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Originally published In Press as doi:10.1074/jbc.M404824200 on August 13, 2004

J. Biol. Chem., Vol. 279, Issue 44, 45926-45934, October 29, 2004
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A Soluble Form of Fibroblast Growth Factor Receptor 2 (FGFR2) with S252W Mutation Acts as an Efficient Inhibitor for the Enhanced Osteoblastic Differentiation Caused by FGFR2 Activation in Apert Syndrome*

Yukiho Tanimoto{ddagger}, Masahiko Yokozeki{ddagger}, Kenji Hiura{ddagger}, Kazuya Matsumoto§, Hideki Nakanishi§, Toshio Matsumoto¶, Pierre J. Marie||, and Keiji Moriyama{ddagger}**

From the {ddagger}Department of Orthodontics and Dentofacial Orthopedics, Institute of Health Biosciences, The University of Tokushima Graduate School, Tokushima 770-8504, Japan, the Departments of §Plastic and Reconstructive Surgery and Medicine and Bioregulatory Sciences, Institute of Health Biosciences, The University of Tokushima Graduate School, Tokushima 770-8503, Japan, and the ||Laboratory of Osteoblast Biology and Pathology, INSERM U 606, Lariboisière Hospital, 75475 Paris, France

Received for publication, April 30, 2004 , and in revised form, August 12, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Apert syndrome is an autosomal dominant disease characterized by craniosynostosis and bony syndactyly associated with point mutations (S252W and P253R) in the fibroblast growth factor receptor (FGFR) 2 that cause FGFR2 activation. Here we investigated the role of the S252W mutation of FGFR2 on osteoblastic differentiation. Osteoblastic cells derived from digital bone in two Apert patients with the S252W mutation showed more prominent alkaline phosphatase activity, osteocalcin and osteopontin mRNA expression, and mineralized nodule formation compared with the control osteoblastic cells derived from two independent non-syndromic polydactyly patients. Stable clones of the human MG63 osteosarcoma cells (MG63-Ap and MG63-IIIc) overexpressing a splice variant form of FGFR2 with or without the S252W mutation (FGFR2IIIcS252W and FGFR2IIIc) showed a higher RUNX2 mRNA expression than parental MG63 cells. Furthermore MG63-Ap exhibited a higher osteopontin mRNA expression than did MG63-IIIc. The enhanced osteoblastic marker gene expression and mineralized nodule formation of the MG63-Ap was inhibited by the conditioned medium from the COS-1 cells overexpressing the soluble FGFR2IIIcS252W. Furthermore the FGF2-induced osteogenic response in the mouse calvarial organ culture system was blocked by the soluble FGFR2IIIcS252W. These results show that the S252W mutation in the FGFR2 gene enhances the osteoblast phenotype in human osteoblasts and that a soluble FGFR2 with the S252W mutation controls osteoblast differentiation induced by the S252W mutation through a dominant negative effect on FGFR2 signaling in Apert syndrome.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Apert syndrome is an autosomal dominantly inherited syndrome characterized by craniosynostosis, which results in skull deformity, and symmetric bony syndactyly of the hands and feet. Its prevalence is ~15.5/1,000,000 newborns (1) and accounts for about 4.5% of all cases of craniosynostosis (2). Mutations of the human fibroblast growth factor receptors (FGFRs)1 have been identified to be the cause of a number of craniosynostosis syndromes such as Crouzon, Pfeiffer, Jackson-Weiss, Apert, Beare-Stevenson, and Muenke syndromes (3–8). With rare exceptions, Apert syndrome is caused by one of the two missense mutations of the FGFR2 gene involving an amino acid substitution, S252W or P253R, in the linker region between the second and third extracellular Ig domains (4, 9, 10). S252W results from a C755G missense mutation and is more common than P253R caused by C758G in Apert patients (11), and each mutation shows differential effects on the phenotype of syndactyly and cleft palate in this syndrome (12, 13). Most of the Apert patients are sporadic cases and are exclusively affected by mutations arising in the paternal germ line (14). Apert mutation in the FGFR2 gene serves as a gain-of-function mutation by decreasing the dissociation rate of FGFs from FGFR2 (15, 16) as well as by evoking the ligand-dependent receptor activation. In addition to the retained ligand dependence for the receptor activation, the loss of ligand specificity of FGFR2 is also elicited by Apert mutations (16, 17).

The importance of FGFs/FGFR2 signaling in the cranial suture and limb bud development has been widely reported (18–25). It is postulated that FGFR2 signaling is essential for osteogenic cell differentiation and proliferation during the process of suture growth and closure because FGFR2 transcripts are expressed at the osteogenic fronts in developing calvarial bone (19, 21). Furthermore calvarial osteoblastic cells derived from Apert patients are reported to exhibit a differentiated osteoblastic phenotype in accordance with the clinical symptom of the premature suture ossification in the patients (26, 27). A recent study has also illustrated that the mesenchymal splice variant form of the mouse Fgfr2 (Fgfr2IIIc) acts as a positive regulator of ossification by affecting cells of osteoblastic lineage (28). These data suggest that the Apert mutation of FGFR2 may stimulate differentiation and/or proliferation in mouse or human cells, although the precise mechanisms are still not completely understood. More specifically, the biological phenotypes of the digit bone cells derived from Apert syndrome patients are totally unknown.

We have recently reported an abnormal rapid mineralization of the callus during distraction osteogenesis of the deformed thumb in an Apert syndrome patient (29). Consistent with the previous genetic and biochemical studies (26, 27), these clinical findings have raised a possibility that the S252W mutation of the FGFR2 may directly cause the unusual differentiation of the digital bone cells. Here we demonstrate that transfection of the gene for human FGFR2IIIc with the S252W mutation (FGFR2IIIcS252W) inhibits the proliferation and reciprocally enhances osteoblastic differentiation of the MG63 human osteosarcoma cell line in vitro. Furthermore the soluble FGFR2IIIcS252W lacking transmembrane and cytoplasmic domains inhibits the osteoblastic phenotype of MG63 cells overexpressing FGFR2IIIcS252W. These results provide direct evidence that activation of FGFR2IIIc caused by the S252W mutation promotes osteoblast phenotype and that a soluble form of FGFR2 with S252W mutation controls osteoblast differentiation.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Patients—All the sampling procedures were undertaken on the basis of informed consent obtained from the patients and parents and were approved by the Ethical Committee of the Tokushima University Medical Hospital. Patients 1 and 2 were male Apert syndrome patients with the germ line mutation of S252W in the FGFR2 gene. Bone samples were obtained during the distraction osteogenesis of the phalanges of the thumbs at 6 years of age in Patient 1 and at 7 years of age in Patient 2. Patients 3 and 4 were male and female patients with no major congenital anomalies except for polydactyly. Bone samples were obtained during distraction osteogenesis for the deformed thumbs at 6 years of age in Patient 3 and at 4 years of age in Patient 4. The detailed information about these patients is described in our previous report (29).

Isolation of Osteoblastic Cells from Bone Samples of the Patients— Osteoblastic cells were isolated from the digit bone samples of Patients 1–4 by collagenase digestion. The samples were minced into small pieces, washed with PBS (pH 7.3) to remove the bone marrow cells, and treated with HEPES buffer containing 1% collagenase (Wako Pure Chemical Industries, Ltd., Osaka, Japan), 25 mM HEPES (Wako), 10 mM NaHCO3, 100 mM NaCl, 1 mM CaCl2, 30 mM KCl, 1 mg/ml bovine serum albumin (Sigma), 5 mg/ml glucose, and 7.5 mM N{alpha}-p-tosyl-L-lysine chloromethyl ketone (Sigma) for 30 min at 37 °C with shaking. After incubation, the supernatant was transferred into a new tube and centrifuged at 1,200 rpm for 10 min. These procedures were repeated six times, and these pellets were gently resuspended in {alpha}-minimum essential medium ({alpha}-MEM, Sigma) supplemented with 10% fetal bovine serum (FBS, Sigma) and antibiotics (100 IU/ml penicillin and 100 IU/ml streptomycin, Invitrogen). The fractionated osteoblastic cells (F1–6) obtained by the collagenase digestion of the bone samples were seeded into a culture dish and maintained in {alpha}-MEM with 10% FBS and antibiotics at 37 °C in a humidified atmosphere of 5% CO2 and 95% air. The medium was changed every 3 days, and the cells were subcultured using 1% trypsin, EDTA solution (0.05% trypsin and 0.53 mM EDTA, Sigma) when they reached confluency. The cells from the second passage were used for the reverse transcription (RT)-PCR analysis, and the cells between the sixth and eighth passages were used for the other experiments. The cells obtained from Patients 1, 2, 3, and 4 were named ApOB1, ApOB2, HOB1, and HOB2, respectively.

RT-PCR and Restriction Enzyme Digestion Analysis—The total RNA was extracted from the cells using ISOGEN (Nippon Gene, Tokyo, Japan) according to the manufacturer's instructions. One microgram of total RNA was reverse-transcribed with an RNA PCR kit (Takara Biomedicals, Shiga, Japan), and PCR was performed for 25–30 cycles of denaturation at 95 °C for 30 s, annealing at each temperature for 30 s, elongation at 72 °C for 1 min, and final extension at 72 °C for 10 min using the oligonucleotide primers described in Table I. The amplified products were separated by electrophoresis on a 1% agarose gel. To detect the expression of mutant FGFR2 mRNA, the PCR products were digested with MboI (Takara) at 37 °C for 1 h, and the DNA fragments were separated by electrophoresis on a 3% agarose gel.


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  		TABLE I
Oligonucleotide primers used in RT-PCR

 
ALP Assay—The cells were cultured in {alpha}-MEM with 10% FBS, and the cultures were washed with PBS twice after bringing them to confluency, scraped with 50 mM Tris-HCl buffer (pH 7.7), and sonicated for 20 s. After centrifugation at 15,000 rpm for 20 min at 4 °C, ALP activity in the collected supernatant was determined with substrate incubation buffer (pH 7.4) containing 10 mM p-nitrophenyl phosphate (Sigma), 20 mM MgCl2, and 1.5 M 2-amino-2-methyl-1-propanediol (pH 10.3) (Wako). The reaction mixture was incubated at 37 °C for 30 min, and the reaction was stopped by the addition of 0.2 volume of 1 N NaOH. The hydrolysis of p-nitrophenyl phosphate was monitored at A405 nm using a multiwell spectrophotometer (Bio-Rad). p-Nitrophenol was used as the standard. The protein concentration was determined using the BCA protein assay reagent (Pierce). The enzyme activity was determined by the rate of hydrolysis of p-nitrophenyl phosphate and expressed as units/mg of protein (units = nmol of p-nitro-phenol formation/incubation min).

Matrix Mineralization—After the cells became confluent, the medium was supplied with the mineralization medium containing 10% FBS, 1 x 10–8 M dexamethasone (Sigma), 10 mM {beta}-glycerophosphate (Sigma), 50 µg/ml ascorbic acid (Sigma) in the case of ApOBs and HOBs, and the diluted mineralization medium containing 10% FBS, 1 x 10–10 M dexamethasone, 100 µM {beta}-glycerophosphate, 500 µg/ml ascorbic acid in the case of MG63 cells. The medium was changed every 3 days. The mineralized matrix was stained by alizarin red-S (AR-S, pH 4.2, Wako). Briefly, at each time point, the cells were washed twice with PBS and fixed with 70% ethanol for 60 min. The fixed cells were incubated with 40 mM AR-S for 10 min with shaking. To minimize any nonspecific staining, the cells were rinsed five times with deionized water and once with PBS for 20 min. The AR-S staining of the mineralization of the extra cellular matrix was photographed.

Construction of FGFR2IIIc and Soluble FGFR2IIIc Expression Vectors—RT-PCR amplification of FGFR2IIIc and FGFR2IIIcS252W was carried out with a specific oligonucleotide primer pair that contained the start or stop codon of FGFR2 (5'-ATGGTCAGCTGGGGTCGTTTCATCT-3' for upstream, 5'-TCATGTTTTAACACTGCCGTTTATAG-3' for downstream). The reverse transcribed total RNA prepared from ApOB1 was added to the PCR as the template. PCR was performed for 40 cycles of penetration at 95 °C for 30 s, annealing at 55 °C for 30 s, elongation at 72 °C for 3 min, and final extension at 72 °C for 10 min. The amplified products were then ligated into pGEM-T Easy vector (Promega, Madison, WI) using T4 DNA ligase, and each isolated plasmid cDNA was sequenced in both 5' and 3' orientations by the FGFR2-specific primers or vector-specific primers. To remove the stop codon of FGFR2 or gain the soluble FGFR2 lacking the transmembrane and cytoplasmic domains, the plasmid cDNA for FGFR2 with or without the S252W mutation was amplified by PCR with the specific primer pairs containing the start codon but not the stop codon of FGFR2 and the XbaI/BamHI site (5'-GCTCTAGAATGGTCAGCTGGGGTCGT-3' for upstream, 5'-CGGGATCCAAGCTGTAATCTCCTTTT-3' for downstream) or amplifying the extracellular domain of FGFR2 and with the XbaI/BamHI site (5'-CGTCTAGAATGGTCAGCTGGGGTCGT-3' for upstream, 5'-CGGGATCCGGAAGCTGTAATCTCCTTTT-3' for downstream). The amplified products were digested with XbaI/BamHI and were subcloned into the 3xFLAG CMV13 expression vector (Sigma) by the XbaI/BamHI site in the frame. The sequence of each expression vector was confirmed by sequencing in both the 5' and 3' orientations by FGFR2-specific primers or vector-specific primers, and these expression vectors were used as constructs encoding the full-length FGFR2IIIc with or without the S252W mutation (FGFR2IIIcS252W-FLAG or FGFR2IIIc-FLAG) and soluble FGFR2IIIc with or without the S252W mutation (sFGFR2IIIcS252W-FLAG or sFGFR2IIIc-FLAG).

Transient and Stable Transfection—COS-1 cells for the transient transfection and MG63 cells for the stable transfection were used in this study. These cells were kind gift from the Institute of Development, Aging, and Cancer, Tohoku University, Japan and maintained with Dulbecco's modified Eagle's medium supplemented with 10% FBS. For the transient transfection, 1 x 104 COS-1 cells were grown in a 100-mm culture dish at a density of 80% confluence and were transfected with 10 µg of expression vectors using SuperFect (Qiagen, Hilden, Germany) according to the manufacturer's instructions. The COS-1 cells transfected with the FGFR2IIIc-FLAG or FGFR2IIIcS252W-FLAG expression vectors and the culture medium from COS-1 cells transfected with the sFGFR2IIIc-FLAG or sFGFR2IIIcS252W-FLAG expression vectors (IIIc-CM or Ap-CM) were used for further analyses. For the stable transfection, MG63 cells were grown in 100-mm culture dishes at a density of 80% confluence and were transfected with 10 µg of FGFR2IIIc-FLAG or FGFR2IIIcS252W-FLAG using SuperFect (Qiagen). After 24 h of transfection, the cells were trypsinized, plated on 100-mm culture dishes, and maintained with {alpha}-MEM containing 10% FBS and 400 µg/ml G418 (Geneticin, Sigma) for 2 weeks. The resulting colonies were isolated using cloning cylinders and transferred to 96-well plates. The generated clones were used for further analyses.

Immunoprecipitation and Western Blotting—The cells were lysed in RIPA buffer (50 mM Tris-HCl, 150 mM NaCl, 1% Nonidet P-40 (Nakarai Chemicals, Ltd., Kyoto, Japan), 0.5% sodium deoxycholate (Wako)) containing protease and phosphatase inhibitors. For immunoprecipitation, 500 µg of protein extract was immunoprecipitated with the antiphosphotyrosine monoclonal antibody (4G10, Upstate Biotechnology, Lake Placid, NY) overnight at 4 °C and incubated with 60 µl of protein A-agarose (Oncogene Research Products, Boston, MA) for 2 h. The protein A-antibody-antigen complexes were washed four times with RIPA buffer, and 100 µl of the sample buffer (4% SDS, 20% glycerol, 10% 2-mercaptoethanol, 0.125 M Tris-HCl, pH 6.8, Sigma) was added and boiled, and the supernatant was used for Western blotting. Forty micrograms of the cell extract and 30 µl of the culture medium were mixed with the sample buffer, boiled, and also subjected to Western blotting. These samples were separated by SDS-PAGE, electrotransferred to a polyvinylidene difluoride membrane (Bio-Rad), and incubated with the anti-FGFR2 polyclonal antibody (Santa Cruz Biotechnology, Inc., Santa Cruz, CA), the anti-phosphotyrosine monoclonal antibody (Upstate Biotechnology), or the anti-FLAG monoclonal antibody (Sigma). Proteins were visualized by an ECL plus kit (Amersham Biosciences). The radiographic film was recorded by image scanner (EPSON, Tokyo, Japan), and the density of the bands was quantified using the image analyzing computer software NIH Image.

3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium Bromide Assay—The rate of cell growth was evaluated by 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide assay. Initially 5 x 103 cells were subcultured in a 96-well plate and then cultured for 24 h to allow the cells to adhere to the plate. After preincubation, the culture medium was changed to a serum-free medium and incubated for 24 h in the presence of either IIIc-CM or Ap-CM with FGF2 (20 ng/ml). After the indicated culture period, 10 µl of the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide solution (5 mg/ml, diluted with PBS pH 7.6) was added to each well followed by 4 h of incubation at 37 °C. The reaction was terminated by adding 100 µl of acid-isopropanol solution (0.78 mg/ml isopropanol and 0.04 N HCl) to each well. The plates were allowed to stand overnight at room temperature to dissolve the formazan crystals. The absorbance of the samples was read at 570 nm in reference to 630 nm with a multiwell spectrophotometer (Bio-Rad).

Immunocytochemistry—The cells were plated on a glass bottom chamber and grown to 80% confluence. The cells were fixed with 4% paraformaldehyde for 1 h, rinsed with PBS, and treated with 0.1% Triton X-100 for 30 min. The cells were then treated with 10% rabbit serum to block nonspecific binding and incubated with anti-FLAG monoclonal antibody. After rinsing with PBS, the cells were incubated with rhodamine-conjugated rabbit anti-mouse IgG. After washing with PBS, the cells were observed using the confocal laser microscope (Radiance 2000, Bio-Rad).

5-Bromo-2'-deoxyuridine (BrdUrd) Incorporation—S phase cells were detected using an in situ cell proliferation kit (Roche Applied Science) according to the manufacturer's instructions. Briefly the cells were grown on the glass bottom chamber for confocal laser microscopic observation or on the 100-mm culture dishes for FACS analysis up to 80% confluence. The cells were then incubated in the presence of 10 µM BrdUrd labeling reagent to facilitate the incorporation of BrdUrd into DNA in place of thymidine. For the confocal laser microscopic observations, the cells were fixed in 70% ethanol diluted in glycine buffer (50 mM glycine, pH 2.0) for 45 min at room temperature and incubated in 4 M HCl for 20 min at room temperature for DNA denaturation. For immunodetection of the incorporated BrdUrd, the cells were incubated with fluorescein isothiocyanate-conjugated anti-BrdUrd monoclonal antibodies for 45 min at 37 °C. After the administration of 1 µg/ml propidium iodide, the cells were observed by confocal microscopy (Bio-Rad). For the FACS analysis, the cells were detached by trypsin/EDTA treatment after the incorporation of BrdUrd. After the DNA denaturation, immune reaction and propidium iodide staining, the cells were analyzed by a flow cytometer (EPICS, Coulter, Miami, FL).

Mouse Calvarial Organ Culture—The isolation and culture of the mouse calvarial explants were performed by a method described previously (30). Briefly calvariae were obtained from 2-day-old C57BL/6N mice (CLEA Japan, Inc., Tokyo, Japan). After removing the skin and brain, all the calvariae were placed upside down on a bed of 1.0% Seakem GTG agarose gel (BioWhittaker Molecular Applications, Rockland, ME) in a 24-well culture dish (Iwaki, Tokyo, Japan). A thin layer of 1% agarose gel covered the calvarial explants. The agarose was dissolved in {alpha}-MEM containing 10% FBS, 100 µg/ml ascorbic acid (Sigma), 5 mM {beta}-glycerophosphate (Sigma), and antibiotics. The conditioned media were changed every 2 days. The cultured tissues were harvested on day 7, fixed with 4% paraformaldehyde, embedded in paraffin, and cut into 6-µm sections. Hematoxylin-eosin staining was performed, and the tissues were photographed.

Statistical Analysis—Data are expressed as means ± S.D. A statistical analysis was performed using the Student's t test for ALP assay or using the posthoc test for proliferation assay. All the data were reproduced in triplicate at least three times. The p values of less than 0.05 or 0.01 were considered to be significantly different in the Student's t test or the posthoc test, respectively.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
The ApOBs Show Highly Differentiated Osteoblastic Phenotype in Vitro—We previously reported a C934G point mutation that elicited an amino acid substitution, S252W, of the FGFR2 gene in Patients 1 and 2 by direct DNA sequence analysis of the genomic DNA prepared from peripheral blood (31). Therefore, we first performed RT-PCR followed by restriction enzyme digestion analysis using fraction 3 (F3) of ApOBs isolated from the two Apert syndrome patients to confirm evidence of the mutant FGFR2IIIc gene expression. The RT-PCR was carried out with the specific primers for FGFR2, and the 429-bp size-matched amplified products for FGFR2 were visualized in all samples (Table I and Fig. 1A, upper panel). We could not detect the gene expression of FGFR2IIIb splice variant form by RT-PCR using the specific primer pair. Hence it was indicated that mRNA for FGFR2IIIc variant form was predominantly expressed in HOBs and ApOBs (data not shown). Restriction enzyme digestion of the PCR products amplified from the total RNA of the HOBs yielded 166-, 145-, and 118-bp fragments, which were generated by two MboI restriction sites in the wild-type FGFR2IIIc gene sequence (Fig. 1A, middle panel). In addition, a 263-bp fragment, which was generated by the destruction of one of the MboI restriction sites due to C934G mutation, was observed in ApOBs (Fig. 1A, middle panel). These results demonstrated that ApOB1 and ApOB2 expressed both the normal and mutated forms of FGFR2IIIc mRNA. Additionally endogenous FGFR2IIIc protein was detected equally from each sample by Western blotting using anti-FGFR2 polyclonal antibody (Fig. 1A, lower panel). Next we examined the mRNA expression of the osteoblastic marker genes in HOBs and ApOBs using the specific primers as described in Table I. The expression of RUNX2 mRNA, which encodes a transcriptional activator of osteoblast differentiation, was observed in both cell fractions of ApOBs and HOBs (Fig. 1B). The mRNA expression for osteopontin and osteocalcin, which are phenotypic osteoblast markers, were detected in all the cell fractions of ApOBs (Fig. 1B). It is noteworthy that the osteopontin and osteocalcin mRNA expressions in HOBs were mostly undetectable except for F2 and F3 of HOB1 in which faint bands were observed by RT-PCR (Fig. 1B). No marked differences in the glyceraldehyde-3-phosphate dehydrogenase mRNA expression were observed in the ApOBs and HOBs (Fig. 1B). To further elucidate the characteristics of the ApOBs and HOBs, the ALP activity and mineralized nodule formation were investigated using F3 cells. The ALP activities of ApOBs were significantly higher than those of HOBs (Fig. 1C). Most strikingly, a rapid mineralized nodule formation was observed within 1–2 weeks in the ApOBs cultured in the differentiation medium, whereas the HOBs exhibited no mineralization before 3 weeks under the same conditions (Fig. 1D). These results confirmed that ApOBs with the S252W mutation of FGFR2 express a phenotype of the highly differentiated osteoblasts.



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  		FIG. 1.
Analysis of osteoblastic phenotypes in HOBs and ApOBs. A, FGFR2 mRNA and protein expression in HOBs and ApOBs and restriction enzyme analysis. The amplified products of 429 bp were detected in all samples (upper panel). The amplified PCR products were digested with the restriction enzyme MboI and then separated by electrophoresis on a 3% agarose gel. Three fragments of 166, 145, and 118 bp were generated in HOBs (middle panel, lanes 1 and 2). In ApOBs (middle panel, lanes 3 and 4), a 263-bp band was present due to the C934G mutation. A schematic diagram of the restriction enzyme analysis shows the size of each fragment generated by MboI. Endogenous FGFR2 protein was detected by Western blotting (WB) using anti-FGFR2 polyclonal antibody as described under "Materials and Methods." B, messenger RNA expression of RUNX2, osteocalcin, osteopontin, and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) in HOBs and ApOBs by RT-PCR. The size of each product is also described in Table I. The specificity of the PCR products was confirmed by DNA sequencing (data not shown). All fractions of both HOBs and ApOBs expressed RUNX2, whereas ApOBs showed higher levels of osteocalcin and osteopontin mRNA expression than HOBs. C, ALP activity in ApOBs and HOBs. Confluent cells were cultured for the indicated periods, and then ALP activity was determined as described under "Materials and Methods." ApOBs showed significantly higher ALP activity compared with HOBs when cells were cultured in {alpha}-MEM containing 10% FBS. Data were normalized to total protein concentration of cell lysates as determined with a BCA kit and described as p-nitrophenol formation/incubation time (30 min)/mg of protein. *, p < 0.05; **, p < 0.01. D, mineralization capacity of ApOBs and HOBs in vitro. Confluent cells were cultured in media supplied with 1 x 10–8 M dexamethasone, 10 mM {beta}-glycerophosphate, and 50 µg/ml ascorbic acid for the indicated period. The media were changed every 3 days until the indicated time. Mineralization was evaluated by AR-S staining as described under "Materials and Methods." Mineralization of ApOBs was clearly enhanced compared with HOBs at 1–2 weeks. W, week(s).

 
Expression of FGFR2IIIc-FLAG or FGFR2IIIcS252W-FLAG in COS-1 Cells and Their Phosphorylation—It is reported that FGFR2IIIc, a splice variant form of FGFR2, is preferentially expressed in cells of mesenchymal origin such as osteoblasts. To investigate the specific roles of this variant form with or without S252W mutation, cDNAs for FGFR2IIIc and FGFR2IIIcS252W were cloned from ApOB1 by PCR and transiently transfected in COS-1 cells with the 3XFLAG CMV13 expression vector (Fig. 2A). In a Western blotting analysis using the anti-FLAG monoclonal antibody, two bands corresponding to the unglycosylated form (the lower molecular size) and the glycosylated form (the higher molecular size) of FGFR2 were detected in the cell lysates prepared from COS-1 cells overexpressing FGFR2IIIc or FGFR2IIIcS252W (Fig. 2B) in accordance with the previous report (32). A tyrosine phosphorylation of the glycosylated form of FGFR2IIIcS252W-FLAG was more evident as compared with that of FGFR2IIIc-FLAG even without exogenous FGF administration (Fig. 2C) probably due to autocrine or paracrine effects of FGFs produced by COS-1 cells. The unglycosylated form of the endogenous FGFR2 was slightly observed in mock-transfected COS-1 cells, although its tyrosine phosphorylation was invisible during immunoprecipitation followed by Western blotting analysis (Fig. 2C).



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  		FIG. 2.
Expression of FGFR2IIIc-FLAG and FGFR2IIIcS252W-FLAG fusion proteins in COS-1 cells and their tyrosine phosphorylation. A, schematic representation of FGFRR2IIIc-FLAG and FGFR2IIIcS252W-FLAG fusion proteins. SP, signal peptide; Ig, immunoglobulin-like domain; Ab, acidic box; TM, trans-membrane domain; TK, tyrosine kinase domain. B, the expression of each FLAG fusion protein in COS-1 cells transiently transfected with FGFR2IIIc-FLAG or FGFR2IIIcS252W. Forty micrograms of the cell extract were mixed with the sample buffer, boiled, and subjected to loading. Western blotting analysis (WB) using anti-FLAG monoclonal antibody ({alpha}-Flag) showed that there was no marked difference of expression level between the fusion proteins. The two bands of the unglycosylated (lower) and glycosylated (upper) forms were observed. C, the tyrosine phosphorylation of each FLAG fusion protein in COS-1 cells transiently transfected with FGFR2IIIc-FLAG or FGFR2IIIcS252W. Each cell lysate was immunoprecipitated (IP) with anti-FGFR2 polyclonal antibody ({alpha}-FGFR2), and Western blotting was carried with anti-FGFR2 polyclonal or anti-phosphotyrosine monoclonal antibody ({alpha}-PY). FGFR2IIIcS252W-FLAG showed a significantly higher phosphorylation level than that of FGFR2IIIc-FLAG.

 
FGFR2IIIc-FLAG and FGFR2IIIcS252W-FLAG Cause Growth Arrest of MG63 Cells—To provide evidence for the functional roles of FGFR2 with S252W mutation on the osteoblastic proliferation and differentiation, we introduced the gene constructs for the fusion proteins of FGFR2IIIc-FLAG and FGFR2IIIcS252W-FLAG into MG63 cells. Western blotting using the anti-FLAG monoclonal antibody proved that three clones expressed FGFR2IIIc-FLAG (MG63-IIIc C1–C3) and four clones expressed FGFR2IIIcS252W-FLAG (MG63-Ap C1–C4) (Fig. 3A). Each clone showed a different level of the FLAG fusion protein expression (Fig. 3B). The similar immunolocalization patterns of the FLAG fusion proteins in the cytoplasm and plasma membrane were observed in the clones of MG63 (MG63-IIIc C3 and MG63-Ap C3) by confocal laser microscopy (Fig. 3C). To investigate the proliferation activities of these cells, BrdUrd incorporation was next investigated by confocal laser microscopy and FACS analysis. About 50% of the parental MG63 cells exhibited positive staining for BrdUrd under normal culture condition (Fig. 4, A and B). However, the rates of BrdUrd-positive cells in the clones of MG63-IIIc and MG-63-Ap were suppressed to less than 10% of the total cell numbers, and the degrees of the inhibition did not significantly correlate with the expression levels of the FLAG fusion proteins (Fig. 4, A and B).



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  		FIG. 3.
Expression and localization of FLAG fusion proteins in MG63-IIIc and MG63-Ap cells. A, expression of FLAG fusion proteins in MG63 cells. An equal amount of protein from MG63-IIIc and MG63-Ap was subjected to Western blotting (WB) with anti-FLAG monoclonal antibody ({alpha}-FLAG). Each clone expressed a different amount of FLAG fusion protein. B, the density of the bands was quantified by NIH Image software. C, localization of the FLAG fusion protein in MG63-IIIc-C3 and MG63-Ap-C3. Both FGFR2IIIc-FLAG and FGFR2IIIcS252W-FLAG proteins were localized in cytoplasm and plasma membrane in these cells. No staining was detectable in mock-transfected MG63 (MG63-MOCK) cells. IF, immunofluorescence.

 



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  		FIG. 4.
BrdUrd incorporation into MG63-IIIc and MG63-Ap. A, confocal microscopic observations. BrdUrd was incorporated into cells and incubated with fluorescein isothiocyanate-conjugated anti-BrdUrd antibody, and propidium iodide staining was performed for nuclear staining. BrdUrd/propidium iodide double positive cells appeared yellow. The number of BrdUrd-positive cells in MG63-IIIc-C1 and -2 and MG63-Ap-C3 and -4 decreased as compared with parental MG63 cells. B, FACS analysis. BrdUrd was incorporated into each cells, incubated with fluorescein isothiocyanate-conjugated anti-BrdUrd antibody, and analyzed by flow cytometer. Almost 50% of parental (P) MG63 cells and 60% of mock (M) cells were BrdUrd-positive. In contrast, the rate of BrdUrd-positive cells in MG63-IIIc-C1 and -2 and MG63-Ap-C3 and -4 was decreased to less than 10%.

 
Effects of Soluble FGFR2IIIc-FLAG (sFGFR2IIIc-FLAG) and Soluble FGFR2IIIcS252W-FLAG (sFGFR2IIIc-FLAG) on FGF2-induced Osteogenic Response—It is known that soluble receptors for various cytokines may function as positive or negative regulators for the biological phenomena that are implicated in the physiological and pathological circumstances. To address the question whether or not soluble forms of FGFR2IIIc with or without S252W mutation have a biological function to regulate the phenotype of osteoblastic cells, the expression vectors for the soluble forms of FGFR2IIIc and FGFR2IIIcS252W were constructed and transiently transfected into the COS-1 cells (Fig. 5A). Conditioned media were collected from the COS-1 cells transfected with the vectors for sFGFR2IIIc-FLAG and sFGFR2IIIcS252W-FLAG (IIIc-CM and Ap-CM) as well as with the empty vector alone (MOCK-CM). The levels of the sFGFR2IIIc-FLAG and sFGFR2IIIcS252W-FLAG fusion proteins in the conditioned media were similar to each other when confirmed by Western blotting using the anti-FLAG monoclonal antibody (Fig. 5B). On the other hand, endogenous soluble forms of FGFR2 were detected equally in each conditioned medium of primary culture of HOBs and ApOBs by Western blotting using anti-FGFR2 antibody recognizing the amino-terminal lesion of FGFR2 (Fig. 5C). The proliferation of MG63 cells was stimulated upon FGF2 administration by more than 3-fold in the media supplemented with 5% of the conditioned medium from the non-transfected COS-1 cells (control-CM) or MOCK-CM (Fig. 5D). However, the accelerated proliferation activity of MG63 cells was significantly reduced in the media supplemented with 5% IIIc-CM or Ap-CM, and the inhibitory effect of Ap-CM was more significant than that of IIIc-CM. Furthermore ultimate inhibition of FGF2-induced osteoblast proliferation and new bone formation in the mouse calvarial organ culture system were observed upon administration of 5% Ap-CM (Fig. 5E, d). IIIc-CM also inhibited the effect of FGF2, but the inhibitory effect was lower than that of Ap-CM, thereby osteoblast scattering on the calvarial bone surface almost completely disappeared (Fig. 5E, c).



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  		FIG. 5.
Soluble form of FGFR2IIIc-FLAG and FGFR2IIIcS252W-FLAG and its biological activity in vitro. A, schematic representation of sFGFRR2IIIc-FLAG and sFGFR2IIIcS252W-FLAG fusion proteins. SP, signal peptide; Ig, immunoglobulin-like domain; TM, transmembrane domain; TK, tyrosine kinase domain. B, FLAG fusion proteins were observed by Western blotting (WB) using anti-FLAG monoclonal antibody ({alpha}-FLAG) in the conditioned media (IIIc-CM and Ap-CM) from COS-1 cells transiently transfected with expression vectors for sFGFR2IIIc-FLAG and sFGFR2IIIcS252W-FLAG. The conditioned media prepared from non-transfected and mock-transfected COS-1 cells (C-CM and MOCK, respectively) were used as controls. There was no marked difference in the expression levels of the fusion proteins in IIIc-CM and Ap-CM. C, the expression of soluble endogenous FGFR2 in HOBs and ApOBs. Endogenous soluble FGFR2 in conditioned medium of HOB1, HOB2, ApOB1, and ApOB2 were determined by Western blotting (WB) using anti-FGFR2 polyclonal antibody recognizing the amino terminus of human FGFR2. The single band was detected from each sample in the region of 50 kDa. D, effects of IIIc-CM and Ap-CM on FGF2-induced proliferation of MG63 cells by 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide assay. The proliferation of MG63 cells was promoted by FGF2 (20 ng/ml) in the presence of 5% C-CM and MOCK, and the increased proliferation induced by FGF2 (20 ng/ml) was significantly reduced in the presence of 5% IIIc-CM and Ap-CM (**, p < 0.001). The inhibitory activity was significantly stronger in Ap-CM than in IIIc-CM (*, p < 0.01). The proliferation of MG63 cells without FGF2 administration was used for normalization. E, the effects of Ap-CM on the number of osteoblastic cells and new bone formation induced by FGF2. Calvariae obtained from 2-day-old mice were cultured in the absence or presence of FGF2, IIIc-CM, and Ap-CM for 7 days. FGF2 (20 ng/ml) increased the number of osteoblastic cells and new bone formation (b) as compared with calvariae without FGF2 treatment (a). Administration of IIIc-CM decreased the number of osteoblastic cells and new bone formation induced by FGF2 to the control level (c). Ap-CM completely inhibited the effect of FGF2, and there were no osteoblasts on the surface of the bone tissue (d). The analysis was performed three times with similar results.

 
FGFR2IIIc with S252W Mutation Stimulates Osteoblastic Differentiation of MG63 Cells, and Soluble FGFR2IIIc with S252W Significantly Inhibits This Phenomenon—Since cell proliferation and terminal differentiation are mutually exclusive in many cell types, it is postulated that the growth arrest observed in the MG63-IIIc and MG63-Ap cells might be related to the enhanced cell differentiation. Therefore, we next analyzed the changes in the osteoblastic phenotype of various MG63 transfectants. RT-PCR exhibited no expression of RUNX2 and osteopontin mRNA in MG63 cells with or without empty vector transfection at 25 and 30 cycles of amplification (Fig. 6), although RUNX2 was slightly detected at 35 cycles of PCR amplification (data not shown). Interestingly enough, RUNX2 mRNA was expressed in MG63-IIIc and MG63-Ap, while a marked expression of osteopontin mRNA was detected only in MG63-Ap (Fig. 6). When the osteoblast marker gene expression was analyzed in the presence of MOCK-CM, IIIc-CM, or Ap-CM, not only the RUNX2 mRNA expression in MG63-IIIc and MG63-Ap but also the osteopontin mRNA expression in MG63-Ap was more significantly inhibited by IIIc-CM and Ap-CM than by MOCK-CM (Fig. 6). In the mineralized nodule formation analysis visualized by AR-S staining, MG63-Ap underwent a rapid matrix mineralization with a 100-fold diluted mineralization medium at 1 week, whereas the parental or mock-transfected MG63 showed no mineralization either in the same media at 1 week (Fig. 7) or in the undiluted mineralization media at 4 weeks (data not shown). No or minimal matrix mineralization was observed in the MG63-IIIc cells in the diluted mineralization medium at 1 week (Fig. 7). The MG63-IIIc and MG63-Ap cells could not survive in the undiluted mineralization medium due to apoptosis. The matrix mineralization of MG63-Ap in the diluted mineralization medium was completely inhibited by the administration of 5% Ap-CM (Fig. 7). These data clearly demonstrate that the soluble form of FGFR2IIIc with the S252W mutation has a potent inhibitory activity for osteoblastic differentiation of MG63 cells transfected with the FGFR2IIIc gene.



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  		FIG. 6.
Expression of the marker genes for osteoblastic differentiation, and the effect of IIIc-CM and Ap-CM in MG63-IIIc and MG63-Ap. The expression of RUNX2 and osteopontin mRNA in MG63-IIIc and MG63-Ap was analyzed by RT-PCR. The expression of RUNX2 mRNA was undetectable in parental (row 1) and mock-transfected (row 2) MG63 cells and was clearly detected in MG63-IIIc (C1–C3) and MG63-Ap (C1–C4) at 25 cycles of PCR. The expression of osteopontin mRNA was limitedly observed in MG63-Ap (C1–C4) at 30 cycles of PCR. Administration of IIIc-CM (lane 3) or Ap-CM (lane 4) reduced the intensity of RUNX2 mRNA expression in both MG63-IIIc (C1–C3) and MG63-Ap (C1–C4) as well as osteopontin mRNA expression in MG63-Ap (C1–4) in comparison to administration of conditioned medium from parental MG63 cells (lane 1) or MOCK-CM (lane 2). GAPDH, glyceraldehyde-3-phosphate dehydrogenase; P, parental; M, mock-transfected.

 



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  		FIG. 7.
Mineralized nodule formation by MG63-IIIc and MG63-Ap. Confluent cells were cultured in Dulbecco's modified Eagle's medium supplemented with 10% FBS, 1 x 10–10 M dexamethasone, 0.1 mM {beta}-glycerophosphate, and 0.5 µg/ml ascorbic acid for 7 days, and the mineralization was evaluated by AR-S staining as described under "Materials and Methods." Positive staining was not observed in parental (P) and mock-transfected (M) MG63 cells. MG63-Ap (C1–C4) showed higher mineralization than MG63-IIIc-C1–3. Weak staining was observed in MG63-IIIc-C1–3. Administration of 5% Ap-CM containing soluble FGFR2IIIcS252W-FLAG completely inhibited the mineralization of MG63-Ap-C1–4.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
In this study, we investigated the role of FGFR2IIIc with the S252W mutation on osteoblast proliferation and differentiation by characterizing the phenotype of the osteoblastic cells derived from syndactylous digit bones of two Apert patients who showed extremely rapid ossification during distraction osteogenesis. Next we identified the phenotype of the MG63 cells transfected with the full-length human FGFR2 gene with the S252W mutation and characterized the biological activities of the soluble form of FGFR2IIIc with the S252W mutation to regulate the phenotypic expression of the osteoblasts. In the light of our new findings, we provide the first evidence that a soluble form of FGFR2IIIc with the S252W mutation acts as a significant and efficient inhibitor of the enhanced osteoblastic proliferation and differentiation caused by the activation of FGF/FGFR2 signaling.

It has been widely reported that stage-specific gene expression occurs during the osteogenic differentiation of the mesenchymal progenitor cells. RUNX2, which is a member of the runt family transcription factors, serves as a transcriptional activator of osteoblasts and regulates a broad spectrum of genes involved in bone and cartilage formation (33–35). In the process of osteoblastic differentiation, RUNX2 has an important role in the commitment of pluripotent mesenchymal cells into the osteoblastic lineage (36) and enhances the expression of the bone matrix genes, including osteopontin and osteocalcin (33). Consistently we found that FGFR2IIIc with S252W mutation increased RUNX2 and osteopontin mRNA expression in osteoblastic cells. This confirms that activation of FGF/FGFR2 signaling induces osteoblast differentiation markers in human osteoblastic cells (26). FGFR1 signaling was also found to induce the expression of Runx2 (37) and other osteoblast differentiation-related genes in vivo and in vitro. Fgfr2IIIc signaling is also required for the Runx2 expression during endochondral bone growth (28). In support of a role of FGFR signaling in the control of osteoblast phenotype is the recent finding that FGF/FGFR signaling stimulates DNA binding and transactivation activity of Runx2 via protein kinase C and mitogen-activated protein kinase pathways whereby osteocalcin gene expression is accelerated (38–40).

MG63 cells have been proven to be p53-deficient (41) and to have a number of typical features of the premature osteoblastic phenotype including the expression of type I collagen and low basal expression of ALP and osteocalcin (42–44). In addition, osteoblastic differentiation of MG63 cells can be promoted by the administration of 1{alpha},25-dyhydroxyvitamin D3 (42–44). Therefore, this cell line seemed a suitable model for investigating the osteoblastic differentiation. We generated three stable MG63 clones (MG63-IIIc C1–C3) overexpressing FGFR2IIIc-FLAG and four stable clones (MG63-Ap C1–C4) overexpressing FGFR2IIIcS252W-FLAG. MG63-IIIc and MG63-Ap showed less proliferative activities and differentiated phenotypes than the parental or mock-transfected MG63 cells. However, the osteoblastic phenotype of MG63-Ap was much more evident when compared with that of MG63-IIIc in terms of the activities for the late osteoblastic marker gene expression and mineralized nodule formation. This strongly suggests that FGFR2IIIc, especially the S252W mutant form, plays important roles in osteoblastic differentiation. In contrast, a stimulation of proliferation and apoptosis as well as an inhibition of differentiation was observed following the overexpression of mouse Fgfr2IIIc with S252W mutation in the osteoblastic cell line established from mouse calvaria (32). The reason for the redundancy of the functional roles of FGFR2IIIc with the S252W mutation in the osteoblastic cells still remains unclear, but one possible explanation might be the species difference in the osteoblastic cells or the difference in the cell maturational stages (25). For example, FGF signaling generally causes potent mitogenic activity and reduces the ALP activity and type I collagen expression in osteoblastic cells (45–47), whereas it increases the ALP activity and matrix mineralization in bone marrow stromal cells (48, 49) and promotes osteogenic responses in mesencephalic neural crest cells (50). On the other hand, the implantation of FGF2-soaked beads in the coronal suture results in increased osteogenic differentiation in vivo (19). Furthermore a significant decrease in the expression of the differentiation markers for osteogenic cells was observed during endochondral bone growth in Fgfr2IIIc-deficient mice (28). Although FGF/FGFR signaling may cause divergent effects on cell proliferation or differentiation dependent on the cell types or the cell maturational stages (32, 51, 52), we show here that FGFR2IIIc with S252W mutation stimulates osteoblastic differentiation in human cells. A significant growth arrest was observed in the MG63-Ap and MG63-IIIc cells regardless of the FGFR2IIIc expression levels or S252W mutation. This might be caused by a p53-independent pathway(s) because MG63 cells have been reported to show inactivation of the p53 gene (41). It is also known that p53 is a negative regulator of cyclin-dependent kinases, which are responsible for cell cycle arrest in the G1 phase (53, 54), and that FGF/FGFR signaling induced growth arrest in chondrocytes through the induction of p21WAF1/CIP1 and subsequent inactivation of cyclin E-cyclin-dependent kinase 2 (55). Taking these notions into consideration, it is suggested that the loss of p53 function and the activation of FGF/FGFR2IIIc signaling may be strongly implicated in the growth arrest of MG63 cells transfected with FGFR2IIIc or FGFR2IIIcS252W.

The ectodomains of numerous transmembrane proteins are released by proteolysis to yield soluble intercellular biological regulators (56). For example, the tumor necrosis factor-{alpha}-converting enzyme, a shedding enzyme belonging to a family of metalloproteinases, which contains a disintegrin domain (ADAM family), plays a role in the processing of cell surface proteins including tumor necrosis factor-{alpha}, the tumor necrosis factor receptor, the L-selectin adhesion molecule, and transforming growth factor-{alpha} (57). It has been suggested that protein ectodomain shedding may play an essential role in normal mammalian development (57) and the regeneration of hematopoietic tissue after radiation injury (58). Interestingly transgenic mice overexpressing a soluble form of Fgfr2IIIb, which acts as a decoy receptor, showed severe aplastic multiorgan response (59). This suggests that the soluble forms of FGFRs may act as dominant negative agents in vivo. Hanneken (60) showed that the secreted form of Fgfr1 was produced by alternative splicing and that the cleaved receptors are released by ectodomain shedding. Furthermore it has also been reported that physiological suture closure was inhibited using an adenoviral vector carrying the dominant negative Fgfr1 in the uterus (61). Therefore, we generated soluble FGFR2IIIcS252W-FLAG and FGFR2IIIc-FLAG lacking transmembrane and cytoplasmic domains as competitive inhibitors for the FGF/FGFR2 interaction. These dominant negative receptors significantly inhibited the FGF2-induced proliferation of MG63 cells, and sFGFR2IIIcS252W-FLAG showed a more potent inhibitory activity on the inhibition than did sFGFR2IIIc-FLAG presumably due to the low dissociation rate of sFGFR2IIIcS252W-FLAG from FGF2 as described previously on the membrane-bound FGFR2 with Apert mutation (15, 16). Although both sFGFR2IIIc-FLAG and sFGFR2IIIcS252W-FLAG induced inhibition of the osteoblastic marker gene expression, only sFGFR2IIIcS252W-FLAG significantly inhibited FGF-2-induced osteoblast proliferation and new bone formation in the mouse calvarial organ culture system. Finally sFGFR2IIIcS252W-FLAG almost completely inhibited mineralized nodule formation of MG63-Ap. These results demonstrate that sFGFR2IIIcS252W-FLAG has a wide effective range to abrogate the biological activities of FGF as a dominant negative agent in human and murine osteoblasts. We could detect the soluble form of endogenous FGFR2 in culture medium from HOBs and ApOBs by Western blotting using rabbit polyclonal antibody raised against the epitope corresponding to amino acids 22–101 mapping near the amino terminus of human FGFR2. We speculate that the endogenous soluble FGFR2 may have a regulatory effect on the FGF/FGFR signaling in physiological condition, and the disturbance of the balance between the soluble form and membrane-bound form receptors may lead osteoblasts to aberrant phenotypes.

In summary, the present study has proved the biological functions of FGFR2 with the S252W mutation to promote differentiation and to suppress proliferation of human osteoblastic cells, which may be implicated in the establishment of abnormal digit and cranial bone formation in Apert syndrome patients. Furthermore we show that a soluble form of FGFR2IIIc with S252W mutation has a biological function to regulate the phenotype of osteoblastic cells, providing a novel model for the regulation of the abnormal osteoblastic differentiation in Apert syndrome.


   FOOTNOTES
 
* The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

** To whom correspondence should be addressed: Dept. of Orthodontics and Dentofacial Orthopedics, Inst. of Health Biosciences, The University of Tokushima Graduate School, 3-18-15 Kuramoto-Cho, Tokushima 770-8504, Japan. Tel.: 81-88-633-7356; Fax: 81-88-633-9138; E-mail: moriyama{at}dent.tokushima-u.ac.jp.

1 The abbreviations used are: FGFR, fibroblast growth factor receptor; FGF, fibroblast growth factor; PBS, phosphate-buffered saline; MEM, minimum essential medium; FBS, fetal bovine serum; RT, reverse transcription; ALP, alkaline phosphatase; AR-S, alizarin red-S; BrdUrd, 5-bromo-2'-deoxyuridine; FACS, fluorescence-activated cell sorter; F, fraction; s, soluble; CM, conditioned medium. Back


   ACKNOWLEDGMENTS
 
We thank G. David Roodman for critical reading of this manuscript and comments on this work.



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