Thyroid Hormones Regulate Fibroblast Growth Factor Receptor ...

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Endocrinology 146(12):5568 –5580 Copyright © 2005 by The Endocrine Society doi: 10.1210/en.2005-0762

Thyroid Hormones Regulate Fibroblast Growth Factor Receptor Signaling during Chondrogenesis Joanna C. Barnard, Allan J. Williams, Be´ne´dicte Rabier, Olivier Chassande, Jacques Samarut, Sheue-yann Cheng, J. H. Duncan Bassett, and Graham R. Williams Molecular Endocrinology Group (J.C.B., A.J.W., J.H.D.B., G.R.W.), Division of Medicine and Medical Research Council Clinical Sciences Centre, Imperial College London, Hammersmith Campus, London W12 0NN, United Kingdom; Institut National de la Sante´ et de la Recherche Me´dicale Unite´ 443 (B.R., O.C.), Universite´ Victor Segalen Bordeaux 2, Bordeaux, France; Laboratoire de Biologie Mole´culaire et Cellulaire de l’Ecole Normale Supe´rieur de Lyon (J.S.), Unite´ Mixte de Recherche 5665 Centre National de la Recherche Scientifique, LA 913 Institut National de la Recherche Agronomique, Lyon, France; and Gene Regulation Section (S.-y.C.), Laboratory of Molecular Biology, National Cancer Institute, National Institutes of Health, Bethesda, Maryland 20892-4264 Childhood hypothyroidism causes growth arrest with delayed ossification and growth-plate dysgenesis, whereas thyrotoxicosis accelerates ossification and growth. Thyroid hormone (T3) regulates chondrocyte proliferation and is essential for hypertrophic differentiation. Fibroblast growth factors (FGFs) are also important regulators of chondrocyte proliferation and differentiation, and activating mutations of FGF receptor-3 (FGFR3) cause achondroplasia. We investigated the hypothesis that T3 regulates chondrogenesis via FGFR3 in ATDC5 cells, which undergo a defined program of chondrogenesis. ATDC5 cells expressed two FGFR1, four FGFR2, and one FGFR3 mRNA splice variants throughout chondrogenesis, and expression of each isoform was stimulated by T3 during the first 6 –12 d of culture, when T3 inhibited proliferation by 50%. FGFR3 expression was also increased in cells treated with T3 for 21 d, when T3 induced an earlier onset of hypertrophic differentiation and collagen X expression. FGFR3 ex-

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HYROID HORMONE (T3) is a key regulator of skeletal development and growth (1, 2). Hypothyroidism in childhood causes growth retardation, delayed bone age, and epiphyseal dysgenesis. Animal studies indicate that these features are due to a lack of the normal effects of thyroid hormones on chondrocyte proliferation and due to impaired hypertrophic chondrocyte differentiation during endochondral ossification. There is also impaired intramembranous ossification resulting in delayed closure of the skull sutures. In contrast, juvenile thyrotoxicosis causes accelerated growth and advanced bone age but paradoxically induces short stature because of premature closure of the epiphyses. In severe hyperthyroidism advanced intramembranous ossification causes early fusion of the skull sutures and craniosynostosis. These features are reproduced in T3 receptor (TR) ␤PV/PV mutant mice, which have skeletal thyrotoxicosis (3). In preFirst Published Online September 8, 2005 Abbreviations: CSS, Charcoal-stripped serum; C-tail, cytoplasmic tail; EGF, epidermal growth factor; FGF, fibroblast growth factor; FGFR, FGF receptor; HSPG, heparan sulfate proteoglycan; Ihh, Indian hedgehog; PDGF, platelet-derived growth factor; PLC, phospholipase C; STAT, signal transducer and activator of transcription; TR, T3 receptor. Endocrinology is published monthly by The Endocrine Society (http:// www.endo-society.org), the foremost professional society serving the endocrine community.

pression was reduced in growth plates from T3 receptor ␣-null mice, which exhibit skeletal hypothyroidism, but was increased in T3 receptor ␤PV/PV mice, which display skeletal thyrotoxicosis. These findings indicate that FGFR3 is a T3target gene in chondrocytes. In further experiments, T3 enhanced FGF2 and FGF18 activation of the MAPK-signaling pathway but inhibited their activation of signal transducer and activator of transcription-1. FGF9 did not activate MAPK or signal transducer and activator of transcription-1 pathways in the absence or presence of T3. Thus, T3 exerted differing effects on FGFR activation during chondrogenesis depending on which FGF ligand stimulated the FGFR and which downstream signaling pathway was activated. These studies identify novel interactions between T3 and FGFs that regulate chondrocyte proliferation and differentiation during chondrogenesis. (Endocrinology 146: 5568 –5580, 2005)

vious studies, we showed that T3 stimulates fibroblast growth factor (FGF) receptor (FGFR)-1 expression and enhances FGFR1 activity in osteoblasts by a mechanism that involves TR␣ (4). Activating mutations of FGFR1 result in Pfeiffer’s craniosynostosis syndrome in humans (5, 6), and increased FGFR1 expression is seen in TR␤PV/PV mice (3). Thus, FGFR1 is implicated in mediating the actions of T3 that regulate intramembranous ossification. However, the mechanisms by which T3 regulates endochondral ossification and linear growth remain unclear. Endochondral bone formation occurs when condensations of mesenchyme cells differentiate to chondrocytes and cells at the mesenchyme border form a perichondrium (7). The cartilage enlarges via proliferation of chondrocytes, which secrete a matrix rich in heparan sulfate proteoglycans (HSPGs), growth factors, and other signaling molecules. Proliferating chondrocytes differentiate into hypertrophic chondrocytes, which increase in size to promote bone growth. Hypertrophic chondrocytes also secrete specific matrix proteins and growth factors that regulate vascular invasion of cartilage and initiate bone deposition and mineralization by osteoblasts. Secondary centers of ossification form at the ends of long bones and are separated from the primary center by a growth plate containing organized columns of proliferating and hypertrophic chondrocytes. During endochondral ossi-

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Barnard et al. • T3 Regulates FGF Signaling in Chondrocytes

fication FGFR2 is expressed in condensing mesenchyme and perichondrium, FGFR1 is present in prehypertrophic and hypertrophic chondrocytes, and proliferating chondrocytes express FGFR3. During linear growth, FGFR2 is not expressed in the growth plate, but FGFR1 expression persists in prehypertrophic and hypertrophic cells, and proliferating chondrocytes express FGFR3 (6, 7). The distinct and adjacent regions of FGFR1 and FGFR3 expression suggest that these receptors possess unique functions, with FGFR3 proposed to regulate chondrocyte proliferation and differentiation and FGFR1 suggested to be involved in the regulation of hypertrophic chondrocyte survival (6). The role of FGFR1 and FGFR2 in endochondral ossification is poorly understood, although the occurrence of digit abnormalities in craniosynostosis syndromes due to activating mutations in FGFR1 and FGFR2 suggest that they regulate limb patterning (8). FGFR3, however, plays a crucial role. FGFR3-activating mutations result in autosomal dominant achondroplasia dwarfism syndromes (5, 6), whereas Fgfr3 knockout mice display limb overgrowth (9, 10), indicating that FGFR3 is a negative regulator of linear growth. There are 22 genes that encode distinct FGF ligands, and many are expressed in mesenchyme during endochondral ossification and in the growth plate during linear growth (6, 7). The roles of individual FGFs in skeletal development and growth are poorly defined probably because many have essential roles in other tissues during embryogenesis, and the effects of others may be masked by functional redundancy (8). Consequently, no FGF mutations have been described that result in abnormalities of endochondral ossification and linear growth. Nevertheless, studies of mutant mice and limb-culture systems have suggested that FGF9 and FGF18 act via FGFR3 to regulate endochondral ossification (6, 11, 12), whereas FGF2 was the first FGF ligand to be isolated from growth-plate chondrocytes. We and others showed that T3 inhibits chondrocyte proliferation (13) and is essential for organization of growthplate proliferating chondrocytes and terminal hypertrophic chondrocyte differentiation (14 –16). T3 appears to control the transition between chondrocyte proliferation and hypertrophic differentiation by blocking cell-cycle progression via induction of the p21 and p27kip1 cyclin/cyclin-dependent kinase inhibitors (17). T3 also influences the composition of sulfated proteoglycans in growth-plate cartilage and regulates expression of components of the Indian hedgehog (Ihh)/PTHrP feedback loop (16), a critical regulator of chondrocyte proliferation during endochondral ossification (7, 18, 19). HSPGs expressed on the cell surface and in cell matrix interact with both FGFs and FGFRs to form a ternary complex essential for FGFR activation and signaling (20, 21). HSPGs are also necessary for migration of secreted hedgehog proteins and have been implicated in the formation of growth factor gradients during bone formation (2). Furthermore, FGFs coordinate chondrocyte proliferation and differentiation by interacting with Ihh/PTHrP signaling via a pathway in which Ihh/PTHrP lies downstream of FGFR3 (22). Thus, the pace of chondrocyte proliferation, the transition between proliferation and hypertrophic differentiation, and the progression of differentiation are integrated by complex paracrine pathways involving HSPGs, FGFR3, and Ihh/PTHrP. Studies that show that T3 regulates HSPG composition and

Endocrinology, December 2005, 146(12):5568 –5580

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the Ihh/PTHrP feedback loop (16) and exerts important effects on chondrocyte proliferation and differentiation (13, 15, 17) suggest that FGFR3 may also be involved in pathways by which alterations of thyroid status regulate endochondral ossification and adjust the rate of linear growth (2). We hypothesized, therefore, that regulation of chondrogenesis involves T3-enhanced activation of FGFR3. To investigate this, we studied chondrogenesis in vitro in ATDC5 cells (23), which express the same TR isoforms as primary growthplate chondrocytes and respond to T3 (13, 24), express FGFRs and respond to FGFs (25), and undergo a well-described reproducible program of chondrogenesis (26). In these studies, we identified the various FGFR isoforms expressed in ATDC5 cells during chondrogenesis and investigated whether T3 regulates FGFR expression and mRNA splicing and function. Materials and Methods Cell culture ATDC5 cells (kindly provided by Dr. A. Grigoriadis, Guy’s Hospital, London, UK) were maintained in DMEM/Ham’s F12 (1:1; Life Technologies, Paisley, UK) containing 5% fetal bovine serum (Bio-Whittaker, Wokingham, UK), 10 ␮g/ml transferrin (Sigma, Dorset, UK), and 3 ⫻ 10⫺8 mol/liter sodium selenite (Sigma) at 37 C in 5% CO2, as described (23, 24). Cells (passage number 20 –23) were harvested at d 0, resuspended in medium containing 5% charcoal-stripped serum (CSS) (27), and plated at 350,000 cells/flask, 100,000 cells/well, 40,000 cells/well, or 4000 cells/well in 75-cm2 flasks, six-, 12-, or 96-well plates, respectively. Differentiation medium, which contained 5% CSS with transferrin, sodium selenite, and 10 ␮g/ml bovine insulin (Sigma), was added on d 1. At d 21, differentiation medium was replaced with mineralization medium, consisting of ␣-modified Eagle medium (Life Technologies) plus 5% CSS, transferrin, sodium selenite, and insulin, in which cells were cultured in 3% CO2 at 37 C to facilitate calcification of chondrocyte nodules (23, 24). Cells were treated without or with 100 nm T3 throughout the culture period and stimulated for 0 –30 min with 5 ng/ml FGF2, FGF9, or FGF18 (Sigma), 50 ng/ml epidermal growth factor (EGF) (Sigma), or 50 ng/ml platelet-derived growth factor (PDGF) (Sigma) before harvesting. We previously showed that FGFR1 mRNA expression is increased by T3 in osteoblastic cells in a concentration-dependent manner, with a threshold response detected after 1 nm T3 treatment and a maximal effect evident after treatment with 100 nm T3 (4). We also documented similar concentration-dependent antiproliferative actions of T3 in growth-plate chondrocytes (13). ATDC5 cell proliferation was determined by counting viable cells in a hemocytometer after staining with trypan blue. Cell growth was also determined by measurement of sulforhodamine-B (Sigma) absorbance at 540 nm after incubating cells with 0.4% sulforhodamine-B in 1% acetic acid (13, 24). ATDC5 cell differentiation was determined by alcian blue staining of chondrocyte nodules and secreted cartilage matrix, alizarin red staining of mineralized nodules, measurement of alkaline phosphatase activity, and analysis of collagen X expression, as described (13, 24).

Experimental animals and primary cell culture Breeding and handling of wild-type and TR␤PV mice were conducted in strict accordance with the National Institutes of Health Guide for Care and Use of Laboratory Animals and were approved by the National Cancer Institute Animal Care and Use Committee. Breeding and handling of wild-type and TR␣0/0 mice were carried out in a certified animal facility at Universite´ Victor Segalen (Bordeaux, France) according to procedures approved by the local animal care and use committee. Primary chondrocytes were prepared from ventral rib cages of newborn TR␣0/0 mice. Ribs were collected into PBS containing amphotericin B (25 ␮g/ml), penicillin (500 U/ml), streptomycin (500 ␮g/ml), and gentamicin (500 ␮g/ml), rinsed in PBS, and incubated in 2 mg/ml protease (Sigma) for 20 min at 37 C. Ribs were rinsed further in PBS and incubated in 2 mg/ml type I collagenase (Sigma) in DMEM/F12 for 20 min. Soft

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tissue was discarded; the ribs were rinsed again in PBS and then incubated in 2 mg/ml collagenase for 4 h. Cells were pelleted and suspended in DMEM/F12 plus 2 mm glutamine, 10% fetal bovine serum, 50 U/ml penicillin, 50 ␮g/ml streptomycin, and 50 ␮g/ml gentamicin and then plated at a density of 50,000/cm2. Confluent cells were washed in PBS and incubated in DMEM/F12 plus antibiotics, 1.25% BSA, transferrin, sodium selenite, and 10 ␮g/ml bovine insulin in the absence or presence of 10 nm T3 for 6 and 12 d and then treated with FGF2, FGF9, or FGF18 for 7.5 min before harvest.

RT-PCR cDNA was synthesized from total RNA (2.5 ␮g) using SuperScript II reverse transcriptase (Invitrogen), and 1 ␮l of cDNA was used for PCR amplification of FGFRs 1– 4, collagen X, and 18S rRNA, as described (13, 28, 29). Primers were derived from GenBank sequences (nucleotide positions are given in the 5⬘-3⬘ direction of the synthesized oligonucleotide): Fgfr1 (GenBank accession no. NM_010206) forward primers SPf (primer 1.1; nucleotides 81–101), VTf (3.1; 1286 –1305), and TKf (5.1; 2035–2056) and reverse primers SPr (2.1; 1175–1149), VTr (4.1; 1385– 1366), and TKr (6.1; 2470 –2451); Fgfr2 (GenBank accession no. NM_010207) forward primers SPf (1.2; 625– 647), VTf (3.2; 1841–1860), and TKf (5.2; 2592–2611) and reverse primers SPr (2.2; 1729 –1708), VTr (4.2; 1940 –1921), and TKr (6.2; 3025–3005); Fgfr3 (GenBank accession no. NM_008010) forward primers SPf (1.3; 231–253), VTf (3.3; 1411–1430), and TKf (5.3; 2155–2176) and reverse primers SPr (2.3; 1300 –1281), VTr (4.3; 1510 –1485), and TKr (6.3; 2590 –2571); Fgfr4 (GenBank accession no. NM_008011) forward primers SPf (1.4; 124 –146), VTf (3.4; 1291–1310) and TKf (5.4; 2067–2086) and reverse primers SPr (2.4; 1163–1141), VTr (4.4; 1395–1376) and TKr (6.4; 2490 –2481); Col10a1 (GenBank accession no. Z21610) forward primer CollXf (236 –255) and reverse primer CollXr (565–546); 18S rRNA (GenBank accession no. X00686) forward primer 18Sf (1577–1596) and reverse primer 18Sr (1727–1708). PCRs were performed with an initial denaturation step at 94 C for 3min, cycles of 30 sec at 94 C, 30 sec at an annealing temperature ranging between 45– 60 C, depending on the primer pairs used, and 30 sec at 72 C, followed by a termination step at 72 C for 5 min. PCR products were subcloned into pGEM T-Easy vector (Promega, Southampton, United Kingdom) and sequenced. Semiquantitative RT-PCR was optimized to detect linear accumulation of FGFR mRNA splice variants, as described (29). A range of input RNA concentrations (0.625–5 ␮g) was tested over a range of 15–35 PCR cycles. Products were Southern blotted and probed with an internal 32 P-labeled oligonucleotide probe specific for each pair of PCR primers. The range of linear accumulation of FGFR mRNAs was determined and assays designed to detect accumulation of PCR products in the middle of the linear range to facilitate the relative quantitation of individual FGFR mRNA isoforms (data not shown). Thus, 1 ␮l of cDNA prepared from 2.5 ␮g of RNA was amplified using the optimized number of PCR cycles for each gene (20 cycles for 18S rRNA; 28 cycles for collagen X; and 28, 28, and 25 cycles for FGFR1, FGFR2, and FGFR3, respectively). FGFR4 expression was not detected in ATDC5 cells.

Western blotting and immunoprecipitation Cells from 6-well plates were lysed with 500 ␮l/well lysis buffer (10 mm Tris-HCl, 75 mm NaCl, 10 mm EDTA, 1% Triton X-100, and 0.5% SDS) in the presence of protease inhibitors (phenylmethylsulfonylfluoride, aprotinin, and leupeptin). Twenty microliters of lysate were resolved on a 10% SDS-PAGE gel and analyzed by Western blotting, as described (4, 30), using an enhanced chemiluminescence detection system (Amersham Biosciences United Kingdom, Little Chalfont, UK). For analysis of MAPK activation, filters were incubated with polyclonal antibodies to nonphosphorylated p42 and p44 components of the MAPK pathway (1:1000 dilution, New England Biolabs, Hitchin, UK). Filters were then stripped at 56 C for 30 sec in 6.25 mm Tris-Cl (pH 6.8), 10 mm ␤-mercaptoethanol, 2% SDS, and reprobed with anti-phospho-p42/p44 antibodies that recognize phosphorylated p42 and p44 proteins (1:1000 dilution, New England Biolabs), as described (4). Additional primary antibodies were used similarly to investigate expression of FGFR downstream signaling pathways by Western blotting or immunoprecipitation. These included anti-signal transducer and activator of transcription (STAT) 1 (Santa Cruz Biotechnology, Santa Cruz, CA), anti-phospho-


STAT1 (Santa Cruz Biotechnology), anti-STAT3 (New England Biolabs), anti-phospho-STAT3 (New England Biolabs), anti-STAT5 (Upstate Biotechnology, Milton Keynes, UK), anti-phospho-STAT5 (Upstate Biotechnology), anti-phospholipase C (PLC) ␥1 (Santa Cruz Biotechnology), anti-phospho-PLC␥ (Santa Cruz Biotechnology), and anti-phospho-tyrosine 4G10 (Upstate Biotechnology). For immunoprecipitations (4), cells from 6-well plates were lysed in 500 ␮l/well lysis buffer as above. Lysates were precleared overnight by incubating with 5 ␮l of goat antimouse IgG (Upstate Biotechnology) at 4 C, followed by 2-h incubation with 50 ␮l of Protein G-Sepharose (Bio-Rad Laboratories, Hemel Hempstead, UK). After centrifugation at 13,000 rpm for 3 min, the supernatant was incubated overnight at 4 C with 5 ␮l of appropriate primary antibody and precipitated by addition of 50 ␮l of Protein G-Sepharose suspension for 2 h at 4 C. Samples were spun at 13,000 rpm for 3 min, and the pellet was resuspended in sample loading buffer (62.5 mm Tris-HCl pH 6.8, 20% glycerol, 2% SDS, 5% ␤-mercaptoethanol) before being denatured at 96 C for 5 min. Sepharose was pelleted at 13,000 rpm for 3 min, and 15 ␮l of supernatant was resolved by 10% SDS-PAGE. Phosphorylated (activated) forms of the immunoprecipitated proteins were then detected by Western blotting as above using an anti-phospho-tyrosine antibody.

In situ hybridization Expression of FGFR3 mRNA was analyzed in growth-plate sections from 3-wk-old TR␣0/0 and TR␤PV mice by in situ hybridization. A bacterial neomycin resistance-gene cRNA probe (Roche Molecular Biochemicals, Lewes, UK) was used as a negative control for all hybridizations, as described in studies in which we optimized in situ hybridization methods (3, 16). A partial mouse FGFR3 cDNA (nucleotides 234 – 680, GenBank accession no. NM_008010) was isolated by RT-PCR from ATDC5 cells, subcloned into the pGEM-T vector (Promega), and sequenced. The FGFR3 construct was linearized with NcoI, and digoxigenin-labeled antisense cRNA probe was synthesized using SP6 RNA polymerase (Roche Molecular Biochemicals). Hybridization of labeled probe to growth-plate sections was detected using alkaline phosphatase-conjugated antidigoxigenin Fab fragments, as described (3, 4, 16).

Statistical analysis Data were analyzed by Student’s t test and by one-way ANOVA followed by Tukey-Kramer’s multiple comparison post hoc test using GraphPad Prism Version 4 (GraphPad Software Inc., San Diego, CA).

Results T3 stimulates chondrogenesis

ATDC5 cells were maintained in the absence or presence of T3 (100 nm) for 28 d. T3 reduced the number of cells by 50% after 6 d, but, thereafter, growth curves of control and T3treated ATDC5 cells followed a parallel course, indicating that T3 inhibited cell proliferation during the first 6 d (Fig. 1A). To determine whether T3 influenced chondrogenesis, three differentiation markers were examined (13, 26, 31–33). As previously shown in primary chondrocytes (13), T3 increased alkaline phosphatase activity (Fig. 1B), enhanced formation of cartilage nodules (Fig. 1C), and stimulated expression of the hypertrophic chondrocyte marker collagen X (Fig. 1D). Thus, T3 inhibited ATDC5 cell proliferation but stimulated the earlier onset of chondrogenesis and hypertrophic differentiation. Numerous FGFR isoforms are expressed in ATDC5 cells

FGFRs 1–3 are expressed in bone and cartilage and implicated in skeletal development (6). We previously showed that T3 stimulates FGFR1 expression and activity in osteoblasts (3, 4). To investigate whether T3 effects on chondro-



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FIG. 1. T3 inhibits ATDC5 cell proliferation but stimulates hypertrophic chondrocyte differentiation. ATDC5 cells were seeded on d 0 (40,000 cells/well) and maintained in the absence or presence of T3 (100 nM). A, Effect of T3 on cell proliferation (n ⫽ 6 experiments, mean number of cells per well ⫾ SEM). B, Effect of T3 on alkaline phosphatase activity (n ⫽ 3 experiments in duplicate). Alkaline phosphatase activity is shown as units of enzyme activity per cell number ⫾ SEM. Results were analyzed by Student’s t test and also by one-way ANOVA, followed by Tukey-Kramer’s multiple comparison post hoc test. *, P ⬍ 0.05; **, P ⬍ 0.01; ***, P ⬍ 0.001. C, Alcian blue staining of chondrocyte colonies in control and T3-treated cultures maintained between 6 and 28 d. D, Effect of T3 on collagen X mRNA expression in ATDC5 cells. Data in C and D are single experiments representative of at least three independent experiments performed in duplicate.

genesis also involved FGFR signaling, we first determined which FGFRs were expressed in ATDC5 cells. The murine Fgfr1–3 genes each contain 19 exons (Fig. 2A). Their mRNAs undergo alternative splicing of exons 8 (Ig-like domain IIIb) and 9 (IgIIIc) to yield transcripts expressed in epithelial tissues (exon 8) or mesenchyme (exon 9) (34). FGFR4 is not expressed in bone (6). RT-PCR analysis of ATDC5 cell RNA revealed numerous FGFR splice variants. Two FGFR1 isoforms (i and ii) were identified (Fig. 2B). Isoform (i) was a full-length transcript containing all three Ig domains, whereas isoform (ii) lacked exon 3 (IgI domain) and included six additional nucleotides at the 3⬘ end of exon 4, which encodes the acid box. The significance of inclusion or exclusion of exon 3 is largely unknown, although FGFR1 variants lacking the IgI domain exhibit a higher affinity for FGF1 and FGF2 than full-length FGFR1 (34). The additional nucleotides at the 3⬘ boundary of exon 4 encode the amino acids Met-Arg and are included by use of an alternative 5⬘-splice donor site (35, 36), but their functional significance has not been determined. Both FGFR1 isoforms contained exon 9 (mesenchyme IgIIIc variant) and a VT motif in the transmembrane domain. The VT motif arises from use of an alternative 5⬘-splice donor site, so that six additional nucleotides encoding Val-Thr are included at the 3⬘ end of exon 10. The Thr residue can be phosphorylated by protein kinase C (37), and the motif is required for interactions between FGFRs and the essential adaptor protein FGF receptor substrate 2, which is necessary for FGFR-mediated MAPK signaling (38).

Four FGFR2 isoforms (i–iv) were identified (Fig. 2C). Isoform (i) was a full-length variant, isoform (ii) lacked exon 3, and isoform (iii) lacked exons 3 and 4 (IgI and acid box domains). The acid box is essential for attachment of HSPGs to an adjacent serine residue, whereas the presence of the IgI domain inhibits HSPG attachment (39). Attachment of HSPGs increases FGFR ligand-binding affinity, enhances receptor autophosphorylation, and sustains downstream signaling responses to FGF. Thus, alternative splicing of exons 3 and 4 regulates the activity of FGFR2 (39). Isoform (ii) in ATDC5 cells can be predicted to contain HSPG attachments, whereas isoforms (i) and (iii) do not, suggesting that their functional responses to FGF stimulation may differ. The fourth FGFR2 isoform (Fig. 2D) differed from the full-length transcript by the inclusion of a 92-bp sequence at the 5⬘ end of exon 19 to produce a previously unreported FGFR2 mRNA. This variant arises by use of an alternative 3⬘-splice acceptor site located in the intron 92 nucleotides upstream of exon 19. The alternative acceptor site has the same sequence as the previously described splice site, and both are preceded by 18 identical nucleotides. The inserted 92-bp sequence contains an in-frame UGA stop codon, such that the new FGFR2 mRNA is predicted to encode a truncated protein that includes an additional 20 amino acids following those encoded by exon 18 but lacks all 54 amino acids encoded by exon 19. The tyrosine kinase II domain is encoded by exons 15–18, and the 54 amino acids encoded by exon 19 form a cytoplasmic tail (C-tail) of unknown function (40, 41). Substitution of the 54 amino acid C-tail with a novel 20 amino

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FIG. 2. Structure of mouse Fgfr genes and FGFR1, FGFR2, and FGFR3 mRNA splice variants in ATDC5 cells. A, General structure of the murine Fgfr genes with representations of transcribed mRNA and translated proteins. Exon 1 contains the 5⬘ untranslated region. The translation start site (ATG, AUG) is located in exon 2, which encodes the signal peptide (SP). Exons 3–9 encode the extracellular region of the FGFR protein, with exon 3 encoding an Ig-like domain (IgI), exon 4 encoding the acid box (AB), exons 5 and 6 encoding the IgII domain, and exon 7, together with either exon 8 (in epithelial cells) or 9 (in mesenchymal cells), encoding the IgIII domain. Exons 10 and 11 encode the transmembrane (TM) and juxtamembrane (JM) regions, and exons 12–18 encode intracellular tyrosine kinase (TK) domains. Exon 19 encodes the C-tail of the receptor and contains the stop codon (TGA, UGA) and 3⬘ untranslated region. Primers used in RT-PCRs are shown as arrows and numbered 1– 6 and also labeled as described in Materials and Methods. B, FGFR1 mRNAs cloned from ATDC5 cells. Two FGFR1 variants are shown. One complete isoform (i) contains exon 9 (mesenchyme cell IgIIIc domain) and a VT motif in the transmembrane region encoded by exon 10 (hatched box). A second isoform (ii) lacks exon 3 but includes an additional six nucleotides in exon 4 (4 ⫹ 6). C, FGFR2 mRNAs cloned from ATDC5 cells. Four FGFR2 variants are shown. A complete isoform (i) contains exon 9, along with the VT motif in exon 10. A second isoform (ii) lacks exon 3. A third isoform (iii) lacks exon 3 and exon 4. A fourth isoform (iv) consists of the complete FGFR2 sequence but includes an additional 92 nucleotides at the 5⬘ end of exon 19. D, Alternative splicing in the fourth FGFR2 isoform and the sequences of the intron/exon boundaries 5⬘ to exon 19 are also shown. Splicing event 1 shows the previously described joining of exon 18 to exon 19. The 92-bp sequence is shown in lower case italics, and exon 19 is shown in upper case. The stop codon in exon 19 (TGA) is indicated in bold, and the 3⬘ donor splice site (ag) is shown in bold underlined italics. Splicing event 2 shows the joining of exon 18 to an alternative 3⬘-splice acceptor site (ag) 92 nucleotides upstream of exon 19. The 92-bp sequence is shown in upper case italics, with its internal stop codon indicated in bold (TGA). E, The single FGFR3 mRNA cloned from ATDC5 cells, which contains exon 9 and the VT motif in exon 10.


acid sequence, therefore, may alter FGFR2 kinase activity or specificity. All FGFR2 isoforms also contained exon 9 and the exon 10 VT motif. A single FGFR3 isoform was identified (Fig. 2E), which was a full-length transcript containing exon 9 and the VT motif in exon 10. An alternative FGFR3 mRNA splice variant lacking the acid box domain has also been identified in ATDC5 cells (25), but this isoform was not detected in these studies. This discrepancy may reflect differences between batches of ATDC5 cells, which were originally derived from an embryonal carcinoma in 1997 (26), or differences in cell culture conditions. Our cells were obtained from a collaborating laboratory in the United Kingdom (Dr. A. Grigoriadis, Guy’s Hospital, London, UK), whereas the previous report was published by the laboratory that originally isolated ATDC5 cells in Japan (25). No FGFR4 mRNAs and no secreted FGFR variants were identified in ATDC5 cells. T3 stimulates FGFR mRNA expression in differentiating ATDC5 cells

The relative concentrations of FGFR mRNAs were determined by RT-PCR and Southern blot analysis using RNA extracted from ATDC5 cells undergoing chondrogenesis in the absence or presence of T3. Primer pairs 1 (SPf) and 6 (TKr), 1 (SPf) and 4 (VTr), or 5 (TKf) and 6 (TKr) amplified fulllength FGFR cDNAs, the extracellular and transmembrane domains, or tyrosine kinase domains (Fig. 2A). T3 stimulated expression of all FGFR mRNAs in ATDC5 cells. The response to T3 was maximal after 6 d (Fig. 3), coinciding with the time at which T3 inhibited cell proliferation and initiated the onset of hypertrophic chondrocyte differentiation (Fig. 1, A and B). The effect of T3 on FGFR3 expression (17.5-fold induction) was much greater than the effects on FGFR1 (6.9-fold) and FGFR2 (6.1-fold). The magnitude of T3 response for all FGFRs diminished as chondrogenesis progressed, although T3 stimulation of FGFR3 was maintained in differentiating chondrocytes until d 21 (Fig. 3). The relative levels of expression of individual FGFR mRNA isoforms were not altered by T3 at any time point, indicating that alternative splicing of FGFR1 and FGFR2 mRNAs was not regulated by T3 during chondrogenesis. Furthermore, in

FIG. 3. Graph showing the effect of T3 treatment on FGFR mRNA expression. The mean T3 induction of all FGFR1, FGFR2, and FGFR3 mRNA isoforms is shown in ATDC5 cells cultured for 0, 6, 12, 21, and 28 d in the absence or presence of T3 (100 nM). Levels of mRNA expression are shown as mean (n ⫽ 4) ⫾ SEM normalized to expression of 18S rRNA. Results were analyzed by Student’s t test and also by one-way ANOVA followed by Tukey-Kramer’s multiple comparison post hoc test. *, P ⬍ 0.05; **, P ⬍ 0.01; ***P ⬍ 0.001.


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the absence of T3 treatment, there was no effect of chondrogenesis per se on expression of FGFR1, FGFR2, or FGFR3 (data not shown). FGFR3 mRNA expression is regulated by T3 in vivo

We previously showed that T3 stimulates FGFR1 in osteoblasts in vitro and in vivo, whereas FGFR2 is not regulated by T3 (3, 4). By contrast, FGFR3 is the major FGFR expressed in growth-plate cartilage (6). Accordingly, mutations of FGFRs 1 and 2 primarily affect osteoblast function and result in craniosynostosis syndromes, whereas FGFR3 mutations affect chondrocyte function resulting in dwarfism and other chondrodysplasias (5, 6). We, therefore, examined FGFR3 expression in growth plates from TR␣0/0 mice, which exhibit skeletal hypothyroidism (4, 42), and in TR␤PV/PV mice with skeletal thyrotoxicosis (3) (Fig. 4). Comparisons between levels of FGFR3 mRNA expression in wild-type and mutant littermates were made by in situ hybridization. In these experiments, samples from TR␣0/0 and wild-type controls were probed and developed in parallel, and the development process was stopped when FGFR3 expression was first detected. Similarly, samples from TR␤PV/PV and their littermate controls were examined in parallel. FGFR3 expression was markedly reduced in TR␣0/0 mice but increased in growth plates from TR␤PV/PV mice (Fig. 4). Thus, FGFR3 mRNA expression in the growth plate is regulated in vivo by changes in thyroid status. In previous studies, we also showed that FGFR1 mRNA is markedly increased in growth-plate chondrocytes from TR␤PV/PV mice (3). FGFR2 mRNA was undetectable by in situ hybridization in the growth plate in both wild-type and mutant mice (data not shown).

FIG. 4. In situ hybridization (⫻200 magnification) showing FGFR3 mRNA expression in tibial growth plates from 3-wk-old wild-type (WT) and homozygous TR␣ knockout (TR␣0/0) and WT and homozygous TR␤ mutant (TR␤PV/PV) mice. Arrows show proliferating chondrocytes staining positive for FGFR3.

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FGFR-mediated activation of MAPK is enhanced by T3 in ATDC5 cells

Because T3 stimulated FGFR3 expression in vitro and in vivo, and also increased expression of FGFRs 1 and 2 in ATDC5 cells, we next investigated the effect of T3 on FGF signaling. FGFs bind to FGFRs and stimulate receptor autophosphorylation and downstream signaling via three main routes in bone cells: the MAPK cascade, via PLC␥, or by activation of STAT pathways (21, 43– 47). T3 enhanced FGF2-induced activation of MAPK maximally (3-fold) after 6 d in culture and also after 12 d (2.5-fold), but no T3 enhancement was observed in cells cultured for 21 or 28 d (Fig. 5). These data, which demonstrate that the


maximal effects of T3 on MAPK responses occurred in cells cultured for up to 12 d, are consistent with the finding that the maximal effects of T3 on FGFR expression were also seen during this time period (Fig. 3), when an antiproliferative response to T3 was observed (Fig. 1A). To investigate whether the effect of T3 on growth factorstimulated activation of MAPK was specific to FGF, we next examined responses to EGF and PDGF. EGF (50 ng/ml) and PDGF (50 ng/ml) stimulated MAPK by 1.5-fold (P ⬍ 0.05) and 2-fold (P ⬍ 0.001), respectively, after 5-min treatment in ATDC5 cells cultured for 6 d in the absence of T3. However, neither EGF nor PDGF activated MAPK in the presence of T3 in cells cultured for 6 d, demonstrating that T3 inhibited EGF

FIG. 5. FGF2-induced MAPK activation in ATDC5 cells cultured for 6 (A), 12 (B), 21 (C), and 28 (D) d in the absence or presence of T3 (100 nM). Cells were stimulated with FGF2 (5 ng/ml) for 2, 5, or 30 min. Proteins were analyzed by Western blotting for expression of basal p44 and p42 components of MAPK and phosphorylated p44 and p42 (pMAPK). Western blots show a single experiment. Data from three experiments are shown in the graphs. Activated pMAPK values at each time point were normalized to the level of basal MAPK. Results are shown as mean ⫾ SEM and represent fold activation of normalized pMAPK at each time point relative to untreated cells. Results were analyzed by Student’s t test and also by one-way ANOVA, followed by Tukey-Kramer’s multiple comparison post hoc test. *, P ⬍ 0.05; **, P ⬍ 0.01.


and PDGF activation of MAPK. There was no MAPK response to either EGF or PDGF in the absence or presence of T3 in cells cultured for 12, 21, or 28 d (data not shown). These findings indicate that T3 enhancement of MAPK signaling does not involve EGF or PDGF receptors in ATDC5 cells. To investigate whether T3 also influenced other FGFR downstream responses, we next examined the effect of T3 on STAT1. In contrast to the enhancement of MAPK responses to FGF2 by T3, FGF2-induced activation of STAT1 in cells cultured for 6 d was blocked by T3 (compare Fig. 6A with Fig. 5A). There was no effect of FGF2 on STAT1 activation in the absence or presence of T3 in cells cultured for 12 d (Fig. 6B). In cells cultured for 21 and 28 d, FGF2 treatment in the presence of T3 resulted in 50% inhibition of STAT1 activation (Fig. 6, C and D). These data demonstrate that T3 exerts differential effects on FGFR downstream responses to FGF2. We next investigated whether FGF2 activated other downstream pathways. Basal expression of STAT3, STAT5, and PLC␥ proteins could be detected readily in ATDC5 cells and HeLa cell-positive controls. By contrast, although activated

FIG. 6. FGF2-induced STAT1 activation in ATDC5 cells cultured for 6 (A), 12 (B), 21 (C), and 28 (D) days in the absence or presence of T3 (100 nM). Cells were stimulated with FGF2 (5 ng/ml) for 2, 5, or 30 min. Proteins were analyzed by Western blotting for expression of phosphorylated pSTAT1 and ␤-actin. Western blots show a single experiment. Data from three experiments are shown in the graphs. Activated pSTAT1 values at each time point were normalized to the level of ␤-actin. Results are shown as mean ⫾ SEM and represent fold activation of normalized pSTAT1 at each time point relative to untreated cells. Results were analyzed by Student’s t test and also by one-way ANOVA, followed by Tukey-Kramer’s multiple comparison post hoc test. *, P ⬍ 0.05.


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phosphorylated pSTAT3, pSTAT5, and pPLC␥ proteins were readily identified in cell extracts from interferon ␣-stimulated HeLa cells, activated proteins could not be detected in ATDC5 cells stimulated with FGF2 (5 ng/ml) for 5 min (data not shown). To investigate whether the effects of T3 on FGFR signaling were restricted to modulation of FGF2 actions, we also investigated MAPK and STAT1 responses to FGF9 and FGF18 in the absence and presence of T3. FGFs 9 and 18 are the major FGF ligands implicated in regulation of endochondral ossification (6, 11, 12). Treatment with FGF9 in the absence or presence of T3 had no effect on MAPK or STAT1 activation (Fig. 7, A and C). In contrast, T3 enhanced FGF18-stimulated activation of MAPK in cells cultured for 6 d but inhibited the stimulation of STAT1 activation by FGF18 at the same time point (Fig. 7, B and D). These data further demonstrate that T3 exerts FGF ligand-specific effects on FGFR signaling. Finally, we determined whether T3 influenced FGFR signaling in primary chondrocytes from wild-type and TR␣0/0 mice (Fig. 8). In wild-type chondrocytes treated for 6 d, T3

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FIG. 7. MAPK (A and B) and STAT1 (C and D) activation in ATDC5 cells cultured for 6 d in the absence or presence of T3 (100 nM) and stimulated with 5 ng/ml FGF9 (A and C) or FGF18 (B and D) for 2, 5, or 30 min. Proteins were analyzed by Western blotting for expression of basal p44 and p42 components of MAPK and phosphorylated p44 and p42 (pMAPK) and for expression of phosphorylated pSTAT1 and ␤-actin. Western blots show a single experiment. Data from three experiments are shown in the graphs. Activated pMAPK or pSTAT1 values at each time point were normalized to the level of basal MAPK or ␤-actin, respectively. Results are shown as mean ⫾ SEM and represent fold activation of normalized pMAPK or pSTAT1 at each time point relative to untreated cells. Results were analyzed by Student’s t test and also by oneway ANOVA, followed by Tukey-Kramer’s multiple comparison post hoc test. *, P ⬍ 0.05; ***P ⬍ 0.001.

enhanced FGF2- and FGF9-stimulated activation of MAPK by 2.0- and 2.9-fold, respectively. Furthermore, T3 enhanced FGF2-, FGF9-, and FGF18-stimulated activation of MAPK by 2.7-, 3.5-, and 4.3-fold, respectively, in wild-type cells treated for 12 d. By contrast, T3 did not enhance FGFR signaling in response to any FGF ligand in TR␣0/0 chondrocytes after 6 or 12 d in culture. These findings are similar to the T3 enhancement of FGFR signaling seen in ATDC5 cells cultured for 6 and 12 d and correlate with the reduction of FGFR3 mRNA expression observed in growth plates from TR␣0/0 mice (Fig. 4). Nevertheless, FGF9 had no effect on MAPK in ATDC5 cells. This altered response in primary chondrocytes may result from differing culture conditions between primary and ATDC5 cells or from the different origins of the two cell types (primary chondrocytes are heterogeneous and derived from rib cartilage, whereas ATDC5 cells do not originate from the same site and are clonal). Despite these dif-

ferences, T3 consistently stimulated FGFR signaling in both systems. Discussion

In previous studies, we found that TR␣ and TR␤ proteins are expressed in ATDC5 cells undergoing chondrogenesis, in primary growth-plate chondrocytes, and in progenitor cells and proliferating chondrocytes in the growth plate in vivo (13, 16, 24). Hypothyroidism or deletion of TR␣ resulted in impaired hypertrophic chondrocyte differentiation in vivo (16, 42), and TRs were not expressed in terminally differentiated hypertrophic chondrocytes (13, 16). In the current studies, we show that T3 stimulates chondrogenesis in cultured ATDC5 cells by inhibiting cell proliferation and stimulating the onset of hypertrophic chondrocyte differentiation. The antiproliferative effects of T3 occurred during the



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FIG. 8. FGF2-, FGF9-, and FGF18-induced MAPK activation in primary wild-type (A and C) and TR␣0/0 (B and D) chondrocytes cultured for 6 (A and B) or 12 (C and D) days in the absence or presence of T3 (10 nM). Cells were stimulated with FGFs (5 ng/ml) for 7.5 min. Proteins were analyzed by Western blotting for expression of basal p44 and p42 components of MAPK and phosphorylated p44 and p42 (pMAPK). Western blots show a single experiment. Data from two experiments performed in duplicate are shown in the graphs. Activated pMAPK values for each treatment were normalized to the level of basal MAPK. Results are shown as mean ⫾ SEM and represent fold activation of normalized pMAPK for each FGF treatment relative to control cells not treated with FGF. Results were analyzed by Student’s t test and also by one-way ANOVA, followed by Tukey-Kramer’s multiple comparison post hoc test. *, P ⬍ 0.05; **, P ⬍ 0.01.

first 6 d of ATDC5 cell culture, and differentiation progressed until the onset of terminal hypertrophic chondrocyte differentiation by 21 d in T3-treated cells. T3 stimulation of FGFR1 and FGFR2 in ATDC5 cells undergoing chondrogenesis occurred until d 12, but stimulation of FGFR3 by T3 was greater and persisted until d 21, coinciding with the period in which T3 inhibited chondrocyte proliferation and advanced the onset of hypertrophic differentiation. We have previously shown that FGFR1 acts downstream of T3 in osteoblasts (4) but, although FGFR1 is expressed in growth-plate cartilage and overexpressed in TR␤PV/PV mice with skeletal thyrotoxicosis (3), the major FGFR involved in endochondral ossification and linear growth is FGFR3 (6). Taken together, these

findings suggest that FGFR3 may be an important mediator of T3 effects on chondrogenesis. FGFR3, which is expressed in proliferating and early hypertrophic chondrocytes, plays an important role in regulating chondrocyte differentiation and growth. Activating mutations of FGFR3 result in several forms of dwarfism in humans (5, 6, 48), and mutant mice with activating mutations of Fgfr3 recapitulate the dwarf phenotypes that result from decreased rates of chondrocyte proliferation and the presence of shortened proliferating and hypertrophic chondrocyte regions in the growth plate (5, 6, 48, 49). In contrast, knockout of Fgfr3 in mice causes an increased rate of chondrocyte proliferation with expanded regions of proliferating

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and hypertrophic chondrocytes (9, 10). Accordingly, FGFR3 has been proposed as a negative regulator of chondrocyte proliferation and differentiation. However, this view has also been challenged by studies showing that FGFs initiate and accelerate chondrocyte differentiation (22, 50) and matrix production via activation of FGFR3 (12). It has also been suggested that FGFR3 may either promote or inhibit chondrocyte proliferation during endochondral ossification according to the stage of development (51). Thus, the precise mechanism by which FGFR3 regulates growth is not yet clear. Hypothyroidism causes delayed bone age and growth retardation. In contrast, thyrotoxicosis leads to accelerated growth and advanced bone age but causes short stature because of premature growth-plate fusion (1, 2). Thyroid hormone replacement in hypothyroidism induces catch-up growth and accelerated bone maturation (52), demonstrating that thyroid hormones, as well as FGFs, regulate the onset and rate of progression of endochondral ossification. Previously, we showed that TR␤PV/PV mutant mice with skeletal thyrotoxicosis have accelerated and disproportionate narrowing of proliferating and hypertrophic chondrocyte zones in the growth plate (3). These findings are similar to the narrowed proliferating and hypertrophic regions observed in achondroplastic Fgfr3 mutant mice (5, 6, 48, 49). Consistent with these observations, we show in these studies that FGFR3 expression is increased in growth plates from TR␤PV/PV mice (Fig. 4). Conversely, we previously demonstrated delayed ossification and impaired hypertrophic chondrocyte differentiation in hypothyroid rats (16) and TR␣ knockout mice (42), and in the current studies, we demonstrate that FGFR3 expression is reduced in growth plates from TR␣0/0 mice (Fig. 4). Deletion of Fgfr3 causes growth-plate widening primarily due to an increase in numbers of proliferating chondrocytes (9, 10, 49), which may result from delayed and impaired hypertrophic chondrocyte differentiation. These findings indicate that both T3 and FGFR3 inhibit chondrocyte proliferation and induce hypertrophic chondrocyte differentiation. Some of the growth inhibitory actions of FGFR3 are mediated via reduced activity of the Ihh/PTHrP feedback loop (22). Similarly, in thyrotoxicosis, expression of the PTHrP receptor is markedly reduced, thereby preventing negative PTHrP signaling and facilitating accelerated hypertrophic differentiation (16). Hypothyroidism, conversely, increases PTHrP expression, resulting in an enhanced negative signal to inhibit hypertrophic chondrocyte differentiation. Importantly, expression of TR proteins, FGFR3, and components of the Ihh/PTHrP feedback loop are colocalized in proliferating and prehypertrophic chondrocytes (5, 6, 13, 16, 22, 48, 49). Taken together, these findings suggest that FGFR3 may play a role in mediating effects of T3 on chondrogenesis. Nevertheless, T3 and FGFR3 must also exert distinct actions in the growth plate because TR␣0/0 mice are growth retarded (42), whereas Fgfr3⫺/⫺ mice have skeletal overgrowth (9, 10, 49). As well as effects on chondrocyte proliferation and hypertrophy, T3 also regulates clonal expansion of chondrocyte progenitors (13), thus limiting the generation of proliferating chondrocyte populations. TR proteins are expressed in progenitor cells within the reserve zone of the growth plate (13, 16), suggesting that these effects may result


from direct T3 actions. Alternatively, they may be mediated via FGFR1 or FGFR2, both of which are coexpressed with FGFR3 and stimulated by T3 in ATDC5 cells (Fig. 3 and Ref. 25). In further support of this possibility, T3 is already known to enhance FGFR1 activity in osteoblasts (4), and FGFR1 expression is increased in growth-plate chondrocytes in TR␤PV/PV mice (3), whereas activation of either FGFR3 or FGFR1 inhibits chondrocyte proliferation in vivo (53). Thus, the effects of T3 on chondrogenesis may involve FGFR1 and FGFR2 as well as FGFR3. These considerations provide new insight into the biological interactions between T3 and FGFR signaling that regulate endochondral ossification and growth, but their mechanisms are undoubtedly complex. In addition to identification of FGFR3 as a major target for T3 in growth-plate chondrocytes, we have demonstrated that FGFR signaling during chondrogenesis primarily involves MAPK and STAT1 in ATDC5 cells because activation of STAT3, STAT5, and PLC␥ was not identified. These findings are consistent with data showing that FGFR3-activated MAPK exerts important actions on chondrocyte differentiation, whereas activation of STAT1 influences chondrocyte proliferation and survival (48). T3 potentiated FGFR signaling via MAPK in response to FGF2 and FGF18 but inhibited STAT1 responses to both ligands, whereas stimulation of ATDC5 cells with FGF9 did not elicit a downstream signaling response in the absence or presence of T3 (Figs. 5–7). Thus, modulation of FGFR activity by T3 is dependent on the downstream signaling pathway and the stimulating FGF ligand involved. The finding that T3 does not enhance FGF-induced activation of MAPK in TR␣0/0 chondrocytes (Fig. 8) suggests that TR␣ is also an important mediator of T3 action in chondrocytes, as previously shown in osteoblasts (4). Despite this, the mechanisms by which these T3 effects are mediated are unknown. Nevertheless, the binding of FGF to FGFR requires interactions with HSPGs, which contact both the receptor and the ligand to form a functional ternary complex (20, 21, 54). Modifications to HSPG structure alter the specificity and affinity of binding interactions between FGFs and FGFRs (55), indicating that extracellular matrix and cell-surface composition are critical regulators of the specificity of FGFR activation. We previously showed that thyroid status or TR␣ deletion results in altered growth-plate matrix composition and the degree of HSPG sulfation (Bassett, J. H. D., R. Swinhoe, O. Chassande, J. Samarut, and G. R. Williams, unpublished data; and Refs. 16, 42). Furthermore, the CD44 hyaluronan receptor, which is expressed in chondrocytes and heparan sulfated, is negatively regulated by T3 at the transcriptional level (56), whereas syndecan 4, a HSPG core protein also expressed abundantly in cartilage, was identified as a T3-target gene in liver (57). Thus, regulation of FGF-FGFR interactions and downstream signaling by T3 may result from T3 regulation of HSPG synthesis or structural modification (2). This possibility represents a novel and uninvestigated mechanism whereby T3 could influence cellular proliferation and differentiation during bone formation. Finally, these studies also provide a complete analysis of the various FGFR1, FGFR2, and FGFR3 isoforms expressed in ATDC5 cells during chondrogenesis. A new FGFR2 splice variant lacking the C-tail was identified in ATDC5 cells, and


we have also identified the same isoform in primary chondrocytes, osteoblastic cell lines, and primary osteoblasts (our unpublished data). The functional consequences of the loss of the C-tail in FGFR2 await characterization. In addition, the roles of FGFR1 and FGFR2 in the regulation of chondrogenesis have not been elucidated. The current studies provide a basis and the reagents required to dissect further the interactions between FGFRs and T3 in the control of endochondral ossification. Acknowledgments We are grateful to Dr. C. B. Harvey for practical advice during optimization of Western blotting and immunoprecipitation methods.


18. 19.

20.

21. 22. 23.

Received June 23, 2005. Accepted September 1, 2005. Address all correspondence and requests for reprints to: Graham R. Williams, Molecular Endocrinology Group, 5th Floor Clinical Research Building, Medical Research Council Clinical Sciences Centre, Hammersmith Hospital, Du Cane Road, London W12 0NN, United Kingdom. E-mail: [email protected]. This work was supported by an Oliver Bird Fund (Nuffield Foundation) Ph.D. Studentship (to J.C.B.) and Arthritis Research Campaign Project Grant 13932 (to G.R.W).

24. 25.

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