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Keywords:

  • craniosynostosis;
  • suture;
  • fibroblast growth factor 2 (FGF2);
  • activator protein 1 (AP1);
  • osteopontin (OP);
  • extracellular signal-regulated kinase (Erk);
  • osteoblast differentiation

Abstract

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES

Cranial sutures are an important growth center of the cranial bones, and the suture space must be maintained to permit the cranial adjustments needed to accommodate brain growth. Craniosynostosis, characterized by premature suture closure, mainly results from mutations that generate constitutively active fibroblast growth factor (FGF) receptors. FGF signaling, thus, is responsible for the pathogenesis of craniosynostosis. Even though FGF activates many different signaling pathways, the one involved in premature suture closure has not been defined. We observed that placing FGF2-soaked bead on the osteogenic fronts of cultured mouse calvaria accelerates cranial suture closure and strongly induces the expression of osteopontin, an early marker of differentiated osteoblasts. FGF2 treatment also induced fos and jun mRNAs and later increased the nuclear levels of activator protein 1 (AP1). FGF2 stimulates the expression of osteopontin by inducing expression of AP1, which then binds to its response element in the osteopontin promoter. Blocking of the Erk pathway by PD98059 suppressed the AP1 and osteopontin expression stimulated by FGF2. Coincidently, blocking of the Erk pathway also significantly retarded FGF2-accelerated cranial suture closure. Thus, the Erk pathway mediates FGF/FGF receptor–stimulated cranial suture closure, probably by stimulating synthesis of AP1 that then stimulates the differentiation of osteoblasts. Developmental Dynamics 227:335–346, 2003. © 2003 Wiley-Liss, Inc.


INTRODUCTION

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES

Cranial sutures are sites of continuous bone growth. During embryonic development and early in life, the sutures are straight bone edges separated by connective tissue called sutural mesenchyme. Thereafter, the spaces between the bone edges gradually narrow and interdigitations develop. It is imperative that the suture space is maintained in early development so that cranial adjustments to accommodate brain and facial growth can be made.

Craniosynostosis (CS) is a congenital anomaly characterized by the precocious obliteration of cranial sutures. Mutations in the genes encoding TWIST, MSX2, and fibroblast growth factor receptors (FGFR) 1–3 are known to cause CS (Jabs et al., 1993, 1994; Muenke et al., 1994; Wilkie et al., 1995; Bellus et al., 1996; El Ghouzzi et al., 1997; Howard et al., 1997). Mutations in FGFRs causing CS render these receptors constitutively active (Muenke et al., 1994; Reardon et al., 1994). Evidence showing the importance of FGF/FGFR signaling in the pathogenesis of CS is supported by the overexpression of the FGF2 gene in transgenic mice, which results in achondroplasia caused by the premature transformation of epiphyseal growth plate cartilage into bone (Coffin et al., 1995), while the disruption of this gene decreases bone formation (Montero et al., 2000). These observations suggest that FGF2 or FGFR activation strongly enhance the differentiation of osteoblasts in both the endochondral and intramembranous bone-forming processes. It is known that the interaction of FGFs like FGF2 with their FGFRs induces receptor dimerization and autophosphorylation and that this, in turn, activates multiple intracellular signal transduction pathways, including those involving mitogen-activated protein kinases (MAPKs; Chaudhary and Avioli, 1997; Debiais et al., 2001). However, the signaling pathways responsible for premature suture closure have not been defined.

MAPK pathways are a key link between membrane-bound receptors and the machinery that regulates gene expression. The three MAPK cascades in mammalian cells involve the extracellular signal-regulated kinase (Erk), the stress-activated protein kinase/c-jun N-terminal kinase (SAPK/JNK), and the p38 MAPK. Of these, the Erk pathway appears to be particularly involved in osteoblast proliferation and differentiation. Studies supporting this notion include the recent study by Lai et al. (2001), who showed that Erk is essential for osteoblast growth and differentiation as well as for osteoblast adhesion, spreading, migration, and integrin expression. The Erk pathway also appears to be involved in the stimulation of osteoblast-related gene expression resulting from an interaction between the extracellular matrix (ECM) and a cell surface integrin receptor (Takeuchi et al., 1997). Furthermore, it has been shown that the binding of growth factors such as FGF2 and platelet-derived growth factor-BB (PDGF-BB) to their receptors on osteoblast cells also activates the Erk MAPK (Chaudhary and Avioli, 1997).

One protein that may play an important part in the signaling pathway that activates osteoblast-related genes is the activator protein 1 (AP1). AP1 represents a family of transcription factors composed of heterodimers of Fos and Jun family members or homodimers of Jun family members joined together by their leucine zipper motifs. The resulting dimers can bind to DNA at a consensus AP1 binding site termed the TPA response element (TRE), thereby transactivating the expression of AP1-responsive genes (St-Arnaud and Quelo, 1998). Experiments where fos and jun expression in the bone was modulated by steroid hormone treatment or by overexpression or ablation of fos or jun genes suggest that the Fos and Jun proteins are important in the regulation of bone formation (Clohisy et al., 1992; Breen et al., 1994). This suggestion is supported by a study by McCabe et al. (1996), who showed that the developmental expression and activity of particular Fos and Jun proteins are functionally related to osteoblast maturation. It has also been reported that transgenic mice overexpressing Fra-1, a Fos protein, progressively increase their bone mass, which eventually leads to osteosclerosis of the entire skeleton (Jochum et al., 2000). This finding is due to an increase in the number of mature osteoblasts. AP1 binding sites are also present in the promoters of many developmentally regulated osteoblast marker genes, including collagenase, osteocalcin (OC), and osteopontin (OP) (McCabe et al., 1996; D'Alonzo et al., 2002; Kim et al., 2002).

Osteopontin (OP) is a major noncollagenous bone matrix protein, produced by osteoblasts, and it is abundantly expressed in mineralized bone. Previous reports showed that OP expression is found only in the cells lining the parietal bones, which are known to contain fully differentiated osteoblasts, while it is not found in the proliferating cells in osteogenic fronts (OFs; Iseki et al., 1997, 1999; Park et al., 2001). In contrast, the expression of FGFR1, 2, and 3 occurred exclusively in OFs (Iseki et al., 1997; Rice et al., 2000). These results indicate that FGFR-positive cells in OFs actively differentiate into FGFR-negative OP-positive cells in the parietal bones. The rapid expansion of OP-positive bone territory consequently results in cranial suture closure. In this context, OP is a good marker that distinguishes fully differentiated osteoblasts (in bones) from differentiating osteoblasts (in OFs) in cranial bone development.

As craniosynostosis results from the premature differentiation of cells in OFs into mature osteoblasts and is associated with constitutively activated FGFRs, it is likely that the FGFR activation stimulates osteoblast differentiation. Determining the mechanism by which FGF/FGFR signaling regulates OP expression, therefore, may reveal how FGFR mutations lead to craniosynostosis. The present study shows that the activation of Erk pathway by FGF2 stimulated AP1 expression, which in turn stimulates osteogenic differentiation of the cells, which is represented by osteopontin expression. Likewise, we also clearly demonstrate that Erk activation by FGF/FGFR signaling is crucial for premature suture closure.

RESULTS

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES

FGF2-Soaked Beads Accelerate Cranial Bone Growth and Induce OP Gene Expression in Cultured Calvaria

To determine the effect of FGF2 treatment on cranial suture closure, we implanted FGF2-soaked heparin acrylic beads over the OFs of the sagittal sutures in calvarial explants from embryonic day (E) 15 mice at a comparable developmental stage (Fig. 1A,C). Compared with the bovine serum albumin (BSA) -soaked beads (Fig. 1B), the FGF2-soaked beads considerably increased the tissue thickness of the sutural mesenchyme (asterisk) and suture closure after 48 hr was accelerated (Fig. 1D). Whole-mount in situ hybridization of the explants with a digoxigenin-labeled riboprobe for OP, a marker of fully differentiated osteoblasts, showed that the FGF2 beads also strongly stimulated the expression of OP and that this specifically occurred in the areas around the beads (Fig. 1F). In contrast, OP expression around the BSA beads was not stimulated (Fig. 1E). Specific signals were not detected by the OP sense probe (Fig. 1G).

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Figure 1. Fibroblast growth factor (FGF) 2 stimulates mouse calvarial bone growth and induces osteopontin (OP) expression. Embryonic day (E) 15 mouse calvaria were cultured for 48 hr with bovine serum albumin (BSA) -soaked beads (A,B) or FGF2-soaked beads (C,D) placed over the osteogenic fronts (OFs) of the sagittal suture. The cultured calvarial explants were then analyzed by in situ hybridization with a digoxigenin-labeled OP antisense (E,F) or sense (G) riboprobe. P, parietal bones. Arrows indicate the OFs of the parietal bones. The asterisks indicate the increase in the thickness of the sutural mesenchyme around the FGF2 beads. Scale bars = 1 mm in A (applies to A–D), in E (applies to E–F).

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FGF2-Induced OP Gene Expression Is not an Immediate Response, as it Requires New Protein Synthesis

We also assessed whether mouse calvarial osteoblast-like cells (MC3T3-E1), mouse bone marrow stromal cells (ST2), and rat osteosarcoma cells (ROS17/2.8) express OP in response to FGF2 treatment (Fig. 2A) by performing Northern blot analysis on RNA extracts with an OP cDNA probe (Kim et al., 2002). We found that FGF2 stimulated significantly OP gene expression all three cell types. In addition, FGF2 treatment induced other bone marker genes such as type I collagen and fibronectin in MC3T3-E1 cells (Fig. 2B). To determine the molecular mechanism by which FGF/FGFR signaling induces osteoblast differentiation, we further assessed the FGF2-mediated regulation of OP gene expression as an early marker of differentiated osteoblasts. OP gene expression is found only in the lining cells of parietal bones, which are known to be differentiated osteoblasts (Park et al., 2001), and OP has been established as a marker gene indicating differentiated osteoblasts in the skull vault by other groups (Iseki et al., 1997; Eswarakumar et al., 2002). As FGF2-stimulated OP gene expression was particularly high in MC3T3-E1 and ST2 cells (Fig. 2A), we used these two cell types in our further experiments. To examine the dosage effect of FGF2 on OP gene expression, ST2 cells were treated with various concentrations of FGF2 and OP expression was analyzed by Northern blotting (Fig. 2C). OP expression was stimulated even when the cells were treated with FGF2 concentrations as low as 0.1 ng/ml and increased in a dose-dependent manner up to 10 ng/ml, after which OP expression reached a plateau. When ST2 cells treated with 10 ng/ml FGF2 were harvested at various time points after treatment and their RNA extracts examined for OP expression, it was found that FGF2-stimulated OP gene expression was initiated 6 hr after treatment. OP expression continued until 24 hr after FGF2 treatment and, thereafter, returned to the basal level at 48 hr (Fig. 2D). The delay in OP expression suggests that new protein synthesis is required after FGF2 treatment. To test this issue, ST2 cells were pretreated with cycloheximide, a protein synthesis inhibitor, and then exposed to FGF2. The pretreatment with cycloheximide completely abrogated FGF2-induced OP gene expression (Fig. 2E), indicating that FGF2-induced OP expression requires new protein synthesis. To rule out the toxicity of cycloheximide in this experiment, we performed a dimethyl thiazolyldiphenyltetrazolium bromide (MTT) assay. The treatment of 10 μg/ml cycloheximide has little toxicity on these cells (Fig. 2F).

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Figure 2. Fibroblast growth factor (FGF) 2 stimulates osteopontin (OP) gene expression in osteoblastic cells, and new protein synthesis is required for the stimulation. MC3T3-E1 cells (MC;), ST2 cells, and ROS17/2.8 cells (ROS; A), MC3T3-E1 cells (B), or ST2 cells (C) were cultured for 3 days and, in the final 24 hr, were treated with 10 ng/ml FGF2. D: Confluent ST2 cells were treated with 10 ng/ml FGF2, and cells were harvested at the indicated time points. E: ST2 cells were cultured for 3 days and treated for final 24 hr with or without FGF2 alone or in combination with cycloheximide (CHX; 10 μg/ml). F: A dimethyl thiazolyldiphenyltetrazolium bromide (MTT) assay was performed for testing the toxicity of CHX. The expression of OP, fibronectin, and type I collagen was determined by Northern blot analysis by using 10 μg of total RNA. That equal amounts of RNA loaded in each lane was visualized by hybridizing 18S ribosomal RNA (18S) in the same blot.

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AP1 Components Are Induced by FGF2, Resulting in Increased AP1 Binding With the TRE Motif on the OP Promoter

Several reports indicate that FGF2 induces the expression of AP1 components (Fos and Jun family members) and that AP1 plays important roles in osteoblast differentiation (McCabe et al., 1996; Newberry et al., 1997; Varghese et al., 2000). Moreover, as the OP promoter contains an AP1 binding site (Kim et al., 2002), it is likely that AP1 is involved in the regulation of OP gene transcription. To assess this involvement, ST2 cells were treated with FGF2 and their RNA extracts were then examined by Northern blotting for the expression of various fos- and jun-related mRNAs, namely, c-fos, FosB, Fra-1, Fra-2, c-jun, Jun-B, and Jun-D. All of the fos- and jun-related mRNAs we examined were up-regulated immediately after FGF2 treatment (Fig. 3). FosB, Fra-1, Fra-2, c-jun, JunB, and JunD mRNA levels peaked between 30 and 60 min after FGF2 treatment and then disappeared. c-fos expression was the briefest, as it peaked at 30 min and then disappeared. JunD and Fra-2 mRNA, which are constitutively expressed by ST2 cells, showed the weakest response to FGF2.

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Figure 3. Fibroblast growth factor (FGF) 2 stimulates the expression of fos- and jun-related genes. Confluent ST2 cells were treated with 10 ng/ml FGF2 for the indicated time periods, after which 10 μg of their total RNA was analyzed by Northern blot hybridization using cDNA probes for c-fos, FosB, Fra-1, Fra-2, c-jun, JunB, and JunD. That equal amounts of RNA were loaded in each lane was visualized by hybridizing 18S ribosomal RNA (18S) in the same blot.

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To determine whether FGF2-induced AP1 mRNA expression is translated into AP1 protein and that this could directly up-regulate OP gene expression, we performed an electrophoretic mobility shift assay (EMSA) using a radiolabeled oligonucleotide representing the AP1 binding site (TRE) in the proximal OP promoter (OP-TRE; Kim et al., 2002). For this purpose, MC3T3-E1 cells were cultured in the presence or absence of FGF2 for the indicated time periods and EMSA was performed on their nuclear extracts. AP1 binding to the probe was strongly enhanced by FGF2 treatment (Fig. 4A). The binding of AP1 increased strongly 3–6 hr after FGF2 treatment and was maintained at lower levels until 12 hr after treatment (Fig. 4A). AP1 bound to the wild-type probe but did not bind to a mutant TRE sequence (Fig. 4B, lanes 8 and 9). That the AP1 binding to the wild-type OP-TRE oligonucleotide was specific was indicated by competition assays, as the FGF2-enhanced AP1 binding activity (Fig. 4B, lanes 2 and 3) was completely competed out by a molar excess of unlabeled wild-type TRE oligonucleotides (Fig. 4B, lanes 4 and 5) but was not affected by the mutant competitor (Fig. 4B, lanes 6 and 7). To identify the members of AP1 superfamily involved in the mobility shift, we performed supershift assay with specific antibodies. Incubation of nuclear extracts, which were obtained 6 hr after the treatment with FGF2, with antibodies against c-Jun led to retardation of the migration of AP1/DNA complex in EMSA (Fig. 4C, lane 6). However, when nuclear extracts were incubated with specific antibodies against c-Fos, FosB, JunB, and JunD, supershifted bands were not detected but the remaining AP1/DNA band intensity was markedly reduced compared with the sample in which no antibody was added (Fig. 4C, compare lanes 4, 5, 7, and 8 with lane 3). Because the absence of a supershifted band in the presence of each specific antibody may derive from the destabilization of these proteins with DNA by each antibody as a result of antibody competition with the oligonucleotide probe or alteration of the conformation of AP1 components by each antibody (Lai and Cheng, 2002), the reduction of the AP1/DNA band intensity in the FGF2-treated sample by specific antibodies against c-Fos, FosB, JunB, and JunD appeared to be specific. To further confirm the up-regulation of the activity of Fos family members by FGF2, we used an anti–pan-Fos antibody, which recognizes the common domain of the Fos family members and can interact with a vast numbers of Fos/Jun dimmers, in EMSA. As shown in Figure 4C, lane 12, this antibody clearly supershifted the AP1/DNA bands in the gel, leaving behind nondetectable AP1/DNA bands in the original location. In contrast, the reduction of AP1/DNA band intensity was shown in the reaction with anti–pan-Jun antibody (Fig. 4C, lane 13). These results indicate that most of the AP1 family members may be involved in the FGF2 induction of OP gene expression in osteoblast. To clarify the discrepancy in timing between AP1 binding to the OP promoter and maximum FGF2-stimulated OP mRNA levels, we checked the stability of OP mRNA. The treatment with actinomycin D, an inhibitor of RNA synthesis, showed that OP mRNA synthesized by FGF2 treatment was stably maintained for an additional 12 hr even in the absence of new RNA synthesis. The presence of FGF2 with actinomycin D did not affect the stability of OP mRNA (Fig. 4D).

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Figure 4. The fibroblast growth factor/receptor (FGF/FGFR) signal enhances the binding of activator protein 1 (AP1) to the TPA response element (TRE) in the osteopontin (OP) promoter. Confluent MC3T3-E1 cells were treated with 10 ng/ml FGF2 for the indicated time periods, and their nuclear extracts were prepared. A: Nuclear extracts were incubated with a radiolabeled oligonucleotide representing the AP1 binding site in the murine OP promoter (OP-TRE). B: Competition was performed with 100- or 1,000-fold molar excess of unlabeled OP-TRE (lanes 4, 5) or mutant OP-TRE (mut; lanes 6, 7) probes. Nuclear extracts treated with or without FGF2 for 6 hr were used in the competition assay. Competitors were incubated with nuclear extracts for 15 min before the addition of labeled probe. In lane 1, no protein was added. C: Extracts were preincubated for 30 min with each antibody against c-Fos, FosB, c-Jun, Jun B, JunD, pan-Fos, or pan-Jun (lanes 4, 5, 6, 7, 8, 12, and 13) before the addition of labeled OP-TRE probe. In lanes 1 and 9, no protein was added. D: Confluent MC3T3-E1 were treated with or without FGF2 for 12 hr to stimulate the level of OP mRNA maximally, then washed twice with phosphate-buffered saline. A total of 0.1 μg/ml actinomycin D (ActD) was added in the presence or absence of FGF2 for the indicated time periods. OP expression was determined by Northern blot analysis by using 10 μg of total RNA. That equal amounts of RNA were loaded in each lane was visualized by hybridizing 18S ribosomal RNA (18S) in the same blot. Arrowheads indicate free probes. S.S means supershifted bands. The asterisks indicate nonspecific binding on these probes.

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FGF2-Stimulated AP1 Expression Is Specifically Mediated by the Erk Pathway

We investigated which of the signaling pathways stimulated by FGF2 are involved in the regulation of AP1 expression. FGF binding to FGFR is known to stimulate receptor dimerization, tyrosine phosphorylation, and the activation of multiple signal transduction pathways, including those involving Ras, Erks, src and p38 MAP kinases, phospholipase C (PLC), and protein kinase C (PKC) (Kuo et al., 1997; Maher, 1999). To investigate this issue, we treated MC3T3-E1 cells with pathway-specific inhibitors before exposing them to FGF2. Northern blot analysis on the RNA extracts of these cells with labeled probes for c-fos, FosB, Fra-1, Fra-2, c-jun, JunB, and JunD revealed that the inhibition of the Erk pathway by PD98059 significantly blocked all the fos- and jun-related gene expression stimulated by FGF2, even though there are slight differences in the blocking levels of these genes (Fig. 5A). Densitometry tracing revealed the following percentage reduction in expression: c-fos, 31%; FosB, 19%; Fra-1, 57%; Fra-2, 80%; c-Jun, 53%; JunB, 51%; JunD, 72%. In contrast, the inhibition of the p38 MAPK and PKC pathways by SB203580 and calphostin C, respectively, had little effect although fra-2 is significantly suppressed by calphostin C (Fig. 5A). To verify that the inhibitors were effective on designated pathways, we examined the state of phosphorylation of Erk1/2 and p38 MAPK by Western blot analysis. FGF2-stimulated phosphorylation of Erk1/2 was effectively down-regulated by PD98059 (Fig. 5B). However, the effect of SB203580 was not detected, because the phosphorylation of p38 MAPK did not occur by FGF2 stimulation, although positive control was detected by a specific antibody against phospho-p38 MAPK (Fig. 5B). These results suggest that p38 MAPK pathway is not involved in the FGF2-stimulated AP1 expression in this experimental condition, although previous reports showed FGF2-stimulated phosphorylation of p38 MAPK (Tanaka et al., 1999; Kozawa et al., 1999). EMSA on nuclear extracts of MC3T3-E1 cells showed that blocking of the Erk pathway abolished the FGF2-stimulated binding of AP1 to the OP-TRE oligonucleotide (Fig. 5C).

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Figure 5. Extracellular signal-regulated kinase (Erk) activation is involved in fibroblast growth factor (FGF) 2-induced fos and jun expression. A: MC3T3-E1 cells were cultured for 3 days and treated for the final 0.5–1 hr with FGF2. One hour before the cells were exposed to FGF2, pathway-specific inhibitors (Erk, 50 μM PD98059; p38, 25 μM SB203580; and protein kinase C, 0.5 μM calphostin C) were added. Total RNA (10 μg) was analyzed by Northern blot hybridization using cDNA probes for c-fos, FosB, Fra-1, Fra-2, c-jun, JunB, and JunD. That equal amounts of RNA were loaded in each lane was visualized by hybridizing with 18S ribosomal RNA (18S). B: MC3T3-E1 cells were cultured for 3 days and treated for 15 min with FGF2. One hour before the cells were exposed to FGF2, PD98059 or SB203580 was treated. Whole cell lysates were prepared and subjected to Western blot analysis (30 μg/lane). Blots were incubated with anti–phospho-Erk1/2 antibody (p-Erk1/2) or anti–phospho-p38 antibody (p-p38). That equal amounts of protein were loaded in each lane were shown by the expression levels of Erk1/2 and p38, respectively. The lysate of U937 cells treated with 1,25(OH)2D for 24 hr was used as positive control for phospho-p38. C: Confluent MC3T3-E1 cells were treated with FGF2 for 6 hr. One hour before the cells were exposed to FGF2, PD98059 was added. An electrophoretic mobility shift assay was then performed by incubating the nuclear extracts of the cells with the labeled osteopontin TPA response element (OP-TRE) probe. The asterisk indicates nonspecific binding.

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Blocking the Erk Pathway Abrogates FGF2-Stimulated OP Expression and Retards FGF2-Stimulated Cranial Suture Closure

To confirm that the Erk pathway mediates FGF2-stimulated OP gene expression, MC3T3-E1 cells were treated with PD98059 for 1 hr before FGF2 treatment and RNA extracts of the cells were assessed by Northern blot analysis by using an OP probe. Blocking of the Erk pathway indeed significantly suppressed FGF2-induced OP gene expression (Fig. 6).

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Figure 6. Inhibiting the extracellular signal-regulated kinase (Erk) pathway blocks fibroblast growth factor (FGF) 2-stimulated osteopontin (OP) gene expression. MC3T3-E1 cells were cultured for 3 days and treated with FGF2 for the final 24 hr. The cells were pretreated with PD98059 (50 μM) 1 hr before FGF2 was added. Total RNA (10 μg) was analyzed by Northern blot hybridization using a cDNA probe for OP. That equal amounts of RNA were loaded in each lane was visualized by hybridizing with 18S ribosomal RNA (18S).

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To evaluate the effect that inhibiting the Erk pathway would have on cranial suture closure, E15 mouse calvarial explants were pretreated with PD98059 for 2 hr before the application of FGF2-soaked beads on the OFs of the parietal bones (denoted here as PD/FGF2 treatment). A slight difference in the mating time greatly affected the initial width of the sagittal suture. To control for such variability, the littermates from individual pregnant mice were divided into three groups, namely, BSA, FGF2, and PD/FGF2. Although slight variations in the increases in the tissue volume and the speed of the suture closure within each group were also observed (Table 1), the increase in the tissue volume of the sutural mesenchyme (Fig. 7F, asterisk) and the acceleration of suture closure induced by applying the FGF2 beads were consistently blocked by the inhibition of the Erk pathway (Fig. 7A–I,M, page 3; Table 1). The inhibition of the Erk pathway also abrogated the OP gene expression around the FGF2 beads (Fig. 7J–L). Statistical analysis on the width of the sagittal sutures (i.e., the distance between the two OFs of the parietal bones) revealed that, at 24 hr, the width of the suture in BSA-, FGF2-, and PD/FGF2-treated calvaria had decreased by 30.2%, 53.4%, and 35.6% relative to the initial width, respectively. A similar situation was observed 48 hr after the beads were applied; by then the widths of the sutures had decreased by 45.1%, 66.6%, and 49.7% relative to the initial width (Table 1). These data showed that suture closure in the PD/FGF2-treated calvaria occurred almost at the same rate as that in the BSA-treated calvaria, whereas the sutures in the FGF2-treated calvaria closed considerably faster. The speed of suture closure in the PD/FGF2-treated calvaria did not differ significantly from that of the BSA-treated calvaria (P > 0.05), but both differed significantly from the suture closure speed of the FGF2-treated calvaria (P < 0.0001). These results indicate that Erk blocking by PD98059 almost completely suppressed FGF2-stimulated cranial suture closure.

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Figure 7. Inhibiting the extracellular signal-regulated kinase (Erk) pathway retards the cranial suture closure accelerated by fibroblast growth factor (FGF) 2. Mouse calvarial explants (embryonic day 15) were cultured for 48 hr with beads soaked with either FGF2 (D–F,K) or bovine serum albumin (BSA; A–C,J). Some explants were pretreated with PD98059 (50 μM) for 2 hr before the FGF2-soaked beads were applied (G–I,L). Arrows indicate the osteogenic fronts of the sagittal suture. J–L: The osteopontin (OP) gene expression was determined by hybridizing the explants with the 35S-labeled OP antisense probe. Red spots indicate OP expression. The asterisk indicates the increase in the thickness of the sutural mesenchyme. M: The results shown in A–I and in Table 1 are presented as a graph. Filled circles, BSA beads; filled squares, FGF2 beads; open triangles, PD98059 pretreatment before the application of the FGF2 beads. Scale bars = 1 mm in A (applies to A–I), 200 μm in J (applies to J–L).

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Table 1. Effect of the Inhibition of the Erk Pathway on FGF2-Accelerated Cranial Suture Closurea
Time (hr)Groups (No. of explants)
BSA (44)FGF2 (36)PD/FGF2 (30)
Mean width (range)% decrease from initial widthMean width (range)% decrease from initial widthMean width (range)% decrease from initial width
  • a

    The calvarial explants were magnified (×70) under a stereomicroscope and changes of suture width were statistically analyzed by repeated measures ANOVA. FGF, fibroblast growth factor; BSA, bovine serum albumin; PD, PD98059.

  • b

    The width between two nearest tangential lines of opposing osteogenic fronts of parietal bones (mean ± SD mm).

  • c

    The suture closure in FGF2-treated explants at 24 and 48 hr occurred significantly faster (P = 0.0001) than that in BSA- or PD/FGF2-treated explants at the same time points.

00.703 ± 0.159b 0.673 ± 0.194 0.726 ± 0.178 
 (0.380–1.079) (0.317–1.047) (0.440–1.111) 
240.491 ± 0.14630.20.314 ± 0.16053.4c0.468 ± 0.11335.6
 (0.158–0.777) (0.079–0.650) (0.158–0.634) 
480.386 ± 0.16245.10.225 ± 0.15169.6c0.365 ± 0.13349.7
 (0.111–0.746) (0.000–0.634) (0.0–0.6) 

DISCUSSION

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES

FGF/FGFR Signaling Accelerates Cranial Suture Closure and Induces the Expression of OP

FGF/FGFR signaling appears to be involved in determining the shape and size of the skeleton during its development (Muenke and Schell, 1995). The importance of these molecules in these processes is supported by the association of several FGFR mutations with dominantly inherited human skeletal disorders (Jab et al., 1994; Reardon et al., 1994; Muenke and Schell, 1995). For example, mutations in FGFR1, FGFR2, and FGFR3 that constitutively activate these receptors are linked to several syndromes, including craniosynostosis, which is characterized by the premature fusion of the cranial sutures, and achondroplasia, which is characterized by shortened long bones. The sutural space in the calvaria and the epiphysis in the long bone are both centers of bone growth. In these areas, stem cells continuously provide osteoprogenitor cells that facilitate long bone growth in childhood and puberty and calvarial expansion to accommodate brain growth. It thus appears that constitutively active mutations in FGFR molecules induce the rapid differentiation of osteoprogenitor cells or preosteoblasts in the bone growth areas and, thereby, accelerate the obliteration of these areas. However, little is known about the underlying molecular mechanisms by which FGF/FGFR signaling induces osteoblast differentiation.

We observed that, when FGF2-soaked beads were implanted over the OFs of the parietal bones in cultured E15 calvaria, the subsequent activation of FGFR signaling accelerated cranial suture closure and induced the expression of the OP gene (Kim et al., 1998; Iseki et al., 1997). Although we focused on the regulation of OP gene expression in the present study, we showed that other bone marker genes such as type I collagen and fibronectin were induced by FGF2 treatment. Furthermore, Runx2, an essential transcription factor for osteoblast differentiation, was also stimulated by FGF/FGFR signaling (Zhou et al., 2000; Kim et al., 2003). These results strongly suggest that FGF/FGFR signaling increases osteoblast differentiation, leading to the acceleration of cranial suture closure.

We observed that the areas around beads increased in thickness in our organ culture experiments, indicating that FGF/FGFR signaling also stimulates cell proliferation. It corresponds with earlier observations that the application of FGF4 beads on the sutural mesenchyme or osteogenic fronts commonly stimulated cell proliferation around beads (Kim et al., 1998). In addition, it is noteworthy that FGF4 beads on the osteogenic fronts accelerated the suture closure, but the beads on the sutural mesenchyme did not (Kim et al., 1998). These observations indicate two important points: first, accelerated cranial suture closure cannot be explained solely by the stimulated cell proliferation induced by FGF/FGFR signaling. A rapid differentiation into OP-positive osteoblast by the signaling must follow. Second, to accelerate the cranial suture closure, FGF ligands should be placed on the committed preosteoblasts in osteogenic fronts in which FGFR1–3 are highly expressed (Rice et al., 2000). The notion also corresponds well with the pathogenesis of craniosynostosis caused by constitutive active mutations of FGFR1–3.

AP1 Is Required for FGF/FGFR-Stimulated OP Gene Expression

When we measured OP mRNA levels in osteoblastic cells over time after FGF2 treatment, we observed that OP mRNA became detectable only 6 hr later. Pretreatment of the cells with an inhibitor of protein synthesis revealed that protein synthesis is needed for OP gene expression, indicating that the increase in OP mRNA levels is not directly due to FGF/FGFR signaling. These observations suggest that newly synthesized transcription factors are involved in converting the FGF/FGFR signal into OP gene expression. We suspect that one of these factors may be AP1, because AP1 is known to be induced by a wide range of osteogenic stimuli (Whitmarsh and Davis, 1996), and AP1 binding sites are believed to be involved in the control of various osteoblast-specific genes, including type I collagen, OP, osteonectin, and OC (Rodan and Noda, 1991; Stein et al., 1996, Kim et al., 2002). When we treated osteoblastic cells with FGF2 and examined their expression of the components that constitute AP1, we found that all the AP1 components were indeed expressed within 15–90 min after FGF2 treatment. Furthermore, most components of the AP1 family participated in the formation of FGF2-stimulated AP1 complex on OP promoter region. These results strongly suggest that the AP1 transcription factor mediates FGF2 stimulation of OP gene expression even if we could not provide evidence that AP1 is directly involved in the accelerated cranial suture closure induced by FGF2 signaling. Further studies are required for verifying the direct involvement of AP1 in cranial suture closure.

The induction of the fos and jun transcripts by FGF2 treatment was only transient; therefore, the question is, How can this early spike of AP1 transcription induce OP expression, which occurs much later? We postulated that, although translation of AP1 mRNA occurs rapidly, thereby depleting the AP1 mRNA pool, the resulting AP1 protein is maintained for a much longer period. This notion is strongly supported by the results of our electrophoretic mobility shift assay. FGF2 treatment of the cells greatly increased the binding of AP1 to the OP-TRE probe 3–6 hr after FGF2 treatment was initiated. This binding continued to be observed, albeit at lower levels, right up to 12 hr after treatment started. However, there appears to be a contradiction; AP1 complex formation disappears by 12 hr but FGF2-induced OP expression was sustained for 24 hr. This discrepancy can be explained by the observed stability of the OP mRNA.

Our present study focused only on the role of AP1 in the regulation of osteoblast differentiation. However, AP1 can interact with other transcription factors, including CREB/ATF, NFκB, Runx2, and nuclear hormone receptors, thereby considerably expanding the repertoire of genes it can control (Karin et al., 1997; Hess et al., 2001; D'Alonzo et al., 2002). These aspects mean that characterization of other transcription factors that are bound with AP1 in response to FGF2 stimulation may assist in understanding AP1-mediated OP gene regulation during osteoblast differentiation and cranial suture closure. As a strong candidate, Runx2 may be involved, because its expression and transcriptional activity are also stimulated by FGF/FGFR signaling (Zhou et al., 2000; Kim et al., 2003) and it regulates the transcription of osteoblast marker genes, including OP and OC (Ducy et al., 1997). Xiao and coworkers (2000, 2002) showed that FGF-stimulated phosphorylation and transcriptional activity of Runx2 are mediated by Erk activation. Also, protein–protein interactions between AP1 and Runx2 have been reported (Hess et al., 2001; D'Alonzo et al., 2002). These reports strongly suggest the importance of posttranslational activation of the transcription factors.

Erk1/2 MAPK Pathway Mediates FGF2-Stimulated Cranial Suture Closure

Extracellular signals are transmitted to the nucleus in a variety of ways by activating kinases. FGF binding to FGFR induces receptor dimerization, intrinsic tyrosine phosphorylation, and activation of multiple signal transduction pathways, including those involving MAPKs, Src, PLCγ, and PKC (Kuo et al., 1997; Maher, 1999). Recent studies demonstrated that PKC and Src-kinase pathways mediate FGF2 stimulation in proliferation, differentiation, and apoptosis of osteoblasts (Lemonnier et al., 2000; Debiais et al., 2001; Marie et al., 2002; Kim et al., 2003). Of interest, when we pretreated osteoblastic cells with the selective Erk inhibitor PD98059, FGF2-stimulated fos and jun mRNA levels and AP1 binding to the OP-TRE probe was dramatically suppressed, resulting in the decrease in OP expression. Inhibitors of other pathways had little effect. Especially, p38 MAPK was not phosphorylated by FGF2 treatment, although Kozawa and coworkers (1999) showed the phosphorylation of p38 MAPK by FGF2 in the same cell line (MC3T3-E1). Blocking Erk pathway by PD98059 significantly suppressed FGF2-induced AP1 components and OP gene expression, but this suppression was incomplete. Two possibilities could be suggested for the incomplete suppression: one is the incomplete blocking of the Erk pathway by PD98059, which was also observed in Western blot analysis, and the other is the involvement of other signaling pathways.

Pretreatment of cultured calvarial explants with the Erk inhibitor significantly blocked the FGF2-accelerated suture closure and the increase in cell volume around FGF2 beads. Thus, these results suggest that the Erk1/2 pathway specifically mediates FGF2-stimulated AP1 expression and is important in the pathogenesis of craniosynostosis. In addition, considering that Erk stimulates transcriptional activity of Runx2 (Xiao et al., 2000) but that Erk activation by FGF2 does not support Runx2 expression (Kim et al., 2003), we suggest that Erk might be involved in the regulation of Runx2 activity by a posttranslational mechanism. First, there is a possibility of a cross-talk between the Erk and PKC pathway. For example, FGFR-stimulated Erk could enhance the transcriptional activity of Runx2 protein through the PKC pathway or vice versa. Second, there is a possibility that Erk-induced AP1 physically interacts with Runx2 protein, which might be crucial for the activation of Runx2 protein by FGF/FGFR signaling. Thus, further study would be required to clarify this issue.

Collectively, the findings of the present study strongly suggest that AP1 transcription factors are critical mediators of FGF/FGFR signaling in osteoblast differentiation. More importantly, we provide direct evidence that the Erk pathway mediates precocious calvarial bone growth by activated FGF/FGFR signal.

EXPERIMENTAL PROCEDURES

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES

Materials

Recombinant human FGF2 was purchased from Promega (Madison, WI). Calphostin C, cycloheximide, proteinase K, actinomycin D, MTT, and BSA were obtained from Sigma Chemical Company (St. Louis, MO). Zeta-probe blotting membranes and the Bradford protein assay kit were acquired from Bio-Rad (Hercules, CA). α-MEM, RPMI1640, and Dulbecco's modified Eagle's medium (DMEM) were purchased from GibcoBRL (Grand Island, NY). Fetal bovine serum and ExpressHyb hybridization solution were obtained from HyClone (Logan, UT) and Clonetech (Palo Alto, CA), respectively. SB203580 and PD98059, purchased from Tocris (Ballwin, MO), were prepared as 25 and 50 mM stock solutions in dimethyl sulfoxide, respectively. Antibodies against c-Fos (sc-7202X), FosB (sc-7203X), c-Jun (sc-45X), JunB (sc-46X), JunD (sc-74X), pan-Fos (sc-253X), pan-Jun (sc-44X), ERK (sc-94), p-ERK (sc-7383), and p38 (sc-535) were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). Antibody against phospho-p38 was from New England Biolabs (Beverly, MA).

Cell Culture

Cells were cultured as previously described (Kim et al., 2002). Cells were washed twice with phosphate-buffered saline (PBS) and then treated for 24 hr with 10 ng/ml FGF2 in serum-free medium supplemented with 0.2% BSA. In some experiments, the cells were pretreated for 1 hr with 0.5 μM calphostin C (a PKC inhibitor), 50 μM PD98059 (a MEK1/2 specific blocker), or 25 μM SB203580 (a p38- MAPK-specific inhibitor).

RNA Preparation and Northern Blot Analysis

Total RNA was prepared and Northern hybridization was performed as previously described (Kim et al., 2002). cDNA probes were labeled with [α-32P]dCTP by using the Megaprime DNA labeling system kit (Amersham Pharmacia Biotech, UK). The cDNAs for fos and jun family members were provided by Dr. MaCabe (MaCabe et al., 1996).

Preparation of Nuclear Extracts and EMSA

Nuclear extracts were prepared as previously described (Kim et al., 2002). For electrophoretic mobility shift assays, previously characterized wild-type and mutant TRE oligonucleotides corresponding to the −80/−63 segment of the mouse OP promoter were used (Kim et al., 2002). Probes were labeled with [α-32P]dCTP and 5 fmol of the labeled probe was incubated with 10 μg of nuclear extract for 30 min at room temperature in the presence of 1× gel shift binding buffer containing 15 mM Tris, pH 7.5, 1 mM MgCl2, 0.5 mM ethylenediaminetetraacetic acid (EDTA), 0.5 mM dithiothreitol (DTT), 50 mM NaCl, 10% glycerol, and 1 μg poly (dI-dC). Competition was performed with a 100- to 1,000-fold molar excess of unlabeled TRE. Antibodies against c-Fos, FosB, c-Jun, JunB, JunD, pan-Fos, and pan-Jun were preincubated with nuclear extracts and binding buffer for 30 min at 4°C before the addition of probe. The reaction mixtures were then separated on a 5% nondenaturating polyacrylamide gel.

Western Blot Analysis

Cells were lysed in 20 mM Tris-HCl pH 7.5, 150 mM NaCl, 0.5% Triton X-100, containing protease inhibitors such as phenylmethyl sulfonyl fluoride and aprotinin. Whole cell lysates were resolved by sodium dodecyl sulfate-polyacrylamide gel electrophoresis and electro-transferred to a polyvinylidene difluoride membrane. After transfer, Western blot analysis was performed as previously described (Kim et al., 2003). The signal was detected by the ECL Plus kit (Amersham Pharmacia Biotech, UK). Antibodies against phospho-Erk1/2, phospho-p38, Erk1/2, and p-38 were used for primary antibody reaction.

Calvaria Organ Culture and Application of FGF2-Soaked Beads

We followed previously described protocols (Kim et al., 1998). Briefly, calvaria were dissected from E15 mice, and explants without skin were placed on a 0.1-μm pore size Nucleopore filter supported by metal grids. A total of 110 calvarial explants were divided into three groups: 44 for the BSA group, 36 for the FGF2 group, and 30 for the PD/FGF2 group. Heparin-coated acrylic beads (125–150 μm in diameter) were soaked in 25 ng/μl FGF2 or in 0.1% BSA in PBS as control for 30 min at 37°C. The beads were then washed in PBS and implanted with a capillary pipette on the OFs of the sagittal suture. The explants were subsequently cultured at 37°C in a humidified atmosphere of 5% CO2 in air for 48 hr in serum-free DMEM containing penicillin/streptomycin. Ascorbic acid was supplemented daily at 100 μg/ml. The cultured tissues were then fixed overnight with 4% paraformaldehyde (PFA) and used for whole-mount in situ hybridization, or after a series of dehydration steps and paraffin embedding, used for sectional in situ hybridization.

In Situ Hybridization of Whole-Mount Calvaria

In situ hybridization was performed with digoxigenin-UTP–labeled sense and antisense riboprobes of OP as described previously (Kim et al., 1998). Briefly, the explants were treated with proteinase K, fixed again with 4% PFA and 0.2% glutaraldehyde in PBS, and hybridized with the riboprobes overnight at 55°C. The tissue was then washed in high stringency conditions, and the color was developed with the digoxigenin RNA labeling kit. The tissues were subsequently fixed and stored in 50% glycerol before photography.

In Situ Hybridization on Histologic Sections

In situ hybridization was performed by using [35S]UTP-labeled riboprobes as described previously (Park et al., 2001) and a hybridization buffer containing 60% deionized formamide, 300 mM NaCl, 20 mM Tris-HCl, pH 8.0, 5 mM EDTA, 10% dextran sulphate, 0.5 mg/ml yeast tRNA, and 1× Denhardt's solution in diethylpyrocarbonate-treated water. After digestion of deparaffinized sections with 7–10 μg/ml proteinase K to enhance probe penetration, sufficient probe was placed over the slide and the slide was covered with Parafilm. Overnight hybridization was carried out in a humidified sealed box at 52°C, followed by high stringency washes with 50% formamide and 20 mM DTT at 65°C. Slides were then prepared for autoradiography. The dehydrated slides were dipped into photographic emulsion (Kodak NTB-2), dried, and exposed for 1–2 weeks at 4°C. The slides were then developed (Kodak D-19), fixed (Kodak Unifix), counterstained with Delafield's hematoxylin, and mounted with DePeX (BDG).

MTT Assay

Confluent ST2 cells were treated with cycloheximide at various concentrations. After 24 hr, 200 μl of 5 mg/ml MTT was added and the cells were returned to the incubator for an additional 4 hr. The reaction was stopped by removing the medium and adding 200 μl of 0.04 N HCl in isopropanol to the cells. The degree of MTT conversion was measured immediately at 570 nm.

Measurement of Suture Width and Statistical Analysis

Three pictures were taken from each explant under a stereomicroscope (×70, Olympus SZH10 Research Stereo): 0 hr, 24 hr, and 48 hr after bead application. By using pictures of the same magnification, two tangential lines parallel to the midline of the sagittal suture were drawn from the nearest points of the opposing OFs of the parietal bones. The width between the two lines was then measured and statistically analyzed. Data were analyzed with SAS 10.2 (SAS Institute, Inc., Cary, NC). The differences among each group were compared by repeated measures analysis of variance. P < 0.05 was considered significant.

Acknowledgements

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES

We thank Dr. Laura R. McCabe for providing the cDNAs of all the AP1 components and Seong-Hwa Jeong for statistical analysis.

REFERENCES

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  2. Abstract
  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES
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