Fax: +55 19 3521-6185.
Spontaneous Healing Capacity of Calvarial Bone Defects in mdx Mice
Article first published online: 23 JAN 2012
Copyright © 2012 Wiley Periodicals, Inc.
The Anatomical Record
Volume 295, Issue 4, pages 590–596, April 2012
How to Cite
Nakagaki, W. R. and Camilli, J. A. (2012), Spontaneous Healing Capacity of Calvarial Bone Defects in mdx Mice. Anat Rec, 295: 590–596. doi: 10.1002/ar.22412
- Issue published online: 12 MAR 2012
- Article first published online: 23 JAN 2012
- Manuscript Accepted: 20 DEC 2011
- Manuscript Received: 15 SEP 2011
- FAPESP. Grant Numbers: 2007/07638-0, 2008/57041-2
- CAPES/PROEX, Brazil.
- mdx mice;
- bone healing;
- calvarial defect
The mdx mouse is an experimental model widely used for the study of Duchenne muscular dystrophy, which is characterized by the lack of dystrophin and cycles of muscle degeneration/regeneration. Studies demonstrated elevated levels of growth factors and accelerated skin wound repair in these animals. We therefore raised the hypothesis that the bone repair process might also be altered in these animals. Thus, the objective of this study was to evaluate the spontaneous healing of calvarial defects in mdx mice by histomorphometric analysis. Animals (45 days old) were divided into mdx and control groups. A defect measuring 2 mm in diameter was produced surgically in the right parietal bone of each animal. The animals were sacrificed 15, 30, and 60 days after surgery, and the skulls were processed by routine histological procedures. No difference in the volume of new bone inside the defect was observed between the two groups at any of the three postoperative time points. There was also no difference between the different periods of healing when each group was analyzed separately. The lower quality of femoral and calvarial bone in mdx mice reported in previous studies and the similar bone regeneration rates seen in two groups suggest that the healing capacity of calvarial defects was more expressive in mdx mice than in control animals. An increase in the amount of osteogenic factors released by damaged myofibers may have favored osteogenesis during bone defect healing in mdx mice. Anat Rec, 2012. © 2012 Wiley Periodicals, Inc.
Duchenne muscular dystrophy (DMD) is the most frequent childhood form of muscular dystrophy and is transmitted in an X chromosome-linked recessive manner. The course of the disease is relatively rapid and progressive, with the occurrence of muscle degeneration and consequent intense and profound muscle weakness (Aparicio et al.,2002). This muscle weakness, in turn, leads to thin and demineralized bone and delays the onset of ossification centers (Hsu,1982). As a consequence, a reduction in bone mineral density occurs as the disease advances (Larson and Henderson,2000; McDonald et al.,2002; Bianchi et al.,2003), leading to osteoporosis and bones that are more vulnerable to fractures (Söderpalm et al.,2007). In addition, the administration of glucocorticoids to treat muscle weakness and inflammation exerts long-term side effects on bone metabolism, increasing osteoblastic apoptosis and osteoclastogenesis (Canalis et al.,2004).
The association between muscular dystrophy and bone tissue metabolism has been studied in humans (Larson and Anderson,2000; Vestergaard et al.,2001; Aparicio et al.,2002; McDonald et al.,2002; Bianchi et al.,2003; Biggar et al.,2005; Söderpalm et al.,2007) and mdx (X chromosome-linked muscular dystrophy) mice (Anderson et al.,1993; Montgomery et al.,2005; Nakagaki et al.,2011). As in human DMD, mdx mice carry a genetic mutation that results in the lack of dystrophin, intense muscle inflammation, and cycles of degeneration/regeneration, which become intense and most marked after the 21st day of life (Pastoret and Sebille,1995; Porter et al.,2003b; Briguet at al.,2004; Grounds and Torrisi,2004; Reed and Bloch,2005). However, in contrast to DMD, the muscle fiber regeneration that occurs in mdx mice seems to compensate for the degenerative process, with the observation of less fibrosis and consequent maintenance of muscle function (Yablonka-Reuveni and Anderson,2006; Evans et al.,2009).
Anderson et al. (1993) observed a smaller cortical thickness and area, increased porosity and lower mechanical strength in the tibia of 28-day-old mdx mice and attributed these deficiencies to muscle weakness resulting from intense muscle fiber degeneration. However, in the absence of signs of quadriceps degeneration, Nakagaki et al. (2011) found that the femur of 21-day-old mdx mice exhibited osteopenia, a smaller number of osteoblasts, lower mineral content, and lower mechanical strength when compared with the control group. These findings suggest that the loss of bone quality in mdx mice is not only due to muscle weakness.
Montgomery et al. (2005) observed higher mineral density and mechanical strength of the femur and higher hindlimb muscle mass in mdx mice (4 months old), despite the presence of systemic muscle weakness caused by episodes of muscle degeneration. In addition, the proximal third of the femur was larger and expanded laterally, demonstrating the existence of localized stimuli of bone formation at the osteotendinous junctions during periods of muscle regeneration. According to the authors, the expansion of bone tissue at these sites is an adaptation to accommodate increases in muscle size and mass even in the absence of an increase in muscle contractile strength. Furthermore, bone tissue metabolism in mdx mice is also influenced by the action of osteogenic factors released from damaged myofibers (Montgomery et al.,2005), and the levels of these factors have been shown to be elevated in mdx mice (Ishitobi et al.,2000; Vetrone et al.,2009) and in patients with DMD (D'Amore et al.,1994).
Growth factors, cytokines and chemokines, are essential for the modulation of bone regeneration and the differentiation of muscle cells (Straino et al.,2004). Studies have shown elevated levels of growth factors and chemokines such as fibroblast growth factor (D'Amore et al.,1994), monocyte chemotactic protein-1 (Porter et al.,2003a), and neural growth factor (Toti et al.,2003) in mdx mice. In this respect, Straino et al. (2004) observed an increase in the number of arterioles after hindlimb ischemia and faster skin wound healing in mdx mice when compared with normal animals. The authors identified a link between the lack of dystrophin and neovascularization and suggested that increased vasculogenesis may represent an adaptive response of the organism to pathological levels of muscle degeneration.
The evidence showing a higher amount of osteogenic factors (Ishitobi et al.,2000; Vetrone et al.,2009), elevated levels of growth factors and acceleration of skin wound healing in mdx mice (Straino et al.,2004) support the hypothesis that bone repair might also be altered in these animals. Therefore, the objective of this study was to investigate by histomorphometric analysis the healing capacity of bone defects surgically created in the skull of mdx mice after different periods of spontaneous healing.
MATERIALS AND METHODS
Male C57BL/10ScN (n = 15; control group) and C57BL/10ScSn-Dmdmdx mice (n = 15; mdx group) were submitted to surgery at 45 days of age. Mdx and control animals (n = 5 per time point) were sacrificed 15, 30, and 60 days after surgery with an overdose of the anesthetic, and their skulls were removed and processed for analysis. The animals were obtained from a breeding colony maintained by our institutional animal care (Multidisciplinary Center for Biological Research—CEMIB), and the origin of this colony is the Institute Pasteur (Paris, France, 1991). The experiment was conducted in accordance with the ethical guidelines adopted by the Brazilian College of Animal Experimentation (COBEA) and the study was approved by the Ethics Committee on Animal Experimentation (CEEA) of Unicamp (protocol 1359-1).
Surgical Procedure for the Creation of Calvarial Defects
The animals were anesthetized by intramuscular administration of a 1:1 solution of xylazine hydrochloride (5 mg/kg) and ketamine hydrochloride (100 mg/kg). After shaving the cranial skin and cleansing the area with ethanol and iodine, a longitudinal incision of ∼6 cm was made to expose the surface of the parietal bones. Next, the periosteum was separated and a full-thickness defect measuring 2 mm in diameter was created with a biopsy punch through the right parietal bone. The defect remained empty. The periosteum was repositioned and the skin was sutured. All animals received an analgesic (500 mg/mL sodium dipyrone) in their drinking water (ad libitum) at a dose of 875 mg/kg for the first five postoperative days. This procedure was based on the study of Silva and Camilli (2006).
The samples were fixed in 10% buffered formalin for 72 hr and decalcified in a solution of formic acid, formalin, and sodium citrate for 35 days. Next, the specimens were submitted to routine histological processing and embedded in paraffin. Five histological sections were obtained from each skullcap in the coronal plane. All histological sections were taken near the greatest diameter (middle third) of the defect. Cross-sections (5-μm thick) were stained with hematoxylin–eosin. The histological sections were examined under a Nikon 80i photomicroscope equipped with 10× and 20× objectives and the images were captured with a Nikon DS-Ri1 camera.
The formation of new bone inside the defect was quantified as described by Gosain et al. (2000), using a combination of two methods. In the first method, the ratio between the linear length of new bone and the initial linear length of the defect measured across the central axis of the bone defect was calculated and is reported as percentage. The linear length of new bone corresponded to the sum of all lengths of each portion of bone to be formed inside the defect. In the second method, the percentage of new bone was calculated based on the ratio between the area of new bone and the total area of the initial defect. The morphometric measurements were made on the images of the histological sections using the NIS-Elements Advanced Research 3.0 software.
Two-way analysis of variance for repeated measures, followed by the Tukey test if necessary, was used for statistical analysis. The results are reported as the mean ± standard deviation. A level of significance of 5% was adopted for all tests (P < 0.05).
Analysis of the histological sections showed the absence of bone formation in the center of the defect in the two groups, irrespective of the time of bone healing. New bone was only observed at the borders of the defect. Portions of soft connective tissue were seen in the center of the defect in front of the dura mater, which remained intact (Figs. 1A–F and 2A–F).
No significant difference in the volume of new bone was observed between mdx and control mice at any of the three postoperative time points. In addition, there was no difference in bone formation between the different postoperative time points in either group. Statistical analysis showed no interaction between disease and time of healing (Fig. 3).
Although the percentage of bone formation inside the defect in terms of area and linear length was statistically similar in the two groups, the mean values of bone regeneration were higher in the mdx group than in the control group (Fig. 3).
Since the mdx mouse is a model widely used for the study of DMD, in which a link between the absence of dystrophin and neovascularization and accelerated skin wound healing has been demonstrated (Straino et al.,2004), this study evaluated the spontaneous healing capacity of calvarial bone defects created in mdx mice after different periods of healing.
The complex process of bone defect regeneration requires the coordinated interaction between growth factors, extracellular matrix proteins, osteoblasts, and osteoprogenitor cells (Aalami et al.,2004). In addition to these factors, the periosteum, dura mater, postsurgery time, and age play a role in the healing of calvarial defects in mice (Aalami et al.,2004). According to these authors, spontaneous healing of calvarial bone defects is compromised when the dura mater is damaged during surgery. In this study, the dura mater did not interfere with bone healing since it was preserved in all animals.
Greenwald et al. (2000a) suggested that dura mater cells of young mice have a greater capacity to differentiate into bone than those of adult mice due to increased alkaline phosphatase activity and osteocalcin synthesis. Other studies showed that the dura mater and calvarial osteoblasts obtained from 6-day-old rats had a higher in vitro osteogenic potential than cells isolated from 60-day-old animals (Greenwald et al.,2000b; Cowan et al.,2003). Aalami et al. (2004) observed that young mice (6 days old) presented a significantly higher bone-forming capacity in calvarial defects than adult animals (60 days old) after 8 weeks of healing. Our results agree with these studies. The adult animals studied here presented similar bone regeneration rates at the three postoperative time points (15, 30, and 60), irrespective of the group analyzed.
Alterations of calvarial bone morphology in mdx mice have been reported in a recent study, with the observation of an increased number of osteoclasts (Rufo et al.,2009). Nakagaki et al. (2011) observed osteopenia in the femur of 21-day-old mdx mice in the absence of signs of quadriceps degeneration, suggesting the existence of some intrinsic factor that reduces bone quality and that might be directly or indirectly related to the lack of dystrophin. This reduction in bone quality and the similar bone regeneration rates in the control and mdx groups found in this study suggest that the osteogenic capacity of calvarial cells was more expressive in mdx mice compared with control animals between 45 and 105 days of age. These ages correspond to the time of surgery and sacrifice during the last postoperative period studied. Thus, the similar regeneration rates observed in the two groups indicate a positive result for mdx mice considering the presence of bone tissue abnormalities in these animals resulting from a genetic mutation.
Growth factors, cytokines and chemokines, are essential for the modulation of bone regeneration and muscle cell differentiation (Straino et al.,2004). According to D'Amore et al. (1994), osteogenic factors such as fibroblast growth factor-2 (FGF-2) are elevated in patients with DMD because of their release from damaged myofibers. Since muscle fiber necrosis reaches its peak between 35 and 90 days and >50% of the muscle fibers of mdx mice are in the process of regeneration (Tanabe et al.,1986), it is possible that specific stimuli derived from damaged myofibers have increased vasculogenesis during this period, stimulating bone healing. In addition, studies have shown that the transforming growth factor-β1 (TGF-β1) and osteopontin (OPN) levels are elevated in skeletal muscle and serum in mdx mice (Ishitobi et al.,2000; Vetrone et al.,2009). OPN displays chemoattractant activity for progenitors cells of osteoblasts and for peripheral blood monocytes (Chenu et al.,1994) and TGF-β1 induces migration of bone marrow stromal cells to the bone surface in response to bone resorption (Pang et al.,2010). Thus, there is a high content of prejudicial factors to the muscles, but that may be capable of stimulating bone formation in these animals.
During episodes of degeneration of muscle fibers of mdx mice, there is an increase in the levels of tumor necrosis factor-α (TNF-α) in skeletal muscle (Grounds et al.,2008) and serum (Pan et al.,2008). Thus, TNF-α is a proinflammatory cytokine that may interfere significantly with bone metabolism of these mice. Studies have verified that this cytokine participates in bone resorption by stimulating osteoclastogenesis (Lam et al.,2000; Kitaura et al.,2005; Ochi et al.,2007; Karieb and Fox,2011). In addition, TNF-α can also act on bone formation, since there is evidence showing that TNF-α is crucial in the initiation of bone healing and in intramembranous bone formation during fracture repair (Kon et al.,2001; Gerstenfeld et al.,2003). Other researchers have suggested that the action of TNF-α is dependent on its concentration (Wahl et al.,2010; Glass et al.,2011). According to Wahl et al. (2010), “a homeostatic level of TNF signaling is required for optimal bone formation but that unregulated or excessive expression results in pathology.” Glass et al. (2011) found that exogenous administration of TNF-α locally accelerated fracture repair and remodeling of the tibia in a murine model, whereas Hashimoto et al. (1989) observed that high doses of this cytokine impaired the process of bone repair. Considering the evidence presented, we suggest that the elevated serum level of TNF-α existing during muscle degeneration could compromise bone quality of the mdx mouse. On the other hand, TNF-α could have its serum level decreased during episodes of muscle regeneration and thus achieve an appropriate level that could be beneficial for calvarial bone healing.
The secretion of cytokines and chemokines plays an important role in the recruitment of leukocytes and macrophages. These cells have been suggested to increase the deposition of collagen and the migration of fibroblasts into the wound area in mdx mice (Straino et al.,2004). Therefore, the secretion of cell signaling molecules that mediate and regulate inflammatory and immune responses seems to compensate for the poor bone quality seen in mdx mice, increasing bone repair capacity. In addition, this bone repair capacity probably depends directly on the age and number of cycles of muscle degeneration/regeneration since these cycles decrease in mdx mice at about 100–120 days of age (Tanabe et al.,1986; Evans et al.,2009).
Studies have demonstrated that long bones of mdx mice are impaired at 21 (Nakagaki et al.,2011), 28 (Anderson et al.,1993), and 49 (Novotny et al.,2011) days of age and also at 24 months of age (Novotny et al.,2011). However, femur bones of mdx mice have presented higher mineral density and mechanical resistance (strength) at 4 months of age compared to control animals (Montgomery et al.,2005). These facts are in contrast with DMD patients that have a progressive bone loss. Thus, the mechanism of compensation suggested in this study can only be observed in mdx mice. Osteogenic factors arising from muscle necrosis associated with other factors released from efficient muscle regeneration in mdx mice, such as insulin-like growth factor-1 (Hamrick et al.,2010), could act together to promote the bone healing.
Nakagaki et al. (2011) concluded that the femur of mdx mice presents a lower degree of collagen organization and suggested a reduction in the quantity of crosslinks and proteoglycans involved in fibrillogenesis. Collagen regulates the spatial sequence during mineralization by defining the site of mineral deposition and growth (Siperko and Landis,2001), whereas the temporal sequence of this process is usually regulated by bone matrix macromolecules (noncollagen proteins) such as proteoglycans (Young,2003). These macromolecules facilitate the nucleation of hydroxyapatite crystals or heterogeneous nucleation and sequester mineral ions to increase the local concentration of calcium and/or phosphorus (Clarke,2008). Taken together, these findings suggest that the mechanism of compensation discussed earlier may have favored the organization of bone extracellular matrix and mineralization during the bone healing.
In conclusion, we suggested that the genetic deficiency of mdx mice might have been compensated for by an adaptive response of the organism to pathological levels of muscle degeneration. In this respect, the age of the animals and the number of muscle degeneration cycles probably interfered with bone regeneration in mdx mice, compensating for the genetic deficiency by increasing bone formation rates during the healing process.
- 2004. Applications of a mouse model of calvarial healing: differences in regenerative abilities of juveniles and adults. Plast Reconstr Surg 114: 713–720. , , , , , , , , , .
- 1993. Recovery from disuse osteopenia coincident to restoration of muscle strength in mdx mice. Bone 14: 625–634. , , .
- 2002. Decreased bone density in ambulatory patients with Duchenne muscular dystrophy. J Pediatr Orthop 22: 179–181. , , .
- 2003. Bone mineral density and bone metabolism in Duchenne muscular dystrophy. Osteoporos Int 14: 761–767. , , , , , , .
- 2005. Bone health in Duchenne muscular dystrophy: a workshop report from the meeting in Cincinnati, Ohio, July 8, 2004. Neuromuscul Disord 15: 80–85. , , , , , .
- 2004. Histological parameters for the quantitative assessment of muscular dystrophy in the mdx-mouse. Neuromuscul Disord 14: 675–682. , , , , .
- 2004. Perspectives on glucocorticoid-induced osteoporosis. Bone 34: 593–598. , , , .
- 1994. Osteocalcin induces chemotaxis, secretion of matrix proteins, and calcium-mediated intracellular signaling in human osteoclast-like cells. J Cell Biol 127: 1149–1158. , , , , , , , , , .
- 2008. Normal bone anatomy and physiology. Clin J Am Soc Nephrol 3: S131–S139. .
- 2003. Age-related changes in the biomolecular mechanisms of calvarial osteoblast biology affect fibroblast growth factor-2 signaling and osteogenesis. J Biol Chem 278: 32005–32013. , , , , .
- 1994. Elevated basic fibroblast growth factor in the serum of patients with Duchenne muscular dystrophy. Ann Neurol 35: 362–365. , , , , , , , .
- 2009. Dysregulated intracellular signaling and inflammatory gene expression during initial disease onset in Duchenne muscular dystrophy. Am J Phys Med Rehabil 88: 502–522. , , , , .
- 2003. Impaired fracture healing in the absence of TNF-alpha signaling: the role of TNF-alpha in endochondral cartilage resorption. J Bone Miner Res 18: 1584–1592. , , , , , , , , .
- 2011. TNF-alpha promotes fracture repair by augmenting the recruitment and differentiation of muscle-derived stromal cells. Proc Natl Acad Sci USA 108: 1585–1590. , , , , , .
- 2000. Osteogenesis in cranial defects: reassessment of the concept of critical size and the expression of TGF-b isoforms. Plast Reconstr Surg 106: 360–371. , , , , , , .
- 2000a. Biomolecular mechanisms of calvarial bone induction: immature versus mature dura mater. Plast Reconstr Surg 105: 1382–1392. , , , , , , , , , .
- 2000b. Regional differentiation of cranial suture-associated dura mater in vivo and in vitro: implications for suture fusion and patency. J Bone Miner Res 15: 2413–2430. , , , , , , , .
- 2008. Implications of cross-talk between tumour necrosis factor and insulin-like growth factor-1 signalling in skeletal muscle. Clin Exp Pharmacol Physiol 35: 846–851. , , , , .
- 2004. Anti-TNFalpha (Remicade) therapy protects dystrophic skeletal muscle from necrosis. FASEB J 18: 676–682. , .
- 2010. Role of muscle-derived growth factors in bone formation. J Musculoskelet Neuronal Interact 10: 64–70. , , .
- 1989. Inhibitory effects of tumor necrosis factor alpha on fracture healing in rats. Bone 10: 453–457. , , , , , , , .
- 1982. Skeletal changes in children with neuromuscular disorders. In: Dixon AD, Sarnat BG, editors. Factors and mechanisms influencing bone growth. New York: Alan R. Liss. p 553–557. .
- 2000. Elevated plasma levels of transforming growth factor beta1 in patients with muscular dystrophy. NeuroReport 11: 4033–4035. , , , , , , , , .
- 2011. Phytoestrogens directly inhibit TNF-α-induced bone resorption in RAW264.7 cells by suppressing c-fos-induced NFATc1 expression. J Cell Biochem 112: 476–487. , .
- 2005. M-CSF mediates TNF-induced inflammatory osteolysis. J Clin Invest 115: 3418–3427. , , , , , .
- 2001. Expression of osteoprotegerin, receptor activator of NF-kappaB ligand (osteoprotegerin ligand) and related proinflammatory cytokines during fracture healing. J Bone Miner Res 16: 1004–1014. , , , , , , , .
- 2000. TNF-alpha induces osteoclastogenesis by direct stimulation of macrophages exposed to permissive levels of RANK ligand. J Clin Invest 106: 1481–1488. , , , , , .
- 2000. Bone mineral density and fractures in boys with Duchenne muscular dystrophy. J Pediatr Orthop 20: 71–74. , .
- 2002. Fracture prevalence in Duchenne muscular dystrophy. Dev Med Child Neurol 44: 695–698. , , , , , , , , .
- 2005. Muscle-bone interactions in dystrophin-deficient and myostatin-deficient mice. Anat Rec A Discov Mol Cell Evol Biol 286: 814–822. , , , .
- 2011. Mechanical, biochemical and morphometric alterations in the femur of mdx mice. Bone 48: 372–379. , , , , .
- 2011. Bone is functionally impaired in dystrophic mice but less so than skeletal muscle. Neuromuscul Disord 21: 183–193. , , , , , .
- 2007. Pathological role of osteoclast costimulation in arthritis-induced bone loss. Proc Natl Acad Sci USA 104: 11394–11399. , , , , , , , , .
- 2008. Curcumin alleviates dystrophic muscle pathology in mdx mice. Mol Cells 25: 531–537. , , , , , , , .
- 2010. IGF-I released during osteoclastic bone resorption induces osteoblast differentiation of BMSCs in the coupling process. J Bone Miner Res 25: S36. , , , , , , .
- 1995. Mdx mice show progressive weakness and muscle deterioration with age. J Neurol Sci 129: 97–105. , .
- 2003a. Persistent over-expression of specific CC class chemokines correlates with macrophage and T-cell recruitment in mdx skeletal muscle. Neuromuscul Disord 13: 223–235. , , , , , , , , .
- 2003b. Dissection of temporal gene expression signatures of affected and spared muscle groups in dystrophin-deficient (mdx) mice. Hum Mol Genet 12: 1813–1821. , , , , .
- 2005. Postnatal changes in sarcolemmal organization in the mdx mouse. Neuromuscul Disord 15: 552–561. , .
- 2009. Mechanisms inducing low bone density in Duchenne muscular dystrophy. Bone 44: S237–S238. , , , , , , , , , , , .
- 2006. Repair of bone defects treated with autogenous bone graft and low-power laser. J Craniofac Surg 17: 297–301. , .
- 2001. Aspects of mineral structure in normally calcifying avian tendon. J Struct Biol 135: 313–320. , .
- 2007. Low bone mineral density and decreased bone turnover in Duchenne muscular dystrophy. Neuromuscul Disord 17: 919–928. , , , , , , .
- 2004. Enhanced arteriogenesis and wound repair in dystrophin-deficient mdx mice. Circulation 110: 3341–3348. , , , , , , , , , .
- 1986. Skeletal muscle pathology in X chromosome-linked muscular dystrophy (mdx) mouse. Acta Neuropathol 69: 91–95. , , .
- 2003. Nerve growth factor expression in human dystrophic muscles. Muscle Nerve 27: 370–373. , , , , , , , .
- 2001. Fracture risk in patients with Duchenne muscular dystrophy and spinal muscular atrophy. J Rehab Med 33: 150–155. , , , , , .
- 2009. Osteopontin promotes fibrosis in dystrophic mouse muscle by modulating immune cell subsets and intramuscular TGF-beta. J Clin Invest 119: 1583–1594. , , , , , , , .
- 2010. Restoration of regenerative osteoblastogenesis in aged mice: modulation of TNF. J Bone Miner Res 25: 114–123. , , , , , , , , , , , , , , , .
- 2006. Satellite cells from dystrophic (mdx) mice display accelerated differentiation in primary cultures and in isolated myofibers. Dev Dyn 235: 203–212. , .
- 2003. Bone matrix proteins: their function, regulation, and relationship to osteoporosis. Osteoporos Int 14: S35–S42. .