif.). Analysis was performed using an Olympus BX51 epifluorescent microscope (Olympus America, Inc., Center Valley, Pa.). Photoshop CS3 (Adobe) was used to segment and quantify the amount of positive CD31 staining. The vascular density of the bony defect was calculated by quantifying the number of CD31 staining vessels (red) per megapixel ( pixels square area) of defect area. Pictures of three random high-power fields were taken of each sample and analysis was performed by a blinded interpreter. Samples to be analyzed for new bone formation were sectioned onto golden glass slides (Fisher Scientific, Pittsburgh, Pa.) and air-dried at room temperature for 30 minutes. After washing, the sections were flooded with 100 mm glycine in Tris buffer (ph 7.4) for 20 minutes at room temperature to eliminate the formalin autofluorescence background. The sections were then treated with 0.1% trypsin in phosphate-buffered saline for 30 minutes at 37 C, blocked with normal goat serum (GIBCO from Invitrogen, Carlsbad, Calif.) for 1 hour, and incubated overnight with polyclonal rabbit anti-mouse osteocalcin primary antibody (1:200 dilution) at 4 C. Subsequently, the sections were stained with Alexa-Fluor 488 conjugated goat anti-mouse immunoglobulin G secondary antibody (1:100 dilution; Molecular Probes) at 4 C for 2 hours and mounted in mounting media with 4=,6-diamidino-2-phenylindole (Vectashield; Vector Laboratories). Osteocalcinpositive staining (green) was quantified as mentioned above. Samples obtained at weeks 4, 8, and 12 were also stained with hematoxylin and eosin and Gomori trichrome. Photographs were taken at 100 magnification. Statistical Analysis All statistical data are expressed as mean SEM. Data were analyzed by t test using SigmaStat Fig. 2.(Continued) calvarialdefecthealing(right) comparedwith sham-treated controls (left). (Below) AMD3100-treated animals had greater calvarial defect healing at postoperative weeks 8(p 0.017) and 12 compared with controls (p 0.001). No significant difference was found between AMD3100-treated mice and controls at week 4 (p 0.31). software (SPSS, Inc., Chicago, Ill.). Values of p 0.05 were considered significant. RESULTS Critical-Sized Defect Model After creation of the 3-mm calvarial defect, animals were allowed to heal for 12 weeks. Although Cowan et al. stated that a cranial bone defect should not heal more than 10 percent, 3-mm bony defects have been shown to remain open up to 12 weeks postoperatively. 20 The 3-mm bony defects were found to be critical sized, as those of control mice had less than a 4 percent difference in closure between weeks 4 ( percent) and 12 ( percent). Based on previously published work, these defects were considered critical size. 21 AMD3100 Improves Calvarial Defect Healing Because progenitor cells are known to be involved in the repair of both soft-tissue and bone defects and AMD3100 increases the number of circulating progenitor cells, we next investigated the effects of AMD3100 treatment on the healing of calvarial defects in wild-type mice. Calvarial defects in mice treated with AMD3100 had improved healing at postoperative weeks 4, 8, and 12 compared with controls ( percent versus percent, p 0.31; percent versus percent, p 0.017; and percent versus percent, p 0.001, respectively) (Fig. 2). Circulating Progenitor Cells Because progenitor cells are known to be released from the bone marrow in response to an ischemic insult, we next investigated the effects of our calvarial bone defect on the number of circulating progenitor cells in our control wild-type mice. The baseline number of circulating progenitor cells (lineage /Sca-1 /c-kit ) as measured in noninjured wild-type mice on day 0 is percent. After the creation of a calvarial defect, this number increases over 2-fold to percent at day 7 (p 0.05), but returns to percent at day 14 after the injury, which is not significantly different from the baseline circulating progenitor cell levels (p 0.76). With AMD3100 treatment, circulating progenitor cell numbers increased to supraphysiologic levels. After five daily doses of AMD3100 (postwounding day 7), the circulating progenitor cell number peaked to percent, a fold increase compared with the sham-treated 399

6 Plastic and Reconstructive Surgery August 2011 control group at the same time point (p 0.003). The AMD3100-induced peak in progenitor cell number was 3.9-fold greater than baseline circulating progenitor cells. After 12 treatments of AMD3100 (postwounding day 14), the circulating progenitor cell number was percent, which was a 2.49-fold increase compared with the control group at the same time point (p 0.014) (Fig. 3). Similar findings were noted when investigating other subpopulations of circulating progenitor cells. For example, the number of circulating lineage /Sca-1 /c-kit cells in wild-type mice increased following a calvarial defect (baseline, percent; day 7, percent; and day 14, percent). With the addition of daily AMD3100 injections, this number increased significantly compared with baseline at days 7 and 14 ( percent and percent, respectively) and when compared with sham-treated controls (postoperative day 7, 38.1 percent, p 0.05; postoperative day 14, percent, p 0.01) (Fig. 4). AMD3100-Improved Bony Healing Is Associated with Increased Neovascularization and Osteogenesis Because AMD3100 treatment significantly improves calvarial defect healing and increases the number of circulating progenitor cells in our model, we investigated its effects on the amount of blood vessel and new bone formation. Image-analysis segmentation of CD31-stained sections was performed to quantify the vascular density, and osteocalcin-stained sections were used to quantify the osteoblast density. The results demonstrated that AMD3100 treatment substantially increased both the vascular density and osteoblast density at postoperative weeks 1, 2, and 4 compared with sham-treated control groups. The vascular density increased from percent to percent at week 1 (p 0.01), from percent to percent at week 2 (p 0.01), and from percent to percent at week 4 (p 0.01) (Fig. 5). The osteoblast density increased from percent to percent at week 1 (p 0.01), from percent to percent at week 2(p 0.01), and from percent to percent at week 4 (p 0.01) (Fig. 6). In addition, Gomori trichrome staining showed that with AMD3100 administration, the regenerating tissue at the bony defect margin was much thicker than in the control group and that there was an increase in new bone formation and more newly formed capillaries in the regeneration tissue (Fig. 7). DISCUSSION During healing, neovascularization is a crucial initiator of bone formation and remodeling. Thus, angiogenesis/vasculogenesis and osteogenesis are critically linked. This interplay has been demonstrated in numerous studies For example, Fang et al. showed that administration of the angiogenic inhibitor TNP-470 not only significantly disrupted local angiogenesis but also prevented normal osteogenesis, resulting in a fibrous union in a rat distraction osteogenesis model. 10 Subsequent studies revealed similar findings, as TNP-470 was also shown to prevent femoral fracture healing in rats. 11 Inhibition of angiogenesis by soluble Flt-l (VEGF antagonist) suppressed not only angiogenesis but also osteogenesis in the rat femur fracture model. 12 Finally, Carvalho et al. showed that angiogenesis-associated genes remained elevated during the time course of distraction osteogenesis, suggesting a relationship between new blood vessel formation and osteogenesis. 13 In 1999, the process of postnatal vasculogenesis was described by Asahara et al., who found that bone marrow derived endothelial progenitor cells contribute to as much as 35 percent of new blood vessel formation. 22,23 Since that time, a flurry of studies investigating the interplay of this process and numerous pathologic conditions have been reported. For example, diabetic wound healing is impaired, in part, because of inadequate neovascularization. Lin et al. and Tanaka et al. demonstrated that application of bone marrow derived progenitor cells to wounds on the dorsum of diabetic mice significantly improved soft-tissue wound healing. 24,25 In addition to their role in wound healing, a number of studies have demonstrated that bone marrow derived progenitor cells play a role in the new blood vessel formation that is critical to bone formation. For example, human endothelial progenitor cells xenotransplanted into a rat were found to contribute to new blood vessel formation in a distraction model. 26 These authors also demonstrated a significant improvement in wound healing with this therapy. Further studies by Lee et al. using both fracture and distraction osteogenesis models found circulating endothelial progenitor cell and VEGF levels to increase following the creation of a bone defect. In addition, endothelial progenitor cell mobilizing cytokines (VEGF, stem cell factor, stromal cell derived factor-1, monocyte chemoattractant protein-1) and 400

7 Volume 128, Number 2 AMD3100 and Calvarial Defect Healing Fig. 3. Circulating progenitor cells. After the creation of our bony defect, the levels of several subpopulations of endothelial progenitor cells increased over time. Compared with baseline, the number of lineage / Sca-1 /c-kit cells in sham-treated control mice increased over 2-fold at postoperative day 7 ( percent) and then decreased back toward baseline values ( percent) at postoperative day 14 ( percent). In AMD3100-treated animals, the increase in circulating endothelial progenitor cells was even more dramatic compared with baseline at postoperative days 7 ( percent) and 14 ( percent). Furthermore, compared with shamtreated control animals, the AMD3100-treated animals showed a 1.87-fold increase at postoperative day 7 (p 0.003) and a 2.49-fold increase at postoperative day 14 (p 0.014). Fig.4. AMD3100treatmentalsoincreasesthenumberoflineage /Sca- 1 /Flk circulating endothelial progenitor cells. Similar to what was found in the lineage /Sca-1 /c-kit endothelial progenitor cell population, analysis of another subpopulation of circulating endothelial progenitor cells that were lineage /Sca-1 /Flk showed an increase in both sham-treated controls (baseline, percent; postoperative day 7, percent; and postoperative day 14, percent) and AMD3100-treated animals (postoperative day 7, percent; and postoperative day 14, percent) as compared with baseline. In addition, comparison of the AMD3100 group to the saline-treated controls showed a 38.1 percent increase at postoperative day 7 (p 0.05) and a greater than 2-fold increase at postoperative day 14 (p 0.01). 401

8 Plastic and Reconstructive Surgery August 2011 Fig. 5. Vascular density of regeneration tissue at the bony defect. (Above and center) More newly formed capillaries shown as red CD31 staining were found in the much thicker regeneration tissue at and around the bony defect margin in the AMD3100-treated group than in the saline control group. (Below) AMD3100 treatment significantly increased the vascular density at postoperative weeks 1, 2, and 4 compared with controls ( percent versus percent at week 1, p 0.01; percent versus percent at week 2, p 0.01; and percent versus percent at week 4, p 0.01). adhesion molecules (specific adhesion molecules) involved in endothelial progenitor cell homing were all expressed at the fracture callus. 14 Similarly, a recent study by Laing et al. found that patients with musculoskeletal trauma and mice with femur fractures had a marked increase in circulating endothelial progenitor cells 2 to 3 days after trauma. The circulating endothelial progenitor cell levels gradually decreased to baseline level 1 week after trauma. 15,16 Interestingly, endothelial progenitor cells may also promote new bone formation through an al- 402

9 Volume 128, Number 2 AMD3100 and Calvarial Defect Healing Fig. 6. Osteoblast density of regeneration tissue at bony defect. (Above and center) More osteoblasts were found in the newly formed bone, and the regeneration tissue was much thicker on the frozen slides stained with fluorescence osteocalcin in the AMD3100-treated group than in the saline control group. (Below) With AMD3100 administration, the osteoblast density of the regeneration tissue was significantly increased at postoperative week 1, 2, and 4 compared with sham-treated control groups ( percent versus percent at week 1, p 0.01; percent versus percent at week 2, p 0.01; and percent to percent at week 4, p 0.01). ternative, nonvasculogenic mechanism; recent reports have surfaced suggesting that endothelial progenitor cells have an osteogenic potential. Chen et al. found that the hematopoietic progenitor cell marker CD34 present on human endothelial progenitor cells is also present on cells capable of osteoblastic activity in culture. 27 Approximately 20 percent of CD34 cells isolated from human volunteers also showed osteocalcin expression, a marker for osteogenic differentiation. 21 Other studies have 403

10 Plastic and Reconstructive Surgery August 2011 Fig. 7. Gomori trichrome stain of the calvarial bony defect. Arrows indicate the original bony defect margin and thicker regeneration tissue, and more capillaries are observed stemming from the defect margin and filling the defect in the AMD3100-treated group as compared with the control group at postoperative week 4. found similar results in mice; murine endothelial progenitor cells have been shown capable of differentiation in either a vasculogenic or osteogenic direction. 28,29 AMD3100 (plerixafor), an antagonist of the CXCR4 chemokine receptor, can quickly mobilize progenitor cells from bone marrow into circulation by partially inhibiting the stromal cell derived factor-1/cxcr4 interaction. 30 In our study, we found that AMD3100 administration increased circulating progenitor cell levels in a mouse calvarial defect model for up to 2 weeks after injury. An increase in circulating progenitor cells was associated with increased new blood vessel and bone formation in the regenerating tissue. Our results demonstrate that AMD3100 has the potential to enhance bone healing through increasing neovascularization and osteogenesis. Whether the mobilized progenitor cells promoted new bone formation through vasculogenic and primary osteogenic mechanisms remains to be determined. Additional experiments using mice with green fluorescent protein labeled bone marrow could give extra insight into the direct function of bone marrow derived cells in this model. CONCLUSIONS AMD3100 improves bone regeneration by means of increased neovascularization and osteogenesis. This finding provides further insight into the interplay of neovascularization and osteogenesis and may provide a novel therapy for endogenous bone regeneration. Stephen M. Warren, M.D. Institute of Reconstructive Plastic Surgery New York University Medical Center 560 First Avenue, TCH-169 New York, N.Y stephen.warren.md@gmail.com REFERENCES 1. Marsh D. Concepts of fracture union, delayed union, and nonunion. Clin Orthop Relat Res. 1998;355(Suppl):S22 S Rodriguez-Merchan EC, Forriol F. Nonunion: General principles and experimental data. Clin Orthop Relat Res. 2004; 419: Flannery T, McConnell RS. Cranioplasty: Why throw the bone flap out? Br J Neurosurg. 2001;15: Hidalgo DA. Fibula free flap: A new method of mandible reconstruction. Plast Reconstr Surg. 1989;84: Clokie CM, Moghadam H, Jackson MT, Sandor GK. Closure of critical sized defects with allogenic and alloplastic bone substitutes. J Craniofac Surg. 2002;13: ; discussion Einhorn TA, Lane JM, Burstein AH, Kopman CR, Vigorita VJ. The healing of segmental bone defects induced by demineralized bone matrix: A radiographic and biomechanical study. J Bone Joint Surg Am. 1984;66: Kretlow JD, Mikos AG. Review: Mineralization of synthetic polymer scaffolds for bone tissue engineering. Tissue Eng. 2007;13: Marchac D, Greensmith A. Long-term experience with methylmethacrylate cranioplasty in craniofacial surgery. J Plast Reconstr Aesthet Surg. 2008;61: ; discussion Lind M. Growth factors: Possible new clinical tools. A review. Acta Orthop Scand. 1996;67: Fang TD, Salim A, Xia W, et al. Angiogenesis is required for successful bone formation during distraction osteogenesis. J Bone Miner Res. 2005;20: Hausman MR, Schaffler MB, Majeska RJ. Prevention of fracture healing in rats by an inhibitor of angiogenesis. Bone 2001;29: Matsumoto T, Kawamoto A, Kuroda R, et al. Therapeutic potential of vasculogenesis and osteogenesis promoted by 404

11 Volume 128, Number 2 AMD3100 and Calvarial Defect Healing peripheral blood CD34-positive cells for functional bone healing. Am J Pathol. 2006;169: Carvalho RS, Einhorn TA, Lehmann W, et al. The role of angiogenesis in a murine tibial model of distraction osteogenesis. Bone 2004;34: Lee DY, Cho TJ, Kim JA, et al. Mobilization of endothelial progenitor cells in fracture healing and distraction osteogenesis. Bone 2008;42: Laing AJ, Dillon JP, Condon E, et al. Mobilization of endothelial precursor cells: Systemic vascular response to musculoskeletal trauma. J Orthop Res. 2007;25: Laing AJ, Dillon JP, Condon ET, et al. A systemic provascular response in bone marrow to musculoskeletal trauma in mice. J Bone Joint Surg Br. 2007;89: Mifune Y, Matsumoto T, Kawamoto A, et al. Local delivery of granulocyte colony stimulating factor-mobilized CD34-positive progenitor cells using bioscaffold for modality of unhealing bone fracture. Stem Cells 2008;26: Matsumoto T, Kuroda R, Mifune Y, et al. Circulating endothelial/skeletal progenitor cells for bone regeneration and healing. Bone 2008;43: Matsumoto T, Mifune Y, Kawamoto A, et al. Fracture induced mobilization and incorporation of bone marrow-derived endothelial progenitor cells for bone healing. J Cell Physiol. 2008;215: Cowan CM, Shi YY, Aalami OO, et al. Adipose-derived adult stromal cells heal critical-size mouse calvarial defects. Nat Biotechnol. 2004;22: Hollinger JO, Kleinschmidt JC. The critical size defect as an experimental model to test bone repair materials. J Craniofac Surg. 1990;1: Asahara T, Murohara T, Sullivan A, et al. Isolation of putative progenitor endothelial cells for angiogenesis. Science 1997; 275: Asahara T, Masuda H, Takahashi T, et al. Bone marrow origin of endothelial progenitor cells responsible for postnatal vasculogenesis in physiological and pathological neovascularization. Circ Res. 1999;85: Lin CD, Allori AC, Macklin JE, et al. Topical lineage-negative progenitor-cell therapy for diabetic wounds. Plast Reconstr Surg. 2008;122: Tanaka R, Wada M, Kwon SM, et al. The effects of flap ischemia on normal and diabetic progenitor cell function. Plast Reconstr Surg. 2008;121: Cetrulo CL Jr, Knox KR, Brown DJ, et al. Stem cells and distraction osteogenesis: Endothelial progenitor cells home to the ischemic generate in activation and consolidation. Plast Reconstr Surg. 2005;116: ; discussion Chen JL, Hunt P, McElvain M, Black T, Kaufman S, Choi ES. Osteoblast precursor cells are found in CD34 cells from human bone marrow. Stem Cells 1997;15: Khosla S, Eghbali-Fatourechi GZ. Circulating cells with osteogenic potential. Ann NY Acad Sci. 2006;1068: Bick T, Rozen N, Dreyfuss E, Soudry M, Lewinson D. Osteogenic differentiation of circulating endothelial progenitor cells. J Bone Miner Res. 2007;22:S Martin C, Bridger GJ, Rankin SM. Structural analogues of AMD3100 mobilise haematopoietic progenitor cells from bone marrow in vivo according to their ability to inhibit CXCL12 binding to CXCR4 in vitro. Br J Haematol. 2006; 134: