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Article |
Address correspondence to Noriyuki Tsumaki, Dept. of Orthopaedics, Osaka University Graduate School of Medicine, 2-2 Yamadaoka, Suita, Osaka 565-0871, Japan. Tel.: 81-6-6879-3552. Fax: 81-6-6879-3559. email: tsumaki-n{at}umin.ac.jp
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Abstract |
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Key Words: chondrocyte; hypertrophy; osteopenia; dwarfism; transgenic mice
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Introduction |
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Members of the TGF-ß superfamily transduce their signals through two types of serine/threonine kinase receptors, types I and II (Heldin et al., 1997; Shi and Massague, 2003). Upon ligand binding, type I and type II receptors form a tetramer consisting of two pairs of type I and type II receptors. Type II receptors phosphorylate type I receptors. Then, type I receptors phosphorylate downstream targets such as Smads. In vertebrates, seven type I receptors and five type II receptors have been found so far. Among them, three type I receptors, BMP type IA receptor (BMPR-IA, or activin receptor-like kinase [ALK-3]), BMPR-IB (ALK-6), and ALK-2, mediate BMP signaling. Smads are the major downstream targets of type I receptors for TGF-ß/BMP superfamily proteins (Heldin et al., 1997). Eight Smads have been identified in mammals and are classified into three subgroups. Receptor-regulated Smads (R-Smads) are phosphorylated at SSXS motifs at their COOH terminus by type I receptors. Smad1, Smad5, and Smad8 are R-Smads that transduce BMP signals, and Smad2 and Smad3 are responsible for TGF-ß and activin signaling. Phosphorylated R-Smads form heteromers with the common-partner Smad (Smad4), and translocate into the nucleus where they interact with transcriptional factors to bind directly or indirectly to specific DNA sequences for the activation of gene transcription.
BMP signaling is subject to delicate regulation at multiple levels: extracellularly, at the membrane site, and intracellularly (Balemans and Van Hul, 2002). In the extracellular space, several molecules antagonize BMPs. Among these antagonists, noggin is expressed in cartilage and binds to BMPs and prevents them from interacting with their receptors. At the intracellular level, inhibitory Smads, Smad6 and Smad7, inhibit phosphorylation of R-Smads by competing with R-Smads for binding to phosphorylated type I receptors. In particular, Smad6 appears to inhibit BMP signaling, whereas Smad7 associates stably with TGF-ß receptor and BMP receptor complexes and inhibits the TGF-ß or BMP-mediated phosphorylation of R-Smads (Hanyu et al., 2001). In addition, ubiquitin-dependent protein degradation plays key roles in Smad signaling. Smad ubiquitin regulatory factor 1 (Smurf1) and Smurf2 induce the ubiquitination and degradation of Smad1 and Smad5 (Zhu et al., 1999; Zhang et al., 2001). Furthermore, Smurf1 and Smurf2 interact with nuclear Smad7 and induce the nuclear export of Smad7. SmurfSmad7 complexes then associate with type I receptor for TGF-ß and enhance its turnover (Kavsak et al., 2000; Ebisawa et al., 2001). Recent biochemical analyses have shown that Smurf1 binds to BMP type I receptors via Smad6 and Smad7, and that it induces the ubiquitination and degradation of these receptors (Murakami et al., 2003). Thus, Smad6 and Smurf1 cooperatively down-regulate BMP signals by degradation of R-Smads as well as BMP receptors. However, the physiological function of Smurfs is unknown.
During development, the limb skeleton is formed through endochondral bone formation (Erlebacher et al., 1995). Mesenchymal cells initially undergo condensation followed by differentiation of cells within these condensations into chondrocytes. Chondrocytes then proliferate and produce ECM to form primordial cartilage. Shortly after the primordial cartilage formation, proliferating chondrocytes in the central region of the cartilage undergo terminal differentiation to hypertrophic chondrocytes. Hypertrophic chondrocytes exit the cell cycle and synthesize an ECM that is different in composition from that of proliferating cartilage. The hypertrophic cartilage is invaded by blood vessels along with osteoblasts, osteoclasts, and hematopoietic cells to form primary ossification centers. Within these centers, the hypertrophic cartilage matrix is degraded, hypertrophic chondrocytes die, and osteoblasts replace the disappearing cartilage with trabecular bone (Olsen et al., 2000). Then, bone formation and maintenance are performed by a balance between the new apposition of bony matrix by osteoblasts and resorption by osteoclasts.
Smad proteins have been identified in growth plate cartilage (Flanders et al., 2001). In vitro analyses have shown that Smad6 regulates the chondrocytic phenotype (Valcourt et al., 2002; Nishihara et al., 2003). However, the physiological roles of Smad6 and Smad signaling in normal endochondral bone formation have not been determined. Here, we generated transgenic mice overexpressing Smad6 or Smurf1 in chondrocytes under the control of the 2(XI) collagen chain gene (Col11a2) promoter/enhancer sequences. We found that overexpression of Smad6 does not significantly affect chondrocyte proliferation, but significantly delays chondrocyte hypertrophy, which may lead to postnatal dwarfism with osteopenia. By using double-transgenic mice, we also found that Smurf1 supports Smad6 function in vivo.
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Results |
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Cartilage-specific expression of transgene in Smad6 transgenic mice
Northern blotting demonstrated Smad6 expression in the limb buds of transgenic mice. Transgene Smad6 mRNA was 2 kb in length, which was smaller than endogenous Smad6 mRNA due to shorter 5' and 3' untranslated regions (Fig. 2 A). Immunoblotting demonstrated the expression of a 70-kD FLAG-tagged Smad6 protein in limb buds of Smad6 transgenic mice (Fig. 2 B, open arrow). Immunohistochemistry using anti-Smad6 antibody showed more intense signals for Smad6 proteins in forelimb chondrocytes of Smad6 transgenic mice (Fig. 2, D, F, H, and J) than from wild-type mice (Fig. 2, C, E, G, and I) from 13.0 through 18.5 days post coitus (d.p.c.).
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To elucidate the mechanism by which osteoclastic bone resorption was activated in Smad6 transgenic mice, we performed bone marrow cell culture and analyzed dexamethasone/parathyroid hormone-induced osteoclastogenesis. The number of tartrate-resistant acid phosphatase (TRAP)positive multinucleated cells in the culture prepared from transgenic mice was equivalent to that of normal mice (Fig. 4, PR). The resorption of hydroxyapatite by cultured osteoclast prepared from transgenic mice was essentially normal (Fig. 4 S). Next, we examined osteoclast formation activities in spleen cell culture in the presence of RANK ligand. RANK ligandinduced osteoclastogenesis of spleen cells from Smad6 transgenic mice was normal, as indicated by the number of TRAP-positive multinucleated cells and the resorption of hydroxyapatite (unpublished data). These results suggested that both osteoclast precursors and osteoclast-supporting activities of osteoblast/stromal cells were normal in the bone marrow of Smad6 transgenic mice.
Skeletal development of Smad6 transgenic mice
Because the transgene was expressed specifically in cartilage, we analyzed the skeleton from earlier stages of development. Whole-mount in situ hybridization using a type II collagen gene (Col2a1) antisense cRNA probe showed that the pattern and intensity of signals did not obviously differ between Smad6 transgenic and normal mice at either 12.5 (Fig. 5, A and B) or 13.0 (Fig. 5, C and D) d.p.c. This suggested that mesenchymal condensation was essentially normal in the transgenic mice. At 13.5 d.p.c., the size and shape of each cartilaginous skeletal component stained with Alcian blue of transgenic mice were essentially identical to those of normal littermates (Fig. 5, E and F). The central regions of metatarsals were mineralized in normal mice as they were stained with Alizarin red S, but not in transgenic mice at 16.5 d.p.c. (Fig. 5, G and H, arrowheads) and at 18.5 d.p.c. (Fig. 5, I and J, arrowheads). The size of mineralized tissue in the humerus and femur was considerably smaller in transgenic mice than in normal mice (Fig. 5, G and H, arrows). At 3 wk of age, skeleton of transgenic mice was much smaller than that of normal mice (Fig. 5, K and L).
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Chondrocyte hypertrophy induced by BMP2 was down-regulated in cartilage explants prepared from Smad6 transgenic mice
To investigate BMP signaling in chondrocytes, we organ cultured primordial metatarsal cartilage at 15.0 d.p.c. Phase-contrast microscopy could distinguish the zones of proliferative and mineralized hypertrophic cartilage in the cultures (Fig. 8 A), which was confirmed by histology (Fig. 8, B and C). At the start of culture, metatarsal rudiments from Smad6 transgenic mice were indistinguishable from those of normal mice (Fig. 8, D and E). After 4 d of culture in control medium, the length of hypertrophic cartilage in normal rudiments was increased (Fig. 8 F), whereas transgenic rudiments lacked hypertrophic cartilage (Fig. 8 G). Culture in the presence of rhBMP2 resulted in excessive outgrowth of the proliferative cartilage at both ends of the rudiments and enhanced formation of a hypertrophic center in the normal rudiments (Fig. 8 H). In transgenic rudiments incubated with rhBMP2, proliferative cartilage expanded (Fig. 8 I) like that of normal mice (Fig. 8 H), but formation of the hypertrophic center was limited (Fig. 8 I) compared with that of normal mice (Fig. 8 H). These findings were confirmed by morphometric analysis of the rudiments (Fig. 8, J and K), indicating that Smad6 overexpression inhibited chondrocyte hypertrophy induced by rhBMP2.
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Discussion |
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Smad6 appears to block BMP signaling, whereas Smad7 blocks that of both TGF-ß and BMP (Hanyu et al., 2001). Skeletal mineralization is delayed and skeletal patterning is defective in mice lacking growth and differentiation factor 5 (Storm et al., 1994), BMPR-IB (Yi et al., 2000), or BMP7 (Luo et al., 1995; Jena et al., 1997). The similarity of delayed mineralization between these mutant mice and Smad6 transgenic mice suggests that BMP signals are blocked by Smad6 during endochondral ossification. This notion was confirmed by our results that chondrocyte hypertrophy and mineralization induced by rhBMP2 was inhibited in cartilage explants from Smad6 transgenic mice. Because the phosphorylation of Smad1/5/8 induced by rhBMP2 was inhibited in explants from Smad6 transgenic mice, we concluded that Smad6 regulates chondrocyte hypertrophy through the inhibition of Smad1/5/8 phosphorylation, thus down-regulating BMP signaling in endochondral bone formation.
The most fundamental abnormality during chondrocyte differentiation in Smad6 transgenic mice was a delay in chondrocyte hypertrophy in humeri at 14.5 d.p.c. This delay was accompanied by the persistent expression of the Col2a1 gene and the retarded expression of the type X collagen gene (Col10a1). Extensive analyses in vitro have shown that BMP signals promote chondrocyte hypertrophy, and BMP-responsive cis-acting elements have been identified in the promoter sequence of the Col10a1 gene (Volk et al., 1998; Drissi et al., 2003). These in vitro analyses and our in vivo results collectively suggest that Smad6 overexpression blocks BMP signaling, thus preventing transcriptional activation of the Col10a1 gene.
Smad6 regulates endochondral ossification in cooperation with Smurf1
Smurf1 binds Smads 1 and 5 and promotes their degradation (Zhu et al., 1999). Smurf1 and Smad6 form complexes and inhibit BMP signaling through the ubiquitin-dependent degradation of BMP receptors as well as of R-Smads (Murakami et al., 2003). Smurf2 may also exhibit functions similar to Smurf1. To examine the in vivo function of Smurf1, we generated transgenic mice expressing Smurf1 in chondrocytes and did not find obvious abnormalities. These results suggest that sufficient Smurf1 already exists in normal chondrocytes. When apparently normal Smurf1 transgenic mice were mated with Smad6 transgenic mice, the endochondral ossification of progenies overexpressing both Smad6 and Smurf1 was more delayed than in transgenic mice overexpressing only Smad6. It is likely that the Smad6 transgenic mice have far less Smurf1/2 than Smad6. In Smad6/Smurf1 double-transgenic mice, Smurf1 derived from the transgene might compensate for this shortage, thus supporting the activities of a large amount of Smad6. When Smurf1 transgenic mice were mated with those of the Smad6 transgenic line 165 in which the expression level was low, the phenotypic severity of the resultant double-transgenic progeny did not differ from those of Smad6 transgenic mice of line 165 (unpublished data). In Smad6 transgenic mice of this line, endogenous Smurf1/2 fully supported the activities of endogenous Smad6 and that derived from the transgene. From these lines of discussion, the expression level of Smad6 appears to be critical in the regulation of conversion from proliferative chondrocytes to hypertrophic chondrocytes. Actually, the expression level of Smad6 is decreased in the transitional zone between proliferative chondrocytes and hypertrophic chondrocytes (Flanders et al., 2001), suggesting that critical regulation of Smad6 expression is responsible for this conversion. The expression level of inhibitory Smads seems to be consistently and strictly regulated through autoregulatory negative feedback during signal transduction of the TGF-ß/BMP superfamily because the inhibitory Smad mRNA is induced by TGF-ß stimulation (Heldin et al., 1997).
Postnatal dwarfism with osteopenia might be associated with the reduced zone of hypertrophic chondrocytes
The most apparent phenotype of Smad6 transgenic mice was postnatal dwarfism and osteopenia. Dynamic bone histomorphometric analysis revealed that osteoblastic bone formation decreased and that osteoclastic bone resorption increased in Smad6 transgenic mice. However, results from cultured bone marrow cells suggested normal osteoclast-supporting activities of osteoblasts/stromal cells prepared from Smad6 transgenic mice. These in vitro results suggest that the increased in vivo osteoclastic bone resorption in transgenic mice was not due to an autonomous abnormality within bone marrow cells. This notion was consistent with the observation that the transgene was specifically expressed in chondrocytes. We speculate that abnormal activities of osteoblasts and osteoclasts might be associated with the dysfunction in cartilage during endochondral bone formation. However, we could not rigorously exclude leaky transgene expression in cells in osteoblast lineage, and thus the possibility that osteoblasts have a primary malfunction.
BrdU labeling revealed that proliferation of chondrocytes in Smad6 transgenic mice was normal at the embryonic stage and at the postnatal stage. Therefore, postnatal dwarfism in Smad6 transgenic mice might develop through a different mechanism from that in transgenic mice overexpressing activated FGF receptor 3 in cartilage, in which chondrocyte proliferation is normal in embryos but postnatally decreased (Naski et al., 1998).
The onset of chondrocyte hypertrophy was delayed by 3 d during the development of Smad6 transgenic mice. Once the zone of hypertrophic chondrocytes formed, the height of the zone of hypertrophic chondrocytes was essentially normal, suggesting that the population of hypertrophic chondrocytes is strictly regulated at late embryonic stages. We did not determine which mechanism maintains the population of hypertrophic chondrocytes in Smad6 transgenic embryos. However, this population decreased in the transgenic mice after birth. It is likely that reduced hypertrophic chondrocyte population lead to a deficiency in signals required for coordination of growth and bone formation after birth. Hypertrophic chondrocytes are known to produce various factors, including angiogenic factors (Karsenty and Wagner, 2002). A reduction in production of such factors may well lead to dysfunction of osteoblast/osteoclast activities, thus resulting in impaired bone growth and osteopenia.
It is also possible that abnormal chondrocyte hypertrophy could result in the impaired structure of the ECM of hypertrophic cartilage. An impaired matrix might not provide a suitable scaffold for osteoblasts and osteoclasts to replace cartilage with bone. This speculation remains to be examined.
Chondrocyte proliferation was not affected in Smad6 transgenic mice
Previously, we generated transgenic mice expressing noggin in chondrocytes under the control of the identical Col11a2 promoter/enhancer sequences used in this paper (Tsumaki et al., 2002). We examined 62 G0 founder mouse embryos for the noggin transgene, and 7 of them displayed a severe phenotype and almost completely lacked cartilage formation during development. On the other hand, 17 of 129 G0 embryos were genetically positive for the Smad6 transgene, and skeletal abnormalities were either relatively minor or almost absent. Although we could not exclude the possibility that the phenotypic difference between noggin and Smad6 transgenic mice might be related to the different level of expression of the transgenes, these results suggest that noggin tends to affect cartilage more than Smad6 when expressed in chondrocytes in transgenic mice.
In addition, chondrocyte proliferation is inhibited in mice lacking BMPR-IB (Yi et al., 2000) and in cartilage explants cultured in the presence of noggin (Minina et al., 2001). Reports have consistently indicated that BMP signaling stimulates chondrocyte proliferation during endochondral bone formation at the embryonic stages of transgenic mice overexpressing BMPs in cartilage (Tsumaki et al., 1999, 2002) and in organ culture of cartilage rudiments in the presence of BMPs (De Luca et al., 2001; Minina et al., 2001).
On the other hand, chondrocyte proliferation appeared normal in Smad6 transgenic mice, as indicated by BrdU labeling. For explanation of the discrepancy between the findings of chondrocyte proliferation obtained from Smad6 transgenic mice and those of other papers, we considered four possibilities. First, we could not exclude the possibility that the expression level of Smad6 transgene was not sufficient to block BMP signaling completely, although immunohistochemical analysis showed strong expression of the transgene (Fig. 2, CJ) Second, Smad signaling might not be blocked by Smad6 alone. Certainly, phenotypes of Smad6/Smurf1 double-transgenic mice were more severe than that of Smad6 transgenic mice. However, it is still milder than that of noggin transgenic mice, and chondrocyte proliferation remained normal, as indicated by BrdU labeling. Third, BMP signals might be mediated by signaling pathways other than Smad proteins. In certain cell types, various MAPKs have been reported to mediate BMP pathways (Iwasaki et al., 1999). Existence of such pathways might account for the discrepancy between chondrocyte proliferation in BMPR-IBdeficient mice and Smad6 transgenic mice. There is likelihood of the fourth possibility as follows: BMPs and noggin are secreted and diffuse. In addition to direct binding to chondrocytes, these proteins might exert indirect effects on chondrocytes. For example, BMPs act on cells around cartilage, and in return these cells secrete factors, affecting chondrocyte proliferation. Thus, the addition or overexpression of BMPs/noggin modulate chondrocyte proliferation directly and indirectly. On the other hand, Smad6 overexpression in chondrocytes might block only the direct effect of BMPs on chondrocytes.
In conclusion, our data on inhibition of endochondral bone formation in Smad6 and Smurf1 transgenic mice suggest a role for Smad signaling in skeletogenesis and growth. By down-regulating Smad1/5/8 phosphorylation and BMP signals, Smad6 plays an important role in regulation of chondrocyte hypertrophy and synergistically cooperates with Smurf1 in vivo.
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Materials and methods |
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Generation of transgenic mice
The plasmids Col11a2-Smad6 and Col11a2-Smurf1 were digested with EcoRI and PstI to release the inserts. Transgenic mice were produced by microinjecting each of the inserts into the pronuclei of fertilized eggs from F1 hybrid mice (C57BL/6x DBA) as described previously (Tsumaki et al., 1996). Transgenic embryos were identified by PCR assays of genomic DNA extracted from the placenta or skin. Genomic DNA was amplified by transgene-specific PCR using primers derived from mouse Smad6 cDNA (5'-CAAGATCGGTTTTGGCATACTG-3') and from the SV40 poly(A) signal region (5'-TCACTGCATTCTAGTTGTGGTTTGTCC-3') to amplify a 411-bp product for Smad6 transgenic mice. To discriminate Smurf1 transgenic mice, genomic DNA was amplified by transgene-specific PCR using primers derived from mouse Smurf1 cDNA (5'-ATGGACTACAAGGACGATGATGACAAGG-3' and 5'-AGGGGCTGGTTCCTCCATGAAGCAG-3') to amplify a 570-bp product. Smad6/Smurf1 double-transgenic pups were generated by mating Smad6 and Smurf1 transgenic mice.
Staining of the skeleton
Mice were dissected and fixed in 100% ethanol overnight, and then stained with Alcian blue followed by Alizarin red S solution according to standard protocols (Peters, 1977).
Micro CT analysis and bone mass measurement
The humeri from 5-wk-old transgenic mice and normal littermates were dissected and analyzed using a micro-focus X-ray CT system (SMX-100CT-SV; Shimadzu). The region extending from the proximal growth plate to the metaphyseal part of the humerus on 350 slices was scanned at a width of 6.75 µm per slice. The data were reconstructed to produce images of the humerus using 3-D visualization and measurement software (Vay Tek, Inc.). Bone mass was quantified by selecting 90 consecutive slices distal to the proximal growth plate (0.6 mm in length). Trabecular parameters in the metaphysis were determined using image analysis software (TRI/3D-BON; RATOC).
Histology and immunohistochemical staining
Embryos were dissected using a stereomicroscope (model SMZ645; Nikon), fixed in 4% PFA, processed, and embedded in paraffin. Serial sections were stained with hematoxylin and eosin, with safranin O/fast green/iron hematoxylin, or by the von Kossa reaction. Dynamic histomorphometric indices were determined by double-fluorescence labeling in vertebral bodies. 4-wk-old normal and transgenic mice were administered i.p. with tetracycline (20 mg/kg body weight; Sigma-Aldrich), followed by calcein label (10 mg/kg body weight; Wako Chemicals) 2 d later. After 24 h, the mice were killed. Bones were fixed with ethanol and embedded in methylmethacrylate. Sections were cut and viewed using a fluorescence microscope (Eclipse E1000; Nikon). The Niigata Bone Science Institute (Niigata, Japan) performed histomorphometric analyses. Immunohistochemistry proceeded using a rabbit pAb against Smad6 (1:200 dilution; Zymed Laboratories) and a rabbit pAb against phospho-Smad1/5/8 (1:200 dilution; Cell Signaling Technology). Immune complexes were detected using streptavidin-peroxidase staining and Histofine SAB-PO kits (Nichirei). Images were acquired using a microscope (Eclipse E1000; Nikon) with a digital camera system (DXM1200; Nikon).
BrdU staining
Pregnant mice bearing 16.5 d.p.c. embryos and 3-wk-old mice were i.p. injected with BrdU labeling reagent (10 µl/g body weight; Zymed Laboratories). 2 h later, the mice were killed. Embryonic limb buds and tibia of the 3-wk-old mice were dissected and sectioned. Incorporated BrdU was detected using a BrdU staining kit (Zymed Laboratories) to distinguish actively proliferating cells. Tissue sections were measured using a micrometer, and the average number of BrdU-positive cells/mm2 cartilage ± SD was calculated.
Northern hybridization and real-time quantitative RT-PCR
Total RNA extracted from the limb buds of 14.518.5 d.p.c. transgenic and normal embryos using RNeasy Mini Kits (QIAGEN) was fractionated by electrophoresis through formaldehyde agarose gels and transferred onto Nytran® membranes (Schleicher & Schuell Bioscience). Complementary DNAs (cDNAs) were labeled with [32P]dCTP using Prime-it® II kits (Stratagene). The membranes were hybridized with 32P-labeled Smad6 cDNA and rehybridized with 32P-labeled probes for mouse 1(II) collagen,
1(IX) collagen, and Sox9.
Total RNAs were digested with DNase to eliminate any contaminating genomic DNA before real-time quantitative RT-PCR. 2 µg of total RNA was reverse transcribed into first-strand cDNA using OmniScript® reverse transcriptase (QIAGEN) and an oligo(dT)12-18 primer. The PCR amplification proceeded in 20 µl containing 1 µl of cDNA, 2 µl of SYBER GreenTM Master Mix (QIAGEN), and 10 pmol of primers specific for Smad6 (5'-GATCCCCAAGCCAGACAGT-3' and 5'-AGCCTCTTGAGCAGCGCGAGTA-3') to generate a 126-bp product (GenBank/EMBL/DDBJ accession no. NM008542). The cDNA was amplified by 35 cycles using a LightCycler® quick system (Roche) according to the following protocol: 94°C for 15 s, 60°C for 20 s, and 72°C for 6 s, each with a temperature transition rate of 20°C, according to the manufacturer's instructions.
In situ hybridization
Digoxigenin-11 UTP-labeled single-strand RNA probes were prepared using a DIG RNA labeling kit (Boehringer) according to the manufacturer's instructions. We generated antisense and sense probes using 1(II) collagen,
1(X) collagen, and osteopontin cDNAs. Hybridization proceeded as described previously (Hirota et al., 1992; Conlon and Herrmann, 1993). A Genius detection system (Boehringer) detected signals according to the manufacturer's instructions.
Immunoprecipitation and Western blotting
Limb buds of 14.5 d.p.c transgenic and normal embryos were lysed with RIPA buffer (50 mM Tris, pH 7.4, 150 mM NaCl, 1% Nonidet P-40, 0.5% sodium deoxycholate, 0.1% SDS, and 10 mM DTT) supplemented with protease and phosphatase inhibitor cocktails (Sigma-Aldrich). The positive control consisted of COS7 cells transfected with the expression construct FLAG-Smad6 that was also lysed with RIPA buffer. The cell lysates were incubated with anti-Smad6 antibody (Zymed Laboratories) for 3 h at 4°C followed by an incubation with protein Gagarose beads (Roche) for 3 h at 4°C. After five washes with lysis buffer (20 mM Hepes, 150 mM NaCl, 10% glycerol, 1.5 mM MgCl2, 1 mM EGTA, and 100 µM orthovanadate), 1x SDS sample buffer was added to the agarose beads. The samples were incubated for 5 min at 95°C, fractionated by 10% SDS-PAGE, transferred onto nitrocellulose membranes (Bio-Rad Laboratories), and Western blotted against anti-FLAG M2 mAb (Sigma-Aldrich). Immunocomplex bands were visualized using the ECL Western blotting detection system (Amersham Biosciences).
Bone marrow cell culture
Osteoclast formation and bone resorbing activity were determined using the modified method described in Azuma et al. (2000). In brief, bone marrow cells prepared from the femurs and tibias of 10-wk-old transgenic mice or wild-type control mice were suspended in -modified essential medium containing 10% FBS, and were cultured in 48-well plates (106 cells/0.5 ml per well) for 7 d in the presence of 0.1 µM dexamethasone (Sigma-Aldrich) and 0.01 µM recombinant human parathyroid hormone (Peptide Institute, Inc.). Cells were then fixed and stained for TRAP using a TRAP staining kit (Hokudo) according to the manufacturer's recommendation. The number of multinucleated TRAP-positive cells with more than three nuclei was counted under a microscope (Eclipse TE300; Nikon). To examine calcified matrix resorption activity, 5 x 105 cells were cultured in 16-well hydroxyapatite-coated slides (Osteologic; Becton Dickinson) for 14 d, and the resorption area was calculated by computer-assisted image analysis.
Metatarsal explant culture
Metatarsal rudiments were cultured as described previously (Haaijman et al., 1997). Metatarsal rudiments were dissected from transgenic and normal embryos at 15.0 d.p.c. and cultured in -modified essential medium without nucleosides (Invitrogen), supplemented with 0.05 mg/ml ascorbic acid (Sigma-Aldrich), 0.3 mg/ml L-glutamine (Merck), 0.05 mg/ml gentamycin (Invitrogen), 0.25 mg/ml Fungizone® (Invitrogen), 1 mM ß-glycerophosphate (Merck), and 0.2% FBS (GIBCO BRL) in a humidified atmosphere of 5% CO2 in air at 37°C. 1 d after starting the cultures, the rudiments were incubated in 400 µl of the same medium containing 500 ng/ml of rhBMP2 (Yamanouchi Pharmaceutical Co., Ltd.) or without rhBMP2 for a further 3 d. For immunohistochemical analysis using anti-phosphorylated Smad1/5/8, rudiments were sectioned before and after a 2-h incubation with rhBMP2. Images of rudiments obtained under inverted phase-contrast microscopy were analyzed morphometrically (Eclipse TE300; Nikon). Areas of proliferative cartilage and length of hypertrophic cartilage were measured using NIH Image software (National Institutes of Health, Bethesda, MD).
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Acknowledgments |
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This work was supported in part by Scientific Research grant 15390458 from the Ministry of Education, Science and Culture of Japan; Health and Labor Sciences Research Grants of Japan; and the Grant of Japan Orthopaedic and Traumatology Foundation, Inc. (0126).
Submitted: 4 November 2003
Accepted: 30 March 2004
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