Skip to main content

Advertisement

Dominant negative Bmp5mutation reveals key role of BMPs in skeletal response to mechanical stimulation

Article metrics

Abstract

Background

Over a hundred years ago, Wolff originally observed that bone growth and remodeling are exquisitely sensitive to mechanical forces acting on the skeleton. Clinical studies have noted that the size and the strength of bone increase with weight bearing and muscular activity and decrease with bed rest and disuse. Although the processes of mechanotransduction and functional response of bone to mechanical strain have been extensively studied, the molecular signaling mechanisms that mediate the response of bone cells to mechanical stimulation remain unclear.

Results

Here, we identify a novel germline mutation at the mouse Bone morphogenetic protein 5 (Bmp5) locus. Genetic analysis shows that the mutation occurs at a site encoding the proteolytic processing sequence of the BMP5 protein and blocks proper processing of BMP5. Anatomic studies reveal that this mutation affects the formation of multiple skeletal features including several muscle-induced skeletal sites in vivo. Biomechanical studies of osteoblasts from these anatomic sites show that the mutation inhibits the proper response of bone cells to mechanical stimulation.

Conclusion

The results from these genetic, biochemical, and biomechanical studies suggest that BMPs are required not only for skeletal patterning during embryonic development, but also for bone response and remodeling to mechanical stimulation at specific anatomic sites in the skeleton.

Background

An area of significant interest in orthopaedics and rehabilitation medicine is the effect of mechanical loading on bone formation and remodeling. Mechanical stimulation plays an important role in determining bone mass and density in the adult skeleton, as well as susceptibility to conditions such as fractures or osteoporosis. It has long been observed that bone mass and mineral density can be altered at very specific sites of the skeleton in response to mechanical stimulation during exercise, as seen in increased size and cortical thickness of the arm bone from the dominant side in tennis players [13] and the increased mineralization seen in the lumbar spine of weight lifters [4] or in the heel bone of runners [5]. In general, increased exercise or muscular loading will increase bone mass [6, 7] or bone density [8, 9]. In contrast, decreased loading will reduce osteogenic activity, as seen in the bones of test animals in space flight [10] or of patients in prolonged bed rest [11].

Since Wolff's observation in 1892 that mechanical stress is a primary determinant in bone adaptation [12], extensive studies have been performed to understand how bone responds to its mechanical environment. Frost proposed a "mechanostat" theory [13] in which the skeleton senses mechanical stimuli that are above a certain threshold and bone formation is activated. After cell-mediated bone remodeling, a feedback system resets this threshold. However, the exact mechanism by which this mechanostat converts biophysical force to a cellular response is unknown. Various mechanisms have been proposed to involve hydrostatic pressure [14, 15], mechanical stretch [1619], fluid shear [2022], and others. The signals activated by these mechanisms have been postulated to act via mechanically sensitive ion channels [2325], the integrin-cytoskeleton pathway [2628], phospholipase C [17, 29, 30], or G protein cascades [31, 32] to trigger a cellular response.

Bone morphogenetic proteins (BMPs) belong to the Transforming growth factor-beta (TGF-β) family of secreted signaling molecules [33]. Although previous studies have revealed much about the important role of BMPs in skeletal patterning in embryogenesis, many of these studies were limited by two issues. First, since BMPs are required for multiple aspects of organogenesis, loss of function mutations often produce animals with prenatal lethality due to pleiotropy [3437]. Second, multiple coexpressing BMPs can produce functional redundancy and mask the effect of loss of function of a single BMP [3840].

Previous null mutations identified at the short ear/Bmp5 locus have shown that early condensation and growth of cartilage precursors in the ear, rib, and vertebra require BMPs [41, 42]. In 1987, a new Bmp5 mutation causing unusually short ears in mice arose spontaneously at The Jackson Laboratory. To gain further insight into the role of Bmp5 in skeletal development, these mice were used to identify the location of this novel Bmp5 mutation and its effect on the processing and activity of BMP5.

To further investigate the role of BMPs in development, mice which were homozygous for this novel Bmp5 mutation were generated. Our findings indicate that the mutation disrupts the processing of the BMP5 peptide and may inactivate BMP5. Furthermore, these mutant mice displayed severe defects at specific skeletal structures that were even more severe than those of Bmp5 null mutants. Some of the skeletal defects were observed at sites of bone-muscle interaction. Biomechanical studies show that mutant osteoblasts from these sites failed to respond appropriately to mechanical strain and may implicate BMPs as the endogenous signals for bone formation in response to mechanical stimulation.

Results

Bmp5cleavage mutation disrupts the proteolytic processing of the BMP5 protein

Sequencing of this newly discovered Bmp5 mutation revealed a correctly spliced Bmp5 transcript with a G-to-A substitution at base 932 of the Bmp5 coding region (Fig. 1a, b). This change destroys a Taq1 site in the second exon (Fig. 1c), providing a simple assay for following the mutation in genetic crosses.

Figure 1
figure1

Bmp5clv, a Bmp5 cleavage sequence mutation, disrupts the proteolytic processing of the BMP5 protein. (a) Schematic of the Bmp5 open reading frame showing the leader signal (LEAD), pro (PRO) and mature (MAT) domains and the putative proteolytic processing site (scissors). The G-to-A mutation (*) at nucleotide 932 of the Bmp5clvallele is predicted to destroy a Taq1 restriction site and to disrupt the first arginine residue in the putative conserved "RXXR" cleavage sequence in BMP5 (box). "1" and "2" denote positions of PCR primers used in typing the Bmp5clvallele. (b) Partial sequence traces showing the G-to-A substitution (*) in Bmp5clvmutant mice. This alteration is the only nucleotide difference in the Bmp5 coding region between wild-type (wt) and Bmp5clvmice. (c) Confirmation of mutation in genomic DNA. A 158-bp PCR product (gray arrowhead) containing the site of the Bmp5clvmutation is cleaved by Taq1 into 67- and 91-bp fragments (black arrowheads) in wild-type mice, partially cleaved in Bmp5clv/+ heterozygous mice, and not cleaved in Bmp5clv/Bmp5clvmice. (d) COS-7 cells were transfected with a mammalian expression vector (vec) or the same vector driving expression of wild-type (wt) or mutant (Bmp5clv) BMP5 protein. Secreted proteins were analyzed by Western blot with antibodies against the pro (anti-PRO) or mature (anti-MAT) domain of murine BMP5. Most of the wild-type protein expressed was in the smaller cleaved form, whereas all the detectable mutant protein was non-processed. No appreciable signal was detected by either antibody in the control cells. (e) A proposed dominant-negative mechanism for the Bmp5clvmutation. Wild-type BMP5 peptides are cleaved at the proteolytic site to form functional dimers with another wild-type copy of BMP5 (gray bar) or another related BMP (black bar). Cleavage mutants produce non-processed BMP5 peptides that bind other non-processed BMP5 peptides to form inactive homodimers or bind and sequester wild-type BMP5 or other related BMPs in defective heterodimers.

Most TGF-β superfamily proteins are synthesized as larger precursors that are cleaved at an RXXR consensus site by proprotein convertase endoproteases to generate an N-terminal pro domain and a C-terminal signaling domain [43, 44]. The G-to-A mutation changes the first arginine in the RXXR processing site of BMP5 to a glutamine (Fig. 1a). To test whether the Arg311Gln mutation disrupts BMP5 protein processing, we expressed the wild-type or mutant forms in COS-7 cells and analyzed conditioned media by Western blot analysis with antibodies raised to the pro and mature regions of BMP5. Cells transfected with the wild-type construct produced BMP5 protein bands of ~40 kDa and ~20 kDa, consistent with the expected sizes of the cleaved BMP5 pro and mature domains (Fig. 1d). Both antibodies also detected a minor protein band of ~55 kDa (Fig. 1d), suggesting that some secreted BMP5 protein was unprocessed. Cells expressing mutant BMP5 produced only the unprocessed ~55 kDa protein form (Fig. 1d), confirming that the mutation in the RXXR site blocks normal proteolytic cleavage. We termed this mutation Bmp5clvto denote the lesion at the cleavage site.

Bmp5clvmutants exhibit an array of skeletal defects

Mutations in the RXXR site of other TGF-β family members have inhibited the processing and activity of the corresponding protein [4548]. Such mutations also have acted as dominant-negative mutations that block the function of other coexpressed TGF-β members, presumably by sequestering them into inactive heterodimer complexes with the unprocessed mutant subunits [45, 47]. Mice heterozygous for the Bmp5clvmutation showed mild skeletal defects not seen in wild-type or +/Bmp5nullmice, including reduction of the spinous process at the second thoracic vertebrae (data not shown). Such defects are consistent with a mild dominant-negative effect observed when the Bmp5clvallele is present in single copy, consistent with the mode of action observed for similar mutations in other TGF-β members [45, 47].

We expect homozygosity for the mutation to further decrease the activity of BMP5 and increase the production of the non-processed BMP5 molecules that may inactivate other coexpressed BMPs. To determine the effect of this mutation on homozygotes carrying this allele, we crossed heterozygous carriers of the Bmp5clvmutation and generated viable Bmp5clv/Bmp5clvhomozygotes, but at rates ~10-times lower than Mendelian predictions (4/156 progeny, P < 0.001). Despite the increased prenatal lethality, some Bmp5clvhomozygotes survived with normal life spans and fertility. Among these surviving homozygous Bmp5clvmice, we noted more severe defects than those seen with age-controlled Bmp5nullhomozygotes, including shorter external ears (wild-type: 6.4 ± 0.4 mm, null: 4.8 ± 0.2 mm, Bmp5clv: 2.9 ± 0.2 mm; P < 0.01), loss of lesser horns of the hyoid (Fig. 2a), more misshapen xiphisternum and missing ribs (Fig. 2f), less calcification of thyroid cartilage (Fig. 2a), abnormal bony fusion (fused posterior sternum; Fig. 2f), and reduced or absent processes on the sixth cervical (Fig. 2b), second thoracic (Fig. 2d), and lumbar vertebrae (Fig. 2e). The spectrum of phenotypes, and the consistent reduction rather than overgrowth of skeletal tissue, both suggest that the Bmp5clvmutation leads to loss rather than gain of BMP5 activity.

Figure 2
figure2

Bmp5clvmutation causes more severe skeletal defects than a Bmp5 null ( Bmp5null) mutation. Alizarin red-stained bones of 12-week +/+, Bmp5null/Bmp5null, and Bmp5clv/Bmp5clvmice show the following: (a) shortening of the greater horn (gh) and lesser horn (lh) of the hyoid bone and decreased calcification of the thyroid cartilage (tc), (b) absence of anterior tubercles (at) and thinning of the neural arch (na) of the 6th cervical vertebra, (c) reduction of the sesamoid (s) and nearly complete loss of the deltoid tuberosity (dt) of the humerus, (d) absence of the spinous process (sp) of the 2nd thoracic vertebra, (e) absence of the transverse process (tp) and anapophysis (a) of the 3rd lumbar vertebra, and (f) abnormal fusion of posterior sternal segments and loss of the xiphoid process (x) at the end of the sternum in Bmp5clv/Bmp5clvmice.

Bmp5clvmutants display a deficiency at a mechanosensitive site of the skeleton

An interesting new phenotype in Bmp5clvmice is marked reduction or complete elimination of the deltoid crest on the humerus bone (Fig. 2c). The deltoid crest is a prominent bony ridge that is the insertion site for the deltoid muscle, and it normally forms in response to mechanical interaction between muscle and bone. In paralyzed animals or those exhibiting genetically defective muscle formation, the deltoid crest does not form [49, 50]. In contrast, mutant animals showing abnormal increases in muscle mass develop bigger deltoid tuberosities [51].

The deltoid muscle remained present in Bmp5clv/Bmp5clvmutants, suggesting that the deltoid crest defect was likely due to changes in the humerus bone. In situ hybridization showed Bmp5 expression in the developing deltoid crest of the humerus (Fig. 3b), and coexpression of the Bmp2 and Bmp6 genes in similar regions (Fig. 3c, d). The coexpression of multiple Bmps may explain why the deltoid crest is only mildly reduced in Bmp5nullmice but completely eliminated in Bmp5clvmice. The expression of multiple Bmp genes at the deltoid crest and the defect of this structure in Bmp5clvmutants suggest that Bmp signaling is important at this mechanosensitive site.

Figure 3
figure3

Multiple Bmps are expressed at the deltoid tuberosity. In situ hybridization analysis of the developing deltoid tuberosity at embryonic day 13.5 with antisense probes to (a) collagen 2, (b) Bmp5, (c) Bmp2, or (d) Bmp6 shows expression of multiple Bmps at the developing deltoid tuberosity (arrowheads). Control sense probes did not detect any appreciable signal (data not shown).

Bmp5clvmutation alters the response of deltoid crest osteoblasts to mechanical stimulation

The loss of a prominent muscle-induced skeletal feature in Bmp5clvmice suggests that Bmp signaling plays a key role in bone cells' response to mechanical activity. To test this hypothesis, we isolated osteoblasts from the deltoid tuberosity of 10-month-old wild-type and Bmp5clvmice and subjected the cells to 24 hours of cyclic uniform radial strain in culture. The stretch regimen we applied (10-second maximum 15% elongation, then 10-second relaxation, frequency 0.05 Hz or 3 cycles/minute) is similar to the mechanical stimulation known to induce cellular responses in cultured osteoblasts [18, 19]. After 24-hour cyclic strain, control osteoblasts became spindle-shaped, showed elongation of cellular processes, and were largely oriented perpendicular to the radial strain field (Fig. 4 and Table 1). In contrast, osteoblasts from the deltoid tuberosity of the Bmp5clvmice displayed no significant changes in morphology or orientation after mechanical strain (Fig. 4 and Table 1), suggesting that the defect in BMP signaling blocked normal response of bone cells to mechanical stimulation.

Table 1 Response of primary osteoblasts and muscle fibroblasts to cyclic mechanical strain.
Figure 4
figure4

Altered response to mechanical stimulation in deltoid tuberosity osteoblast cells of Bmp5clv/ Bmp5clvmutants. Cultured cells from the indicated sites were subjected to 0- or 24-h cyclic mechanical strain and visualized afterwards by indirect immunofluorescence using antibodies against collagen 1 (osteoblast) or vimentin (fibroblast). After subjection to 24-h strain, most osteoblasts from the deltoid tuberosity of wild-type (wt) mice become spindle-shaped and reorient perpendicular to the strain axis (arrow). In contrast, Bmp5clvdeltoid tuberosity osteoblasts display a random orientation after mechanical strain. Femur trochanter osteoblasts from wild-type or Bmp5clvmice both realign after mechanical strain. Deltoid muscle fibroblasts from wild-type or Bmp5clvmice also respond appropriately to mechanical strain.

Osteoblasts cultured from an independent bone-muscle interaction site that does not show morphologic defects in Bmp5clvmice responded normally to mechanical strain in vitro (femur trochanter osteoblasts; Fig. 4). The anatomic site-specificity of osteoblast response to mechanical stimuli in Bmp5clvmice is consistent with the specificity of the skeletal defects seen at the tissue level of these mutants. Furthermore, muscle fibroblasts isolated from the deltoid of both wild-type and Bmp5clvmice showed normal changes in morphology and orientation after mechanical stimulation in vitro (Fig. 4 and Table 1), suggesting that the mutation primarily affects bone cells and not muscle cells at these sites.

To further characterize the relationship between mechanical stimulation and BMP signaling, we studied the effect of mechanical strain on cellular translocation of SMAD proteins, key transcription factors that translocate from the cytoplasm to the nucleus upon activation of BMP receptors [52]. Non-strained wild-type deltoid tuberosity osteoblasts exhibited SMAD1/5 immunoreactivity predominantly in the cytoplasm (Fig. 5 and Table 1). Within 30 minutes of applying cyclic strain to these cells, we detected a significant increase in nuclear localization of SMAD1/5 (Fig. 5 and Table 1). Peak nuclear localization occurred 1 hour after mechanical stimulation, SMAD1/5 was then predominantly cytoplasmic again by 24 hours (Fig. 5 and Table 1), at which point cells also had reoriented perpendicularly to the strain axis. In contrast, SMAD1/5 in Bmp5clvdeltoid tuberosity cells remained mostly cytoplasmic before and after cyclic stretch (Fig. 5 and Table 1), demonstrating that perturbation of BMP signaling by the Bmp5clvmutation disrupts bone cells' rapid response to mechanical stretch.

Figure 5
figure5

SMAD1/5 nuclear relocalization after mechanical stimulation in deltoid tuberosity cells. Resting wild-type (wt) deltoid tuberosity osteoblasts display SMAD1/5 immunoreactivity in the cytoplasm; however, with increased duration of mechanical strain SMAD immunoreactivity becomes more nuclear. At 24 hours, most of the reoriented cells again display cytoplasmic localization of SMAD 1/5. In contrast, cyclic strain failed to elicit any significant nuclear translocation of SMAD1/5 immunoreactivity in Bmp5clvdeltoid tuberosity osteoblasts during the time period tested. Nuclei were counterstained with propidium iodide (P.I.).

While mutant osteoblasts from deltoid crest displayed altered response to mechanical stimuli, it was unclear whether this was due to an ongoing requirement for BMP-mediated signaling or abnormal cell development at the deltoid crest of Bmp5clvmice. To distinguish between these possibilities, we tested the effect of transiently inhibiting BMP signaling in wild-type osteoblasts cultured with increasing concentrations of noggin, a secreted protein that binds BMP and inhibits its activity [53]. Incubation of adult wild-type osteoblasts with noggin produced dose-dependent decreases in their reorientation response to mechanical stimulation (Table 1), confirming that BMP signaling is important in maintaining the normal mechanical responses of osteoblasts at postnatal stages.

Discussion

Although the role of BMPs in the formation of cartilage, bone, and other tissues during embryonic development is well-established, studies of their functions during postnatal development are complicated by both the requirement of BMP signaling for many developmental events (pleiotropy) and the overlapping expression and roles of multiple BMPs at particular sites (partial functional redundancy). The spontaneous Bmp5clvallele is a missense mutation at a site that encodes the consensus cleavage sequence of BMP5 and blocks the post-translational processing of the BMP5 protein. Interestingly, this allele mimics a form of dominant-negative mutation that researchers have artificially induced in other TGF-β members and injected into frog or fish embryos [46, 48] or expressed in transgenic animals [45, 47] in attempts to inhibit endogenous activity. The processing-defective Bmp5clvallele identified here may be able to inactivate multiple BMPs but only at sites of endogenous Bmp5 gene expression, providing a unique tool to uncover new functions of Bmp genes during embryonic and postnatal development.

Skeletal analyses show that the Bmp5clvallele has a dominant negative effect. Bmp5clvheterozygote animals show skeletal defects not seen in Bmp5nullheterozygotes, and Bmp5clvhomozygote animals have a worse phenotype compared to that of Bmp5nullmutants. Interestingly, studies on others TGF-β members have also described a dominant negative phenomenon displayed by similar cleavage sequence mutants. In those studies, the non-processed mutant proteins exert their effect on coexpressed wild type proteins by sequestering them into inactive complexes [4548]. It remains to be shown whether BMP5clvheterodimerizes with BMP2 or BMP6, which are co-expressed at the deltoid crest. Inactivation of BMP2 and BMP6 by the mutant BMP5 protein may explain why the deltoid crest is only mildly reduced in Bmp5nullmice but severely reduced in Bmp5clvmice.

While the skeletal analysis was performed in adult Bmp5clvmutants, given the expression of Bmp5 seen at the early embryonic stages of humeral development in our expression studies, it is possible that BMPs exert their effect on this structure at an early developmental stage. It will be interesting to perform histological analysis at various development time points to better characterize the phenotype of this mutation at different stages of development.

The skeletal defects we observed in the Bmp5clvmutants are localized to specific structures. Interestingly, previous studies show that the expression of Bmp5 is controlled by an array of cis-acting regulatory sequences that drive Bmp5 expression at highly specific anatomical locations in the skeleton [54, 55]. Accordingly, the unique expression pattern of Bmp5 may account for the specificity of skeletal defects seen in the Bmp5clvmutants.

The deltoid tuberosity defect in Bmp5clvmice suggests that the formation of a prominent muscle-induced bony structure in the humerus requires BMP signaling. Our biomechanical studies confirm that BMP signaling is fundamental to the early response of bone cells to mechanical stimulation. While we have shown that osteoblasts respond to mechanical stimulation by increasing the nuclear translocation of SMAD1/5, others have also shown similar in vitro activation of BMP signaling with mechanical stimulation. Previous studies have shown elevated BMP mRNA levels in response to 6 hours of tensile stress applied to neonatal mouse calvaria [56] or cyclic stress applied to cultured chick chondrocytes [57]. Our results demonstrate SMAD translocation in normal osteoblasts in vitro within 30 minutes of mechanical stimulation and show that Bmp5clvcells fail to undergo SMAD translocation or reorientation in response to similar mechanical strain. The altered response to mechanical loading in vitro and the lack of muscle-induced features in Bmp5clvmice provide strong genetic evidence that BMP signaling is integral to the normal response of bone cells to mechanical strain. While our data show that this mutation has an impact on osteoblasts, it remains to be determined whether perturbation of BMP signaling has any effect on chondrocytes from the deltoid crest.

While mutant osteoblasts from deltoid crest displayed altered response to mechanical stimuli, it was unclear whether this was due to an ongoing requirement for BMP-mediated signaling or abnormal cell development at the deltoid crest of Bmp5clvmice. The altered response of wildtype deltoid crest osteoblasts to mechanical strain when the BMP signaling pathway was blocked with noggin suggests that BMPs are important in maintaining the ability of bone cells to respond to mechanical strain postnatally. It will be interesting to perform further biomechanical studies on Bmp5clvosteoblasts in the presence of exogenous BMP5 protein to ascertain whether the Bmp5clvmutation also affects the proper cell development of osteoblasts at the deltoid crest.

Conclusion

Over the years, researchers have recognized that the mechanical response of bone strongly influences human health and disease. Weight-bearing exercise can increase bone mass and density and can reduce the risk of fracture in millions of persons with predisposing factors including osteoporosis. Numerous studies have provided a better understanding of the mechanical stimuli that bone cells detect, the signaling pathways that transduce mechanical signals to the cell, and the nature of the cellular response. This study on Bmp5clvmice shows that BMP signaling is an important part of this mechanotransduction system in bone. Further studies may elucidate how the BMP pathway interacts with other signaling pathways in this process. Modulation of BMP signaling through recombinant protein or gene therapy may enable clinicians to potentiate the benefits of weight-bearing activity to the skeleton or improve the treatment of bony diseases caused by prolonged lack of mechanical stimulation.

Methods

Mouse strains and skeletal preparation

The Bmp5clvmutation was originally designated "se-4J" when discovered as a spontaneous mutation in a backcross between C57Bl/6J and B6.Cg-Otop1tlt(for further strain details, see JAX stock number 001496). The Bmp5clvstock was maintained in the laboratory of DMK by intercrossing Bmp5clv/+ heterozygotes. Mutants were identified by the short ear phenotype, and their genotype was confirmed by molecular typing as described in this paper. The classical se mutation was also maintained on the C57Bl/6J background (Jackson Laboratory stock number 000056). Skeletons from age- and sex-matched mice from different genotypes were fixed in 95% ethanol, processed using a potassium hydroxide/alizarin red (Sigma) staining procedure [58], disarticulated, and analyzed.

Identification of the Bmp5clvmutation

The Bmp5 open reading frame from total lung RNA of a 1-month-old C57Bl/6J male mouse and an age- and sex-matched Bmp5clv/Bmp5clvmutant mouse was amplified. Reverse transcription was carried out with Superscript RT (Gibco) using 1.5 μg of RNA and 2.5 μL of 20-μM reverse primer (5'-CGC GGA TCC CTA GTG GCA GCC ACA CGA-3') in a 20-μL reaction at 37°C for 1 h. Polymerase chain reaction (PCR) was performed with Amplitaq DNA polymerase (Perkin-Elmer) in a 50-μL reaction containing 1 μL each of 20-μM forward (5'-CGC GGA TCC ACC ATG CAT TGG ACT GTA TTT TTA C-3') and reverse (as above) primers and 1 μL of the RT reaction product from above. The PCR products were purified from a 1% regular agarose gel with the Gene-clean kit (Bio 101), digested with BamHI, and cloned into pBluescript II SK(+) (Stratagene). The Bmp5 cDNA insert on both strands from multiple clones of each genotype was sequenced with the Sequenase kit (United States Biochemicals) and primers that span the Bmp5 open reading frame (primer sequences available upon request).

The genomic region affected by the Bmp5clvmutation was amplified with primers 1 (5'-AAA TCT GCT GGT CTT GTG GG-3') and 2 (5'-GGG TCC TGA TGA GAG TTG GA-3') in a 50-μL PCR reaction containing 2.5 μL each of 20-μM primers 1 and 2 and 1.5 μL of genomic DNA. The amplified products were digested by TaqI and separated by electrophoresis on a 3% low melt agarose gel.

Expression and detection of BMP5 protein

COS-7 cells were cultured in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% fetal bovine serum (FBS, Hyclone), 2-mM GlutaMax, 0.1-mM non-essential amino acids, penicillin G (100 U/mL), and streptomycin (100 μg/mL) at 37°C in 5% CO2/95% air and were passaged at confluency and maintained according to standard tissue culture practices. All tissue culture reagents were from Gibco except as indicated otherwise.

The Bmp5 open reading frame was amplified from wild-type or Bmp5clvhomozygous mice as described above, subcloned into the mammalian expression vector pCDNA3 (Invitrogen), and sequenced to ensure that no mutations were introduced in the cloning steps. In transfection experiments, 2 × 106 COS-7 cells were plated on a 100-mm tissue culture dish (Corning) and transfected with 3.3 μg of DNA and 10 μL of lipofectamine (Gibco) per dish under reduced serum condition for 7 h (as instructed by the manufacturer). The transfected cells were allowed to recover in full medium for 5 h, at which point the normal medium was replaced with reduced serum medium (growth medium with 1% FBS, 3 mL/plate). After ~60 h, the conditioned media from 2 identically treated plates were collected and pooled, clarified by centrifugation, made 2% SDS by addition of a 10% SDS stock, boiled for 10 min, and aliquoted and stored at -20°C.

Recombinant protein containing murine BMP5 pro region (amino acids 1–310) fused with thioredoxin was expressed under the IPTG-inducible pET32 system in BL21 E. Coli bacterial cells as instructed by the manufacturer (Novagen). Briefly, 8 L transformed bacterial culture was grown to a density of OD600 ~0.5 and induced with 1-mM IPTG for 4 h at 37°C. Cells were collected by centrifugation and resuspended in Buffer A (6-M Guanidine HCl, 0.1-M NaH2PO4, 0.01-M Tris pH 8.0, 5 mL/g cell pellet weight), and the clarified supernatant was incubated with Ni-NTA nickel resin beads (Qiagen) for 1 h at room temperature (RT). The beads were washed with 10 volumes of Buffer A, 10 volumes of Buffer B (8-M urea, 0.1-M NaH2PO4, 0.01-M Tris pH 8.0), 10 volumes of Buffer C (8-M urea, 0.1-M NaH2PO4, 0.01-M Tris pH 6.3) with 5-mM imidazole, 10 volumes of Super Buffer C (Buffer C plus 250-mM NaCl, 0.5% Tween-20, 10-mM β-mercaptoethanol, 100-mM imidazole), and 2 washes of 10 volumes of Buffer C with 20-mM imidazole. Bound protein was eluted by boiling the beads in 2.5× Laemmli buffer. Polyclonal antibodies against the BMP5 pro domain (anti-PRO) were raised by immunizing rabbits with recombinant BMP5 pro region protein at the Berkeley Antibody Company. Monoclonal antibodies against the human BMP5 mature domain (anti-MAT) were provided by Genetics Institute, Inc., and were shown to cross-recognize the highly conserved murine BMP5 mature region.

For western blot analysis, 60 μL of the processed SDS samples were boiled for 5 min in reducing Laemmli buffer, fractionated on a 15% SDS-polyacrylamide gel, and transferred onto nitrocellulose. The membrane was incubated at RT for 3 h in blocking solution (5% non-fat dried milk in 150-mM NaCl, 10-mM Tris pH 8.0, and 0.05% Tween-20), probed at RT for 12 h with anti-BMP5 primary antibodies (anti-PRO 1:667 or anti-MAT 1:172) diluted in blocking solution, and incubated for 1 h at RT in blocking solution containing donkey anti-rabbit-IgG coupled to horseradish peroxidase diluted 1:5000 (anti-PRO) or sheep anti-mouse-IgG coupled to horseradish peroxidase diluted 1:1000 (anti-MAT). Signals on the blot were detected by the ECL system (Amersham).

In situhybridization

Forelimbs were dissected from E13.5 CD1 embryos, frozen immediately in O.C.T. embedding medium (VWR), and stored at -80°C. Twelve-micrometer cross sections of the humerus at the level of the deltoid tuberosity were collected with a cryostat microtome and were processed, prehybridized, and hybridized with digoxygenin-labeled cRNA probes, as described in previously published protocols [40]. Bmp probes were generated by in vitro transcription of linearized constructs containing 200- to 500-bp DNA inserts from the pro region or untranslated region of Bmp genes as described previously [59]. Descriptions of the washing conditions and the methods used to detect the labeled signal have been published previously [40].

Isolation of primary osteoblasts and muscle fibroblasts

Primary osteoblast and muscle fibroblast isolates from sex-matched 10-month-old C57Bl/6J and Bmp5clvmice were established. Under microscopy, the humerus was exposed and the surrounding musculature was dissected off the bone. A ~3-mm segment of the humerus containing the deltoid tuberosity (wild type) or the deltoid muscle attachment site (Bmp5clv) was removed. Similarly, a ~5-mm segment of the femur shaft containing the femoral trochanter was dissected from each mouse. The marrow cavity of the bone segments was flushed with ice cold phosphate-buffered saline, and the bone was rinsed in phosphate-buffered saline containing penicillin G (200 U/mL), streptomycin (200 μg/mL), and amphotericin B (0.5 μg/mL). The rinsed bones were minced into small pieces with a bone cutter in 2 mL dissociation medium (21.3-mM Tris pH 7.4, 111.2-mM NaCl, 5.4-mM KCl, 1.3-mM MgCl2, 0.5-mM ZnCl2, and 13-mM glucose supplemented with type II collagenase [5 U/mL], elastase [6.25 U/mL], D-sorbitol [18.22 mg/mL], and chondroitin sulfate [6 mg/mL]) and digested at 37°C for up to 180 min. At 10, 20, and 30 min and every 30 min thereafter, the dissociated cells were removed with the medium and added to 1 volume of FBS (Invitrogen), and a fresh 2 mL of dissociation medium was added to the bone fragments. Cells from the 10–20 min, 30–90 min, and 120–180 min time points were pooled and recovered by centrifugation, resuspended in osteoblast growth medium (αMEM with 10% FBS, 10% horse serum [Invitrogen], 2-mM GlutaMax, and 0.1-mM nonessential amino acids) supplemented with penicillin G (100 U/mL), streptomycin (100 μg/mL), and amphotericin B (0.25 μg/mL), and plated in a 175-cm2 tissue culture flask. The plated cells were allowed to attach for 72 h at 37°C in 5% CO2/95% air, and non-adherent cells were rinsed off. Fresh growth medium with antibiotics and antimycotics was replaced every 4 days until the osteoblasts reached confluency (typically ~14 days after initial plating), when they were passaged and maintained similarly afterwards. An aliquot of cells from each flask was plated separately at first passage and assayed for alkaline phosphatase activity, an indicator of osteogenic differentiation, using the AP assay kit (Sigma). Cultures containing fewer than 50% alkaline phosphatase-positive cells were discarded. Antibiotics and antimycotics were omitted from the growth medium after the second passage. Experiments were typically performed on cells from the second and third passages.

A previously described fibroblast isolation protocol [60] was used to harvest primary fibroblasts from the distal third of the deltoid muscle by enzymatic dissociation. The fibroblasts were maintained in DMEM with 15% FBS, 2-mM GlutaMax, and 0.1-mM non-essential amino acids supplemented with penicillin G (100 U/mL), streptomycin (100 μg/mL), and amphotericin B (0.25 μg/mL).

Mechanical stimulation and indirect immunofluorescence

A total of 5 × 104 osteoblasts or 1 × 104 muscle fibroblasts per well were plated on BioFlex 6-well plates with collagen 1-coated flexible membrane bottoms (Flexcell) and allowed to attach overnight. For noggin treatment, fresh medium containing no (control) or recombinant mouse noggin protein (R&D, 0.1–10 μg/mL) was replaced 3 h before mechanical strain. The plates were mounted onto the base plate of a Flexercell FX-4000T system (Flexcell) and left undisturbed for 1 h before initiation of mechanical stimulation to minimize the extraneous strain and fluid stress secondary to handling of the plates.

Cyclic uniform radial strain was applied with -70 kPa negative pressure to produce a 15% maximum elongation of the membranes in square wave cycles of 10 sec strain and 10 sec relaxation at a frequency of 0.05 Hz or 3 cycles per minute.

We visualized the cells using a previously described indirect immunofluorescence protocol [60] with the following modifications. The cells were fixed in ice-cold 50% methanol/50% acetone at -20°C for 20 min; blocked in 10% serum with 0.5% Triton-X (Biorad); and probed with primary antibodies against type I collagen (Chemicon), vimentin (Sigma), or SMAD1/5 (Chemicon) and secondary antibodies coupled to FITC or Cy3 (Sigma). Nuclei were counterstained with propidium iodide (Molecular Probes). The round flexible membrane was cut into equal quadrants and each quadrant was mounted onto a slide. This allowed the angles between the cell axis and the strain field to be measured accurately. Angles were measured from 80–150 cells per condition over 3 independent experiments, and SMAD subcellular localization was determined from 80–150 cells per condition over 3 independent experiments. Images were captured with a Retiga 1300 digital camera (QImaging) attached to a Leica DM IRB microscope and were processed with Northern Eclipse 6.0 software (MVIA). P values and statistical significance between the means of each group were determined by the modified Bonferroni's t test.

References

  1. 1.

    Jones HH, Priest JD, Hayes WC, Tichenor CC, Nagel DA: Humeral hypertrophy in response to exercise. J Bone Joint Surg Am. 1977, 59: 204-208.

  2. 2.

    Haapasalo H, Kontulainen S, Sievanen H, Kannus P, Jarvinen M, Vuori I: Exercise-induced bone gain is due to enlargement in bone size without a change in volumetric bone density: a peripheral quantitative computed tomography study of the upper arms of male tennis players. Bone. 2000, 27: 351-357. 10.1016/S8756-3282(00)00331-8.

  3. 3.

    Huddleston AL, Rockwell D, Kulund DN, Harrison RB: Bone mass in lifetime tennis athletes. Jama. 1980, 244: 1107-1109. 10.1001/jama.244.10.1107.

  4. 4.

    Granhed H, Jonson R, Hansson T: The loads on the lumbar spine during extreme weight lifting. Spine. 1987, 12: 146-149. 10.1097/00007632-198703000-00010.

  5. 5.

    Williams JA, Wagner J, Wasnich R, Heilbrun L: The effect of long-distance running upon appendicular bone mineral content. Med Sci Sports Exerc. 1984, 16: 223-227.

  6. 6.

    Doyle F, Brown J, Lachance C: Relation between bone mass and muscle weight. Lancet. 1970, 1: 391-393. 10.1016/S0140-6736(70)91520-5.

  7. 7.

    Pocock N, Eisman J, Gwinn T, Sambrook P, Kelly P, Freund J, Yeates M: Muscle strength, physical fitness, and weight but not age predict femoral neck bone mass. J Bone Miner Res. 1989, 4: 441-448.

  8. 8.

    Sinaki M, McPhee MC, Hodgson SF, Merritt JM, Offord KP: Relationship between bone mineral density of spine and strength of back extensors in healthy postmenopausal women. Mayo Clin Proc. 1986, 61: 116-122.

  9. 9.

    Colletti LA, Edwards J, Gordon L, Shary J, Bell NH: The effects of muscle-building exercise on bone mineral density of the radius, spine, and hip in young men. Calcif Tissue Int. 1989, 45: 12-14. 10.1007/BF02556654.

  10. 10.

    Wronski TJ, Morey ER: Effect of spaceflight on periosteal bone formation in rats. Am J Physiol. 1983, 244: R305-9.

  11. 11.

    Donaldson CL, Hulley SB, Vogel JM, Hattner RS, Bayers JH, McMillan DE: Effect of prolonged bed rest on bone mineral. Metabolism. 1970, 19: 1071-1084. 10.1016/0026-0495(70)90032-6.

  12. 12.

    Wolff J: Concerning the interrelationship between form and function of the individual parts of the organism. By Julius Wolff, 1900. Clin Orthop Relat Res. 1988, 2-11.

  13. 13.

    Frost HM: The mechanostat: a proposed pathogenic mechanism of osteoporoses and the bone mass effects of mechanical and nonmechanical agents. Bone Miner. 1987, 2: 73-85.

  14. 14.

    Ozawa H, Imamura K, Abe E, Takahashi N, Hiraide T, Shibasaki Y, Fukuhara T, Suda T: Effect of a continuously applied compressive pressure on mouse osteoblast-like cells (MC3T3-E1) in vitro. J Cell Physiol. 1990, 142: 177-185. 10.1002/jcp.1041420122.

  15. 15.

    Imamura K, Ozawa H, Hiraide T, Takahashi N, Shibasaki Y, Fukuhara T, Suda T: Continuously applied compressive pressure induces bone resorption by a mechanism involving prostaglandin E2 synthesis. J Cell Physiol. 1990, 144: 222-228. 10.1002/jcp.1041440207.

  16. 16.

    Weyts FA, Bosmans B, Niesing R, van Leeuwen JP, Weinans H: Mechanical control of human osteoblast apoptosis and proliferation in relation to differentiation. Calcif Tissue Int. 2003, 72: 505-512. 10.1007/s00223-002-2027-0.

  17. 17.

    Jones DB, Nolte H, Scholubbers JG, Turner E, Veltel D: Biochemical signal transduction of mechanical strain in osteoblast-like cells. Biomaterials. 1991, 12: 101-110. 10.1016/0142-9612(91)90186-E.

  18. 18.

    Buckley MJ, Banes AJ, Levin LG, Sumpio BE, Sato M, Jordan R, Gilbert J, Link GW, Tran Son Tay R: Osteoblasts increase their rate of division and align in response to cyclic, mechanical tension in vitro. Bone Miner. 1988, 4: 225-236.

  19. 19.

    Buckley MJ, Banes AJ, Jordan RD: The effects of mechanical strain on osteoblasts in vitro. J Oral Maxillofac Surg. 1990, 48: 276-82; discussion 282-3.

  20. 20.

    McGarry JG, Klein-Nulend J, Mullender MG, Prendergast PJ: A comparison of strain and fluid shear stress in stimulating bone cell responses--a computational and experimental study. Faseb J. 2005, 19: 482-484.

  21. 21.

    Kapur S, Baylink DJ, Lau KH: Fluid flow shear stress stimulates human osteoblast proliferation and differentiation through multiple interacting and competing signal transduction pathways. Bone. 2003, 32: 241-251. 10.1016/S8756-3282(02)00979-1.

  22. 22.

    Reich KM, Gay CV, Frangos JA: Fluid shear stress as a mediator of osteoblast cyclic adenosine monophosphate production. J Cell Physiol. 1990, 143: 100-104. 10.1002/jcp.1041430113.

  23. 23.

    Charras GT, Williams BA, Sims SM, Horton MA: Estimating the sensitivity of mechanosensitive ion channels to membrane strain and tension. Biophys J. 2004, 87: 2870-2884. 10.1529/biophysj.104.040436.

  24. 24.

    Walker LM, Publicover SJ, Preston MR, Said Ahmed MA, El Haj AJ: Calcium-channel activation and matrix protein upregulation in bone cells in response to mechanical strain. J Cell Biochem. 2000, 79: 648-661. 10.1002/1097-4644(20001215)79:4<648::AID-JCB130>3.0.CO;2-Q.

  25. 25.

    Morris CE: Mechanosensitive ion channels. J Membr Biol. 1990, 113: 93-107. 10.1007/BF01872883.

  26. 26.

    Pavalko FM, Chen NX, Turner CH, Burr DB, Atkinson S, Hsieh YF, Qiu J, Duncan RL: Fluid shear-induced mechanical signaling in MC3T3-E1 osteoblasts requires cytoskeleton-integrin interactions. Am J Physiol. 1998, 275: C1591-601.

  27. 27.

    Salter DM, Robb JE, Wright MO: Electrophysiological responses of human bone cells to mechanical stimulation: evidence for specific integrin function in mechanotransduction. J Bone Miner Res. 1997, 12: 1133-1141. 10.1359/jbmr.1997.12.7.1133.

  28. 28.

    Wang N, Butler JP, Ingber DE: Mechanotransduction across the cell surface and through the cytoskeleton. Science. 1993, 260: 1124-1127. 10.1126/science.7684161.

  29. 29.

    Binderman I, Zor U, Kaye AM, Shimshoni Z, Harell A, Somjen D: The transduction of mechanical force into biochemical events in bone cells may involve activation of phospholipase A2. Calcif Tissue Int. 1988, 42: 261-266. 10.1007/BF02553753.

  30. 30.

    Jones D, Leivseth G, Tenbosch J: Mechano-reception in osteoblast-like cells. Biochem Cell Biol. 1995, 73: 525-534.

  31. 31.

    Hara F, Fukuda K, Ueno M, Hamanishi C, Tanaka S: Pertussis toxin-sensitive G proteins as mediators of stretch-induced decrease in nitric-oxide release of osteoblast-like cells. J Orthop Res. 1999, 17: 593-597. 10.1002/jor.1100170420.

  32. 32.

    Kuchan MJ, Jo H, Frangos JA: Role of G proteins in shear stress-mediated nitric oxide production by endothelial cells. Am J Physiol. 1994, 267: C753-8.

  33. 33.

    Kingsley DM: Genetic control of bone and joint formation. Novartis Found Symp. 2001, 232: 213-22; discussion 222-34, 272-82.

  34. 34.

    Ripamonti U, Teare J, Petit JC: Pleiotropism of bone morphogenetic proteins: from bone induction to cementogenesis and periodontal ligament regeneration. J Int Acad Periodontol. 2006, 8: 23-32.

  35. 35.

    Lawson KA, Dunn NR, Roelen BA, Zeinstra LM, Davis AM, Wright CV, Korving JP, Hogan BL: Bmp4 is required for the generation of primordial germ cells in the mouse embryo. Genes Dev. 1999, 13: 424-436. 10.1101/gad.13.4.424.

  36. 36.

    Solloway MJ, Robertson EJ: Early embryonic lethality in Bmp5;Bmp7 double mutant mice suggests functional redundancy within the 60A subgroup. Development. 1999, 126: 1753-1768.

  37. 37.

    Zhang H, Bradley A: Mice deficient for BMP2 are nonviable and have defects in amnion/chorion and cardiac development. Development. 1996, 122: 2977-2986.

  38. 38.

    Bandyopadhyay A, Tsuji K, Cox K, Harfe BD, Rosen V, Tabin CJ: Genetic analysis of the roles of BMP2, BMP4, and BMP7 in limb patterning and skeletogenesis. PLoS Genet. 2006, 2: e216-10.1371/journal.pgen.0020216.

  39. 39.

    King JA, Storm EE, Marker PC, Dileone RJ, Kingsley DM: The role of BMPs and GDFs in development of region-specific skeletal structures. Ann N Y Acad Sci. 1996, 785: 70-79.

  40. 40.

    Storm EE, Kingsley DM: Joint patterning defects caused by single and double mutations in members of the bone morphogenetic protein (BMP) family. Development. 1996, 122: 3969-3979.

  41. 41.

    Kingsley DM, Bland AE, Grubber JM, Marker PC, Russell LB, Copeland NG, Jenkins NA: The mouse short ear skeletal morphogenesis locus is associated with defects in a bone morphogenetic member of the TGF beta superfamily. Cell. 1992, 71: 399-410. 10.1016/0092-8674(92)90510-J.

  42. 42.

    King JA, Marker PC, Seung KJ, Kingsley DM: BMP5 and the molecular, skeletal, and soft-tissue alterations in short ear mice. Dev Biol. 1994, 166: 112-122. 10.1006/dbio.1994.1300.

  43. 43.

    Dubois CM, Blanchette F, Laprise MH, Leduc R, Grondin F, Seidah NG: Evidence that furin is an authentic transforming growth factor-beta1-converting enzyme. Am J Pathol. 2001, 158: 305-316.

  44. 44.

    Constam DB, Robertson EJ: Regulation of bone morphogenetic protein activity by pro domains and proprotein convertases. J Cell Biol. 1999, 144: 139-149. 10.1083/jcb.144.1.139.

  45. 45.

    Eimon PM, Harland RM: Effects of heterodimerization and proteolytic processing on Derriere and Nodal activity: implications for mesoderm induction in Xenopus. Development. 2002, 129: 3089-3103.

  46. 46.

    Hawley SH, Wunnenberg-Stapleton K, Hashimoto C, Laurent MN, Watabe T, Blumberg BW, Cho KW: Disruption of BMP signals in embryonic Xenopus ectoderm leads to direct neural induction. Genes Dev. 1995, 9: 2923-2935. 10.1101/gad.9.23.2923.

  47. 47.

    Zhu X, Hadhazy M, Wehling M, Tidball JG, McNally EM: Dominant negative myostatin produces hypertrophy without hyperplasia in muscle. FEBS Lett. 2000, 474: 71-75. 10.1016/S0014-5793(00)01570-2.

  48. 48.

    Wittbrodt J, Rosa FM: Disruption of mesoderm and axis formation in fish by ectopic expression of activin variants: the role of maternal activin. Genes Dev. 1994, 8: 1448-1462. 10.1101/gad.8.12.1448.

  49. 49.

    Felts WJ: In vivo implantation as a technique in skeletal biology. Int Rev Cytol. 1961, 12: 243-302.

  50. 50.

    Rudnicki MA, Schnegelsberg PN, Stead RH, Braun T, Arnold HH, Jaenisch R: MyoD or Myf-5 is required for the formation of skeletal muscle. Cell. 1993, 75: 1351-1359. 10.1016/0092-8674(93)90621-V.

  51. 51.

    Hamrick MW, McPherron AC, Lovejoy CO: Bone mineral content and density in the humerus of adult myostatin-deficient mice. Calcif Tissue Int. 2002, 71: 63-68. 10.1007/s00223-001-1109-8.

  52. 52.

    Derynck R, Zhang YE: Smad-dependent and Smad-independent pathways in TGF-beta family signalling. Nature. 2003, 425: 577-584. 10.1038/nature02006.

  53. 53.

    Balemans W, Van Hul W: Extracellular regulation of BMP signaling in vertebrates: a cocktail of modulators. Dev Biol. 2002, 250: 231-250.

  54. 54.

    DiLeone RJ, Marcus GA, Johnson MD, Kingsley DM: Efficient studies of long-distance Bmp5 gene regulation using bacterial artificial chromosomes. Proc Natl Acad Sci U S A. 2000, 97: 1612-1617. 10.1073/pnas.97.4.1612.

  55. 55.

    DiLeone RJ, Russell LB, Kingsley DM: An extensive 3' regulatory region controls expression of Bmp5 in specific anatomical structures of the mouse embryo. Genetics. 1998, 148: 401-408.

  56. 56.

    Ikegame M, Ishibashi O, Yoshizawa T, Shimomura J, Komori T, Ozawa H, Kawashima H: Tensile stress induces bone morphogenetic protein 4 in preosteoblastic and fibroblastic cells, which later differentiate into osteoblasts leading to osteogenesis in the mouse calvariae in organ culture. J Bone Miner Res. 2001, 16: 24-32. 10.1359/jbmr.2001.16.1.24.

  57. 57.

    Wu Q, Zhang Y, Chen Q: Indian hedgehog is an essential component of mechanotransduction complex to stimulate chondrocyte proliferation. J Biol Chem. 2001, 276: 35290-35296. 10.1074/jbc.M101055200.

  58. 58.

    Green MC: A rapid method for clearing and staining specimens for the demonstration of bone. Ohio J Sci. 1952, 52: 31-33.

  59. 59.

    Marker PC, King JA, Copeland NG, Jenkins NA, Kingsley DM: Chromosomal localization, embryonic expression, and imprinting tests for Bmp7 on distal mouse chromosome 2. Genomics. 1995, 28: 576-580. 10.1006/geno.1995.1192.

  60. 60.

    Ho AM, Johnson MD, Kingsley DM: Role of the mouse ank gene in control of tissue calcification and arthritis. Science. 2000, 289: 265-270. 10.1126/science.289.5477.265.

Download references

Acknowledgements

Genetic and biochemical analyses of the Bmp5clvmutants were performed in the Kingsley lab, and biomechanical studies were carried out in the Huard lab. We thank M. Wagener, M. Johnson, and L. Pantalena-Filho for excellent animal care, K. Corsi, D.W. Humiston, and R. Sauder for manuscript editing, and members of the Kingsley and Huard labs for discussions. This work was supported by 5T32GM07365 (NIGMS) and a Pittsburgh Foundation Albert Ferguson grant to AMH, NIH DE 13420-03 and the Henry J. Mankin and William F. and Jean W. Donaldson Chairs to JH, and NIH AR42236 and a Howard Hughes Medical investigatorship to DMK.

Author information

Correspondence to David M Kingsley.

Additional information

Authors' contributions

AMH and PCM carried out the genetic sequencing of the Bmp5clvallele. AMH performed the genetic, biochemical, and biomechanical studies on the mutant. DMK and AMH contributed to the conception, design, analysis, and interpretation of the genetic and biochemical analyses. AMH, AJQ, HP, and JH participated in the conception, design, analysis, and interpretation of the biomechanical studies. All authors were involved in the drafting, review, and revision of the manuscript, and have read and agreed to its final content.

Authors’ original submitted files for images

Rights and permissions

Reprints and Permissions

About this article

Keywords

  • Mechanical Stimulation
  • Mechanical Strain
  • Deltoid Muscle
  • Skeletal Defect
  • BMP5 Protein