Introduction
Nonunion is a serious complication of a bone fracture that may occur in any bone of the skeletal system. It occurs when a broken bone fails to heal. Recently, the rapid development of tissue engineering technology has provided an effective method of dealing with this issue. Mesenchymal stem cells (MSCs) are best suited for regenerative medicine due to their extensive proliferation and differentiation potential (1). MSCs are non-hematopoietic multipotent cells which can self-renew and differentiate into cell types of mesodermal tissue, including bone, cartilage, adipose tissue, muscle and tendon (1–3). Previous studies have demonstrated that several signaling pathways are involved in regulating the osteogenic differentiation of MSCs (1,4). Bone morphogenetic proteins (BMPs), belonging to the transforming growth factor-β (TGF-β) superfamily, are known to perform pivotal functions in the areas of embryogenesis, multiple growth and differentiation processes (5–9).
At present, more than twenty BMPs have been identified. Among these, BMP2 and BMP7, have been shown to promote osteogenic differentiation and are currently used as adjunctive therapy to improve bone healing in the clinical setting (10–12). However, it remains unclear as to whether these two factors are in fact the most potent BMPs in promoting osteogenic differentiation and bone formation.
Previous studies have demonstrated that bone morphogenetic protein 9 (BMP9) is more potent in inducing the osteogenic differentiation of MSCs both in vitro and in vivo through a comprehensive analysis of the 14 types of BMPs (6,13). It has been reported that a variety of signaling pathways and cytokines, such as p38 and extracellular signal-regulated protein kinase (ERK)1/2 mitogen-activated protein kinases (MAPKs), JNKs, insulin-like growth factor-2 (IGF-2), fibroblast growth factor-2 (FGF-2) and epidermal growth factor (EGF) are involved in the BMP9-induced osteogenic differentiation of MSCs (14–18). Despite these meaningful discoveries, BMP9 remains the least studied BMP, and the signaling mechanisms through which BMP9 regulates the osteogenic differentiation of MSCs remain unclear and warrant extensive investigation.
Hedgehog (Hh) signaling was initially identified in Drosophila, which includes three members, Sonic hedgehog (Shh), Indian hedgehog (Ihh) and Desert hedgehog (Dhh) signaling (19,20). Hh signaling acts through two transmembrane proteins, the transmembrane receptor, Patched (Ptch) and the seven-pass transmembrane protein Smoothened (Smo); following Hh ligand binding to Ptc1, the suppression of Smo is reversed and this subsequently leads to the activation of the Gli family of transcription factors that mediate the transcription of Hh signaling target genes in cells (19–21). Hh signaling plays a critical role in the regulation of pattern formation, growth, stem cell maintenance and self-renewal in a number of organs during development (21). Several studies have indicated that Hh signaling may act as a key modulator in bone homeostasis (22–25). Furthermore, Hh signaling regulates osteoblast and osteoclast differentiation together with BMP2, BMP4 and BMP7 (24,26,27). Therefore, we spontaneously raised the issue whether Hh signaling is also relevant to the BMP9-induced osteogeneic differentiation of MSCs.
In this study, we sought to investigate the possible involvement and the detailed role of Hh signaling in the BMP9-induced osteogenic differentiation of MSCs. We found that BMP9 exerts an effect on Hh signaling in MSCs and that this leads to alterations in the expression of Hh signaling molecules. The BMP9-induced early and late osteogenic differentiation of MSCs was effectively decreased by the Hh signaling inhibitor, cyclopamine, whereas it was promoted by the Hh signaling agonist, purmorphamine. Furthermore, cyclopamine was shown to inhibit the BMP9-induced transcriptional activity of Smad1/5/8, and to disrupt the BMP9-induced expression of pivotal osteogenic transcription factors. On the contrary, treatment with purmorphamine promoted the BMP9-induced transcriptional activity of Smad1/5/8 and enhanced the BMP9-induced activation of pivotal osteogenic transcription factors. Our data suggest that Hh signaling plays an important regulatory role in the BMP9-induced osteogenic differentiation of MSCs.
Materials and methods
Cell culture and chemicals
C3H10T1/2 and C2C12 cells were obtained from the American Type Culture Collection (Manassas, VA, USA) and maintained in complete Dulbecco’s modified Eagle’s medium (DMEM) with 10% fetal bovine serum (FBS) (both from Gibco, Grand Island, NY, USA), 100 U/ml of penicillin and 100 μg/ml of streptomycin at 37°C in 5% CO2.
Anti-phospho-Smad1/5/8 (#9511), anti-ERK1/2 (#4695), anti-phospho-ERK1/2 (#4370), anti-p38 (#9212) and anti-phospho-p38 (#4511) antibodies were purchased from Cell Signaling Technology (Danvers, MA, USA). Anti-Smad1/5/8 (sc-6031), anti-osteopontin (OPN; sc-21742), anti-osteocalcin (OCN; sc-23790), anti-Runx2 (sc-12488), anti-distal-less homeobox 5 (Dlx5; sc-18151) and anti-β-actin (sc-47778) antibodies were obtained from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA, USA). The Hh signaling inhibitor, cyclopamine, was obtained from Selleckchem (Houston, TX, USA) and the Hh signaling agonist, purmorphamine, was obtained from Cayman Chemical Co. (Ann Arbor, MI, USA). These reagents were dissolved in dimethyl sulfoxide (DMSO) and stored at −80°C. The recombinant adenoviruses, Ad-BMP9 and Ad-GFP, were kindly provided by Dr Tong-Chuan He (University of Chicago Medical Center, Chicago, IL, USA). Other chemicals were purchased from Sigma-Aldrich (St. Louis, MO, USA) or Fisher Scientific (Pittsburgh, PA, USA).
Isolation of mouse embryonic fibroblasts (MEFs)
MEFs were isolated from mice on postcoital day 13.5, as previously described (14). Each embryo, voided of its internal organs, was dissected into 10 ml of sterile pbosphate-buffered saline (PBS), and sheared through an 18-gauge syringe in the presence of 1 ml of 0.25% trypsin and 1 mM ethylenediaminetetraacetic acid (EDTA). After 15 min of incubation with gentle shaking at 37°C, 10 ml DMEM with 10% FBS were added to inactivate trypsin. The cells were plated in 100 mm dishes and incubated for 24 h at 37°C. The adherent cells were used as MEFs. Aliquots were kept in a liquid nitrogen tank. All MEFs used in this study were at passage 5.
RNA isolation and semi-quantitative reverse transcription-polymerase chain reaction (RT-PCR)
Total RNA was isolated using TRIzol reagent and used to generate the cDNA templates by reverse transcription (RT) reaction with hexamer and Superscript II RT (both from Invitrogen, Carlsbad, CA, USA). Semi-quantitative RT-PCR was carried out as described previously (14). The first strand cDNA products were further diluted 5- to 10-fold and used as templates for PCR. All samples were normalized with the expression level of mouse glyceraldehyde 3-phosphate dehydrogenase (GAPDH). PCR primers were designed using the Primer3 program (Free Software Foundation, Inc., Boston, MA, USA) to amplify the genes of interest as follows: Smo forward, 5′-TTG TGC TCA TCA CCT TCA GC-3′ and reverse, 5′-TGC CAA ACA TGG CAA ATA GA-3′; Hedgehog-interacting protein (Hhip) forward, 5′-CCT GTC GAG GCT ACT TTT CG-3′ and reverse, 5′-GGG CAG GTT GAA CTG TGA CT-3′; Ptch1 forward, 5′-CTC AGG CAA TAC GAA GCA CA-3′ and reverse, 5′-GAC AAG GAG CCA GAG TCC AG-3′; Gli family zinc finger 1 (Gli1) forward, 5′-GAA GGA ATT CGT GTG CCA TT-3′ and reverse, 5′-GCA ACC TTC TTG CTC ACA CA-3′; Gli family zinc finger 2 (Gli2) forward, 5′-ACC ATG CCT ACC CAA CTC AG-3′ and reverse, 5′-CTG CTC CTG TGT CAG TCC AA-3′; Ihh forward, 5′-CGT GCA TTG CTC TGT CAA GT-3′ and reverse, 5′-CTC GAT GAC CTG GAA AGC TC-3′; Dhh forward, 5′-CTT GGA CAT CAC CAC GTC TG-3′ and reverse, 5′-GTA GTT CCC TCA GCC CCT TC-3′; Shh forward, 5′-CTG GCC AGA TGT TTT CTG GT-3′ and reverse, 5′-GAT GTC GGG GTT GTA ATT GG-3′; OPN forward, 5′-ACA CTT TCA CTC CAA TCG TCC-3′ and reverse, 5′-TGC CCT TTC CGT TGT TGT CC-3′; OCN forward, 5′-TCT GAC AAA GCC TTC ATG TCC-3′ and reverse, 5′-AAA TAG TGA TAC CGT AGA TGC G-3′; Runx2 forward, 5′-CCG GTC TCC TTC CAG GAT-3′ and reverse, 5′-GGG AAC TGC TGT GGC TTC-3′; Dlx5 forward, 5′-CTC AGC CAC CAC CCT CAT-3′ and reverse, 5′-TGG CAG GTG GGA ATT GAT-3′; inhibitor of DNA binding 1 (Id1) forward, 5′-ACG ACA TGA ACG GCT GCT-3′ and reverse, 5′-CAG CTG CAG GTC CCT GAT-3′; inhibitor of DNA binding 2 (Id2) forward, 5′-CAG CAT CCC CCA GAA CAA-3′ and reverse, 5′-TCT GGT GAT GCA GGC TGA-3′; inhibitor of DNA binding 3 (Id3) forward, 5′-CTA CGA GGC GGT GTG CTG-3′ and reverse, 5′-GCG CGA GTA GCA GTG GTT-3′; GAPDH forward, 5′-GGC TGC CCA GAA CAT CAT-3′ and reverse, 5′-CGG ACA CAT TGG GGG TAG-3′. A touchdown cycling program was carried out as follows: 94°C for 5 min for 1 cycle, 94°C for 30 sec, 68°C for 30 sec, and 72°C for 12 cycles with a decrease in 1°C/cycle and then at 94°C for 30 sec, 55°C for 30 sec, and 72°C for 30 sec for 18–27 cycles depending on the abundance of a given gene. The specificity of PCR products was confirmed by resolving the PCR products on 2% agarose gels.
Alkaline phosphatase (ALP) activity assay
ALP activity was assessed by a modified Great Escape SEAP Chemiluminescence Assay (BD Clontech, Mountain View, CA, USA) and/or histochemical staining assay (using a mixture of 0.1 mg/ml of napthol AS-MX phosphate and 0.6 mg/ml of Fast Blue BB salt) as described previously (14,28). The C3H10T1/2 cells, MEFs and C2C12 cells were infected with Ad-GFP or Ad-BMP9 and/or treated with various concentrations of cyclopamine or purmorphamine. For the chemilluminescence assays, each assay condition was performed in triplicate, and the results were repeated in at least 3 independent experiments. ALP activity was normalized to the total cellular protein level. For ALP histochemical staining, the induction of ALP expression was detected at different time points following treatment using histochemical staining assays, and then recorded using bright-field microscopy.
Alizarin Red S staining
The C3H10T1/2 cells, MEFs and C2C12 cells were seeded in 24-well culture plates and infected with Ad-GFP or Ad-BMP9 and/or treated with cyclopamine or purmorphamine. These cells were cultured in the presence of ascorbic acid (50 mg/ml) and β-glycerophosphate (10 mM). At 14 days after treatment, mineralized matrix nodules were stained for calcium precipitation by means of Alizarin Red S staining assay, as previously described (28,29). The cells were fixed with 0.05% (v/v) glutaraldehyde at room temperature for 10 min. After being washed with distilled water, the fixed cells were incubated with 0.4% Alizarin Red S for 5 min, followed by extensive washing with distilled water. The results were repeated in at least 3 independent experiments. The staining of calcium mineral deposits was recorded under brightfield microscopy.
Western blot analysis
Western blot analysis was performed as described previously (15). The cells were plated in a 100 cm2 cell culture dish and treated as scheduled. At the indicated time points, the cells were collected and lysed in Laemmli buffer. Cleared total cell lysate was denatured by boiling and loaded onto a 8–15% gradient sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE). Following electrophoretic separation, the proteins were transferred onto an Immobilon-P membrane. The membrane was blocked with SuperBlock blocking buffer for 2 h at 37°C and probed with the primary antibody (diluted 1:1,000) overnight at 4°C, followed by incubation with a secondary antibody-conjugated to horseradish peroxidase (diluted 1:5,000; Zhongshan Golden Bridge Biotechnology, Beijing, China). Finally, the images of the target bands were developed with SuperSignal West Pico Chemiluminescent Substrate (Thermo Scientific, Rockford, IL, USA). Each assay was carried out in triplicate.
Transfection and luciferase reporter assay
The C3H10T1/2 cells were seeded in 25 cm2 cell culture flasks and transfected with 3 μg per flask of BMP receptor Smad-binding element luciferase reporter (p12xSBE-Luc) using Lipofectamine 2000 (Invitrogen). At 24 h after transfection, the cells were replated in 24-well plates and treated with Ad-BMP9 and/or cyclopamine or purmorphamine. At 24 and 48 h after treatment, the cells were lysed and collected for luciferase assays using the Luciferase assay kit (Promega, Madison, WI, USA) as previously described (14,29). Each assay condition was performed in triplicate. The results were repeated in at least 3 independent experiments. Luciferase activity was normalized by total cellular protein concentrations among the samples.
Statistical analysis
For all quantitative assays, each assay condition was performed in triplicate, and the results were repeated in at least 3 independent experiments. Data are expressed as the means ± SD. Statistical analysis was performed using SPSS software version 14 (SPSS Inc., Chicago, IL, USA). P<0.05 was considered to indicate a statistically significant difference. All data collected were subjected to statistical analysis.
Results
BMP9 affects Hh signaling
First of all, we sought to determine whether BMP9 exerts an effect on Hh signaling in MSCs. The C3H10T1/2 cells were infected with Ad-BMP9 or Ad-GFP, and the expression of pivotal Hh signaling molecules, including Smo, Hhip, Ptch1, Gli1, Gli2, Dhh, Ihh and Shh was assessed by semi-quantitative RT-PCR. We found that BMP9 exerted an effect on Hh signaling, leading to an altered expression pattern of pivotal Hh signaling molecules (Fig. 1). Similar results were obtained with the MEFs and C2C12 cells (data not shown). These results suggest that BMP9 affects Hh signaling in the MSCs at least partially by altering the expression of related molecules.
Effect of Hh signaling on the BMP9-induced early osteogenic differentiation of MSCs
To further determine the detailed role of Hh signaling in the BMP9-induced osteogenic differentiation of MSCs, we used cyclopamine (Cy) to block the activity of Hh signaling (30,31), and purmorphamine (Pur) to activate Hh signaling (23,32). The C3H10T1/2 cells were transfected with Ad-BMP9 or Ad-GFP in the presence of various concentrations of cyclopamine (6, 10, 14 and 18 μM) or purmorphamine (0.4, 0.6, 0.8 and 1 μM). Of note, cyclopamine significantly inhibited the BMP9-induced ALP activity in a dose-dependent manner (Fig. 2A and C). Conversely, treatment with purmorphamine markedly enhanced the BMP9-induced ALP activity (Fig. 2B and C). Similar phenomena were observed with the MEFs (Fig. 3A and B) and C2C12 cells (Fig. 3C and D). The above results suggest that the BMP9-induced early osteogenic differentiation of MSCs is markedly blocked by the inhibition of Hh signaling, whereas it is enhanced by the activation of Hh signaling.
Effect of Hh signaling on the BMP9-induced late osteogenic differentiation of MSCs
Although ALP is a well-established early marker, it is hardly an accurate predictor of the late stage of osteogenic differentiation (15,29). We further determined the effect of Hh signaling on BMP9-induced late stage of osteogenic differentiation. Fistly, C3H10T1/2 cells, MEFs and C2C12 cells were infected with Ad-BMP9 in the presence of cyclopamine (14 μM) and purmorphamine (0.8 μM) respectively. At 14 days post treatment, Alizarin Red S staining was conducted to determine the effects of Hh signaling on BMP9-induced matrix mineralization. We found that the BMP9-induced matrix mineralization was decreased by cyclopamine, whereas it was increased by purmorphamine (Fig. 4). Subsequently, the effect of Hh signaling on the BMP9-induced expression of the late osteogenic markers, osteopotin (OPN) and osteocalcin (OCN), were assessed by semi-quantitative RT-PCR and western blot analysis in the C3H10T1/2 cells and MEFs. We found that treatment with cyclopamine resulted in a significant decrease in the BMP9-induced expression of OPN and OCN; however, treatment with purmorphamine led to a marked increase in the BMP9-indcued OPN and OCN expression at the gene and protein levels (Fig. 5). Taken together, the above results suggest that Hh signaling plays a pivotal role in regulating both the early and late stages of the BMP9-induced osteogenic differentiation of MSCs.
Effect of Hh signaling on the BMP9-induced activation of the Smad1/5/8 and MAPK pathways
We then sought to explore the possible mechanisms behind the effects of Hh signaling on the BMP9-induced osteogenic differentiation of MSCs. Previous studies have reported that Smad-dependent Smad1/5/8 canonical signaling and Smad-independent MAPKs pathways are important in regulating BMP9 osteoinductive signaling (16,33). Therefore, we wished to determine whether the BMP9-induced activation of the Smad1/5/8 and MAPK signaling pathways is also affected by Hh signaling. Firstly, using the BMP responsive Smad1/5/8 reporter, p12xSBE-Luc (14,15), we found that the BMP9-induced transcriptional activity of Smad1/5/8 was augmented by purmorphamine, and it was inhibited by cyclopamine (Fig. 6A). However, we found that the BMP9-induced phosphorylation of Smad1/5/8, ERK1/2 and p38 was not altered by treatment with cyclopamine or purmorphamine (Fig. 6B and C). Collectively, these above-mentioned results suggest that Hh signaling regulates the BMP9-induced osteogenic differentiation of MSCs at least partially by affecting the transcriptional activity of canonical Smad1/5/8 directly rather than altering the phosphorylation status of Smad1/5/8.
Effect of Hh signaling on the BMP9-induced expression of pivotal osteogenic transcription factors
It has been demonstrated in previous studies that pivotal osteogenic transcription factors, such as Id1, Id2, Id3, Runx2 and Dlx5, are targets of BMP9 and are critical to the osteogenic differentiation of MSCs (6,34). In order to determine the effects of Hh signaling on the BMP9-induced expression of pivotal osteogenic transcription factors, the C3H10T1/2 cells were infected with Ad-BMP9 in the presence of cyclopamine (14 μM) or purmorphamine (0.8 μM). Using semi-quantitative RT-PCR, we found that the BMP9-induced gene expression of Id1, Id2, Id3, Runx2 and Dlx5 was markedly increased by purmorphamine, while it was inhibited by cyclopamine (Fig. 7A). Subsequently, we further examined the protein expression levels of Dlx5 and Runx2 by western blot analysis. Consistently, the BMP9-induced expression of Dlx5 and Runx2 was decreased by treatment with cyclopamine, while it was enhanced by treatment with purmorphamine (Fig. 7B). These results suggest that Hh signaling regulates the BMP9-induced expression of pivotal osteogenic transcription factors. Taken together, these data indicate that Hh signaling is involved in regulating the BMP9-induced osteogenic differentiation of MSCs.
Discussion
BMPs are potent growth factors which are important for cell differentiation and proliferation and over 20 BMPs have been identified to data; among these, BMP2, BMP4, BMP6 and BMP7 have been validated to commit MSCs to the osteoblast lineage (9–11,13,35,36). BMP9 [also known as growth differentiation factor 2 (GDF-2)] was originally isolated from fetal mouse liver cDNA libraries and is a potent stimulant of hepatocyte proliferation (37). BMP9 induces the cholinergic phenotype of embryonic basal forebrain cholinergic neurons, maintains the homeostasis of iron metabolism and regulates glucose and lipid metabolism in the liver (38,39). Previous studies have validated that BMP9 is a potent factor which induces the osteogenic differentiation of MSCs (6,13). It has been demonstrated that TGF-β type I receptors activin receptor-like kinase (ALK)1 and ALK2, as well as TGF-β type ΙΙ receptors BMP receptor type II (BMPRΙΙ) and ActRΙΙ are essential for the BMP9-induced osteogenic differentiation of MSCs (28,29). Furthermore, a distinct set of targets that may play important roles in the BMP9-induced osteogenic differentiation of MSCs have been identified (6,34). Various signaling pathways with diverse functions have been found to play roles in BMP9-induced osteogenesis (14–18). Nevertheless, BMP9 remains to be the least studied BMPs, and little is known about the specific molecular mechanisms underlying BMP9-induced osteogenic differentiation.
In this study, we analyzed the detailed roles of Hh signaling in the BMP9-induced osteogenic differentiation of MSCs, and the possible mechanisms involved. We found that BMP9 affected Hh signaling at least partly by affecting the expression of the related molecules, Smo, Hhip, Ptch1, Gli1, Gli2, Dhh, Ihh and Shh, in the MSCs. Furthermore, we found that the BMP9-induced activity of the early osteogenic marker ALP and the expression of late osteogenic markers, such as OPN and OCN, as well as the matrix mineralization in MSCs were reduced by the Hh signaling inhibitor, cyclopamine, while they were promoted by the activator of Hh signaling, purmorphamine. Mechanistically, we found that the BMP9-induced transcriptional activity of Smad1/5/8 and the expression of pivotal osteogenic transcription factors were enhanced by purmorphamine, while they were inhibited by cyclopamine. These results suggest that Hh signaling plays an important role in the BMP9-induced osteogenic differentiation of MSCs.
Several biological studies have indicated that Hh signaling, most notably Shh signaling and Ihh signaling, plays an important role in osteogensis and bone development (22–25). In a previous study, C3H10T1/2 cells transfected with a plasmid encoding N-terminal Shh showed an increased expression of ALP and OCN (40). The complete knockout of Shh (Shh−/−) results in mice lacking vertebrae and having major defects in the distal bones of the limbs (41). Hh signaling in mature osteoblasts regulates both bone formation and resorption by upregulating the osteoblast expression of parathyroid hormone-related protein (PTHrP), which promotes RANKL expression through PKA and its target transcription factor, CREB (24). The loss of Ihh signaling by genetic knockout (Ihh−/−) has been shown to result in decreased secondary palate ossification (42). Furthermore, interaction between Hh and BMP signaling has been found to regulate osteogenic differentiation and bone formation. For example, Shh signaling induces osteoblast differentiation by interacting with BMP2 (26,40). Ihh and the BMP2 gene synergistically increase the osteogenic potential of human MSCs (43). Shh signaling, acting as a negative effector of BMP signaling, suppresses osteo/dentinogenic differentiation in stem cells from apical papilla (44). Although these studies on the precise role of Hh signaling in the skeletal system did not lead to complete unanimity, it is well accepted that Hh signaling plays a functional role in bone development and bone metabolism. In the present study, we found that Hh signaling is involved and may exert synergistic effects on the BMP9-induced osteogenic differentiation of MSCs.
BMPs transduce the signaling activity by binding to BMP receptors, and subsequently activating BMP receptor kinases. These activated receptors phosphorylate the transcription factors, Smad1/5/8, which in turn form a heterodimeric complex with Smad4 and regulate downstream target genes in concert with co-activators (5). It has been previously reported that canonical Smad1/5/8 signaling is involved in the BMP9-induced osteogenic differentiation of MSCs (33). In the present study, we found that Hh signaling increased the BMP9-induced transcriptional activity of Smad1/5/8 without altering the phosphorylation status of Smad1/5/8. We thus hypothesized that this phenomenon may be due to the following reasons: i) Hh signaling may promote the translocation of phosphorylated Smad1/5/8, and may thus result in the acceleration of these proteins in the nucleus; ii) the target transcription factors of Hh signaling, such as Gli1 may act as co-activators which interact with Smad1/5/8 to enhance the transcriptional activity of Smad1/5/8. However, more intensive methods, such as co-immunoprecipitation (Co-IP) and western blot analysis of nuclear proteins should be conducted to verify these above-mentioned hypotheses. It has been proven that p38 and ERK1/2 MAPKs signaling are also involved in the BMP9-induced osteogenic differentiation of MSCs and bone formation (16). In this study, however, we found that Hh signaling had no effect on the phosphorylation of p38 and ERK1/2. Nevertheless, the potential role of p38 and ERK1/2 in regulating the effects of Hh signaling on the BMP9-induced osteogenic differentiation of MSCs cannot be eliminated rashly without careful consideration and validation.
It has been previously demonstrated that Id1, Id2, Id3, Dlx5 and Runx2 are critical to the BMP9-induced osteogenic differentiation of MSCs (6,34). Studies have indicated that Hh signaling affects the expression of Runx2 in the process of osteogenic differentiation (45,46). In this study, we found that the activation of Hh signaling promoted the expression of these pivotal osteogenic transcription factors induced by BMP9, while the inhibition of Hh signaling had the opposite effect. These results suggest that Hh signaling is involved in the BMP9-induced osteogenic differentiation of MSCs possibly by exerting effects on BMP9 downstream pivotal osteogenic transcription factors directly. Various experiments, such as western blot analysis and chromatin immunoprecipitation (ChIP) assay should be conducted to further explore the role of these transcription factors in this process. Moreover, other signaling molecules which may also be involved need to be intensively characterized and illustrated.
In conclusion, the findings of the present study indicate that BMP9 affects Hh signaling in MSCs at least partly through altering the expression of related molecules. The inhibition of Hh signaling decreased the BMP9-induced early and late osteogenic differentiation of MSCs, while the activation of Hh signaling promoted this osteogenic differentiation. Mechanistically, we found that Hh signaling exerts regulatory effect on the BMP9-induced osteogenic differentiation of MSCs partly through the modulation of the transcriptional activity of Smad1/5/8 and the expression of pivotal osteogenic transcription factors. These findings may contribute not only to promote the development of BMP9-mediatied bone tissue engineering, but also to provide a rational basis for its clinical application.