Htra1 is a Novel Transcriptional Target of RUNX2 That Promotes Osteogenic Differentiation
Paul P.R. Iyyanara,b Merlin P. Thangaraja B. Frank Eamesc Adil J. Nazaralia
aLaboratory of Molecular Cell Biology, College of Pharmacy and Nutrition and Neuroscience Research Cluster, University of Saskatchewan, Saskatoon, SK, Canada, bDivision of Developmental Biology, Cincinnati Children’s Hospital Medical Center, Cincinnati, OH, USA, cDepartment of Anatomy and Cell Biology, University of Saskatchewan, Saskatoon, SK,Canada
Key Words
Runx2 • Htra1 • Osteoblast differentiation • Matrix mineralization
Abstract
Background/Aims: Runt-related transcription factor 2 (Runx2) is a master regulator of osteogenic differentiation, but most of the direct downstream targets of RUNX2 during osteogenesis are unknown. Likewise, High-temperature requirement factor A1 (HTRA1) is a serine protease expressed in bone, yet the role of Htra1 during osteoblast differentiation remains elusive. We investigated the role of Htra1 in osteogenic differentiation and the transcriptional regulation of Htra1 by RUNX2 in primary mouse mesenchymal progenitor cells. Methods: Overexpression of Htra1 was carried out in primary mouse mesenchymal progenitor cells to evaluate the extent of osteoblast differentiation. Streptavidin agarose pulldown assay, chromatin immunoprecipitation assay, and dual luciferase assay were carried out to investigate the interaction of RUNX2 protein at the Htra1 promoter during osteoblast differentiation. Results: Overexpression of Htra1 increased the production of mineralized bone matrix, upregulating several osteoblast genes, such as Sp7 transcription factor (Sp7) and Alkaline phosphatase, liver/bone/kidney (Alpl). In addition, Htra1 upregulated osteogenesis-related signalling genes, such as Fibroblast growth factor 9 (Fgf9) and Vascular endothelial growth factor A (Vegfa). A series of experiments confirmed Htra1 as a direct RUNX2 transcriptional target. Overexpression of Runx2 resulted in the upregulation of Htra1 mRNA and protein. Chromatin immunoprecipitation and streptavidin agarose pull-down assays showed that RUNX2 binds a proximal -400 bp region of the Htra1 promoter during osteogenic differentiation. Dual luciferase assays confirmed that RUNX2 activates the proximal Htra1 promoter during osteogenic differentiation. Mutation of putative RUNX2 binding sites revealed that RUNX2 interacts with the Htra1 promoter at -252 bp and -84 bp to induce Htra1 expression. Conclusion: We demonstrate that Htra1 is a positive regulator of osteogenic differentiation, showing for the first time that Htra1 is a direct downstream target of RUNX2.
Introduction
RUNX2 is a master regulatory transcription factor critical for osteogenic differentiation and bone formation [1]. Runx2 null mutant mice exhibit neonatal lethality with complete lack of ossification [1], and RUNX2 haploinsufficiency in humans results in cleidocranial dysplasia, a skeletal disorder characterized by bone and dental abnormalities [2–4]. RUNX2 regulates several osteogenic genes, such as Integrin binding sialoprotein (Ibsp) [5], Matrix metalloprotease-13 (Mmp13) [6, 7], and Sp7 [8, 9], by binding a consensus site RCCRC(A/T) [10] in their proximal promoter regions. However, a likely majority of the transcriptional targets of RUNX2 during osteogenic differentiation remain to be characterized [11].
HTRA1 is one of four members of the HTRA family of serine proteases. HTRA1 has been implicated in several diseases, including age-related macular degeneration [12], cerebral autosomal recessive arteriopathy with subcortical infarcts and leukoencephalopathy (CARASIL) [13–15], and cancer [16, 17]. HTRA1 is expressed in hypertrophic chondrocytes and osteoblasts under both physiological [18] and pathological conditions, including osteoarthritis [19] and the fracture callus [20]. Despite Htra1 expression in bone, published studies conflict in concluding whether Htra1 is a positive or negative regulator of osteogenesis. Some studies have demonstrated that Htra1 is an inducer of osteogenic differentiation, where it is involved in the lineage commitment of mesenchymal stem cells to form osteoblasts [20] at the expense of adipocytes [21]. Silencing Htra1 abolishes osteogenic differentiation induced by all-trans retinoic acid in adipose-derived stromal cells [22]. In contrast, other studies have found Htra1 to be a negative regulator of osteogenesis in 2T3 osteoblasts [23] and the KusaO cell line [24]. Therefore, the role of Htra1 in osteogenesis remains unclear.
Very little information is available on the gene networks and signalling pathways downstream of Htra1 in osteogenic differentiation. HTRA1 has proteolytic activity, degrading various substrates in the extracellular matrix (ECM), such as Fibronectin, Fibulin, and Nidogen [15, 25–27]. In Xenopus embryos, Htra1 shares similar expression domains with FGF ligands, such as FGF8 and FGF4, and Htra1 promotes long-range FGF signalling via cleavage of proteoglycans, such as Biglycan, Syndecan-4, and Glypican-4 [28]. Such an impact of Htra1 on FGF signalling could influence bone formation. Fgf9 induces Vegfa expression for skeletal vascularization of the proximal limb during development [29]. Furthermore, FGF9 promotes Vegfa-mediated Mmp9 expression for the bone remodelling process in the long bones [30]. FGF8 also induces Htra1 in developing chick facial mesenchyme [31], suggesting reciprocal feedback. Htra1 promotes ERK/MAPK-dependent MMP expression during adipogenic differentiation, potentially via breakdown of ECM-related components such as Fibronectin [21]. In human bone marrow stem cells, HTRA1 suppresses the expression of Sclerostin (Sost), an inhibitor of WNT signalling-dependent osteogenesis [20]. Further insights into the molecular signalling downstream of Htra1 is necessary to decipher its role in osteogenesis.
Mesenchymal progenitor cells from the embryonic palate differentiate directly into osteoblasts to form the palatal bones through the process of intramembranous ossification [32–34]. In the present study, we have investigated the impact of Htra1 overexpression on the expression of osteogenic markers and matrix mineralization during osteogenic differentiation of primary palatal mesenchymal progenitor cells in vitro. Furthermore, we examined the transcriptional regulation of Htra1 by RUNX2 during osteogenic differentiation. Data reveal that Htra1 is a positive modulator of osteogenic differentiation, regulating the expression of several key osteogenic genes. In addition, our data demonstrate for the first time that RUNX2 binding to the proximal Htra1 promoter is necessary and sufficient to induce Htra1 expression, thus elucidating Htra1 as a novel direct downstream target of RUNX2 that promotes osteogenesis.
Materials and Methods
Immunohistochemistry
Immunohistochemistry was carried out as previously described [34]. Briefly, mouse embryonic heads of stage E16.5 were fixed with freshly prepared 4% paraformaldehyde overnight and rehydrated in 30% sucrose at 4°C. Frozen coronal sections (10 μm) were rehydrated with PBS for 45 min, permeabilized with 0.1% Triton X-100 and blocked with 3% skim milk containing 0.1% Triton X-100 in 1X PBS for 1 h at room temperature. Sections were then incubated overnight with the following primary antibodies: HTRA1 (1:200; Abcam ab38611), RUNX2 (1:200; Abcam ab23981) or SP7 (1:800; Abcam ab22552) in 1X PBS with 0.1% Triton X-100 at 4ο C. Sections were washed three times and treated with secondary antibodies conjugated with Alexa Fluor® 594 (1:400; Invitrogen) in 1X PBS with 0.1% Triton X-100 at room temperature for 1.5 h. Stainings with rabbit IgG served as a negative control to assess specific staining of primary antibodies employed in the study. Sections were then washed three times, five mins each and mounted in Prolong gold antifade reagent (Invitrogen) for imaging.
Primary mesenchymal progenitor cells derived from mouse embryonic palate
Primary mesenchymal progenitor cells were isolated from the palatal shelves of wild-type C57BL/6J mouse embryos at embryonic day (E)13.5 as previously described [34, 35]. Briefly, palatal shelves were micro-dissected from the maxilla of E13.5 embryos and incubated with 0.25% trypsin (Sigma) for 15 min at 37οC [36]. Cells were gently pipetted up and down, passed through 70-µm-cell strainer (BD Falcon) and plated on poly-D-lysine-coated T25 flasks. Cells were cultured in Dulbecco’s Modified Eagle’s Medium (DMEM): Ham’s F12 (DMEM/F12) (1:1) media containing 10% FBS, 1% antibiotic-antimycotic solution (Sigma) and passaged no more than three times. Osteogenic differentiation was induced as described previously [34, 37] with minor modifications. Briefly, mesenchymal progenitor cells were seeded on PDL-coated 24-well plates at a cell density of 5 X 104 cells per well in triplicate and cultured until they reach confluence. Osteogenic differentiation was induced with osteogenic differentiation medium comprising of DMEM with 4, 500 mg/L glucose, 10% FBS, 2mM L-glutamine, 1% antibiotic-antimycotic solution, 50µg/ml L-ascorbic acid 2-phosphate sesquimagnesium salt (Sigma), 10 mM β-glycerophosphate (Sigma) and 100 nM dexamethasone (Sigma). Media was changed every third day for up to 8, 14 or 21 days. Cells were harvested for RNA isolation or fixed for alkaline phosphatase (ALP) staining at day 8 and 15.
Alkaline phosphatase (ALP) staining
ALP staining was carried out as previously described [34, 38]. Primary mesenchymal progenitor cells were washed once with PBS (pH 7.4), followed by fixation in 4% paraformaldehyde for 15 min and then washed twice with PBS. Cells were then treated with ALP buffer (100 mM NaCl, 100 mM Tris–HCl pH 9.5, 50 mM MgCl2, 0.1% Tween-20) for 10 min prior to staining with 4.5 μl/ml of 5-Bromo-4-chloro-3-indolyl phosphate and 3.5 μl/ml of nitro blue tetrazolium in ALP buffer for 10 min. The reaction was stopped with PBS containing 20 mM EDTA buffer. Images were taken using a bright field microscope and the percentage area of ALP-positive blue staining was quantified using ImageJ (NIH) [39].
Alizarin red S (ARS) staining and quantitation
ARS staining and quantitation was carried out as described [34, 40] with minor modifications. Briefly, primary mesenchymal progenitor cells were washed once with PBS, pH 7.4, followed by fixation in 4% paraformaldehyde for 15 min and then washed two times with PBS. Cells were subsequently washed with deionized water to remove excess PBS, prior to the addition of 250 µl of 40 mM ARS (Sigma) solution (pH 4.1). Cells were incubated at room temperature for 20 min with gentle shaking. Excess dye was aspirated, and cells were washed with deionized water and imaged using bright field microscopy. For quantitation of ARS, 400 µl of 10% acetic acid (Fisher Scientific) was added to each well and incubated for 30 min with shaking. The cells were then dislodged using a cell scraper (Fisher Scientific) and transferred to a microcentrifuge tube containing 10% acetic acid. After vortexing for 30 s, samples were heated at 85οC for 10 min and cooled on ice for 5 min. The slurry was centrifuged at 15, 000 g for 15 min and 300 µl of the supernatant transferred to a new micro-centrifuge tube followed by the addition of 150 µl of 10% solution of ammonium hydroxide (Sigma). After vortexing, 150 µl was transferred from this mixture to an opaque walled 96-well plate for measurement of absorbance at 405 nm. A standard plot of ARS concentration was constructed by serially diluting 40 mM ARS in a buffer containing 10% (v/v) acetic acid and 10% (v/v) ammonium hydroxide. Absorbance values of standard concentrations were used to interpolate concentrations of the test sample.
Quantitative real-time polymerase chain reaction (qPCR)
Total RNA was isolated from the mesenchymal progenitor cells using RNA mini spin column as per the manufacturer’s protocol (BioRad). First strand cDNA synthesis (Reverse transcription) was performed in 20 µl reactions with 500 ng of total RNA using High-Capacity cDNA Reverse Transcription Kit (Invitrogen). qPCR was carried out using SYBR green master mix (Invitrogen) in 7300-real-time PCR system (Applied Biosystems). The primer pairs used for qPCR are listed in Table 1.
Htra1 overexpression transfections
Htra1 vector (Origene) was digested using SalI and XhoI restriction enzymes to separate the Htra1 cDNA, and the empty vector (pCMV6-myc-ddk) was generated by self-ligation of SalI and XhoI compatible sticky ends. Site-directed mutagenesis of Htra1 at nucleotides corresponding to serine 328 to alanine; Htra1 (mut) [41] was carried out by PCR using QuikChange Lightning kit (Agilent Technologies) with the following primers: 5′CATCAATTATGGAAATGCCGGAGGCCCGTTAG 3ʹ and 5′GTAGTTAATACCTTTACGGCCTCCGGGCAATC 3ʹ. Primary mesenchymal progenitor cells were transfected with polyethylenimine (PEI) (Linear Mol. Wt 25, 000, Polysciences, Inc.). For each well of a 24-well plate, 700 ng of Htra1, Htra1 (mut) or empty vector were transfected with 1.5 µl of PEI (1 mg/ml), every third day in antibiotic-free media in the presence of serum. Cells were either fixed for ALP or ARS staining or lysed for RNA isolation at indicated time points. Similarly, the Runx2 vector (Origene) was transfected as described above for the overexpression of Runx2 in primary mesenchymal progenitor cells.
Osteogenic array
Osteogenesis and bone remodelling real-time PCR array (Bio-Rad), which contains a set of 84 osteogenic pathway related genes in 96-well format, was used to determine differentially regulated genes after overexpression of Htra1. Primary mesenchymal progenitor cells were transfected as described above, with Htra1 or empty vector and allowed to differentiate for seven days. 1000 ng of RNA was converted to cDNA in 20 µl reactions. The cDNA sample was diluted with 100 µl of nuclease-free water and 1 µl of cDNA was used to load each well of the 96-well plate. Gapdh was used as an endogenous control and relative gene expression was determined using the ΔΔCt method. Three biological replicates each for samples treated with Htra1 overexpression vector or empty vector were run individually and analyzed together to get the relative gene expression data. Gene expression in Htra1-overexpressed samples greater than 1.5-fold was considered upregulated and lesser than 0.6-fold was considered as downregulated, relative to the expression in the empty vector-treated samples.
Biotinylated probes
Biotinylated probes were
synthesized by PCR amplification from mouse genomic DNA using 5' biotin-labeled
primers (Invitrogen). The primer pairs used for the synthesis of 1 kb Htra1
promoter probe (-994 to +72 bp) were forward 5'-biotin-TCCTCCTTGAGTCAGGGTCA-3' and reverse 5'-biotin-GCAGCAGTAGCAAAGACAGG-3'. The primers
for the -400 bp probe were forward primer 5'-biotin-AAGTTCACAGCCACAGTCCC-3' and reverse
primer 5'-biotin-TAGCAAAGACAGGAGCGTGG-3'. The annealing
temperature for the primers was 60οC. All PCR
amplified biotinylated probes were electrophoresed on an agarose gel and
purified using a gel extraction kit (Thermo Fisher).
Streptavidin agarose pull-down assay (SAPA)
SAPA was carried out as previously described [42, 43] with minor modifications. Primary mesenchymal cells were harvested 5 days after differentiation and lysed with RIPA buffer containing 50 mM Tris–HCl, pH 7.4, 1% NP-40, 0.25% sodium deoxycholate,150 mM NaCl, 1 mM NaF, 1 mM Na3VO4, 1 mM EDTA, and 1% protease inhibitor cocktail (Sigma). Lysates were vortexed, passed through a 28-gauge needle, incubated on ice for a minimum of 30 min, and centrifuged. Protein concentration in the supernatant was measured using the Bradford assay. SAPA was carried out using 5 µg biotinylated DNA probe added to the cell lysates (250 µg) and incubated at 4οC overnight with rotational mixing. Streptavidin-agarose beads (20 µl) were added to each sample and incubated for 4 h at 4οC with rotation. The beads were then washed four times with 1 ml RIPA buffer and centrifuged at 4οC. Beads were then suspended in 30 µl of 2X SDS loading buffer, boiled for 15 min and centrifuged. The resulting supernatant was used for western blot analysis.
Western blot analyses
Western bot analyses were carried out as previously described [34]. Briefly, proteins separated on a 12% sodium dodecyl sulfate (SDS)-polyacrylamide gel and blotted to PVDF membranes (Bio-Rad) for 2 hours at 4οC. Blots were blocked with 4% skimmed milk for 1 hour and incubated with primary antibody anti-RUNX2 rabbit polyclonal (1:500; Cell signaling D1H7), anti-HTRA1 rabbit polyclonal (1:1000; Abcam 38611) and β-actin mouse monoclonal (1:3000; Cell Signaling) overnight at 4οC. Anti-rabbit IgG HRP-conjugate (Bio-Rad) (1:3000) secondary antibody was incubated for 1 hour at room temperature. Bands were visualized with Clarity™ Western ECL Substrate (BioRad) and imaged using CCD-camera in Syngene imaging system.
Chromatin immunoprecipitation assay
Chromatin immunoprecipitation (ChIP) assay was carried out [43] using a rabbit monoclonal antibody against RUNX2 (Cell Signaling D1H7). Briefly, mesenchymal cells were cross-linked with 0.75% formaldehyde in 1x PBS for 10 min and were lysed with cell lysis buffer containing 50mM HEPES-KOH (pH 7.5), 140mM NaCl, 1mM EDTA (pH 8), 1% Triton X-100, 0.1% sodium deoxycholate, 0.1% SDS and 1% protease inhibitors. Samples were then sonicated to shear the DNA with four rounds of 12 sec pulses each at 50% power output and 90% duty cycle using a Branson Sonicator 200. 50 µg of sonicated DNA was suspended in 1 ml of ChIP dilution buffer containing 1% Triton X-100, 2mM EDTA (pH 8), 20mM Tris-HCl (pH 8), 150mM NaCl, and 1% protease inhibitors for each immunoprecipitation. The precleared samples were then mixed with either the RUNX2 monoclonal antibody or normal rabbit IgG and rotated at 4οC overnight. This was followed by the addition of 40 µl of protein A/G beads to the immunoprecipitated complex and further mixing by rotation at 4οC for 4 h. Beads were then washed four times with low salt wash buffer containing 0.1% SDS, 1% Triton X-100, 2mM EDTA (pH 8), 20mM Tris-HCl (pH 8), and 150mM NaCl and two times with high salt buffer containing 0.1% SDS, 1% Triton X-100, 2mM EDTA (pH 8), 20mM Tris-HCl (pH 8), and 500 mM NaCl at 4οC for 2 min each. The immunoprecipitated complex was eluted using elution buffer containing 1% SDS and 100mM sodium bicarbonate at 30οC for 15 min. The eluted DNA samples were treated with proteinase K (10 µg/ml) and RNase A (20 µg/ml) and incubated at 65οC for 2 h. Following incubation, DNA concentration was measured, and PCR was carried out using the primer set to amplify -295 to +72 bp region of the Htra1 promoter.
Cloning of Htra1 promoter regions
Five different regions of the 1 kb Htra1 promoter were amplified by PCR using Phusion High-Fidelity DNA polymerase (NEB) with the primers listed in Table 2. Using dATP, an A-overhang was introduced to the 3' end of the PCR amplicons by non-proof reading Taq DNA polymerase (Lucigen). The PCR products were then purified by gel extraction and cloned into the pGEM-T easy vector (Promega). From the positive clones, the respective Htra1 promoter fragments were digested with KpnI and XhoI for cloning into the destination pGL3 promoter luciferase vector (Promega). All clones were verified by Sanger sequencing at National Research Council (NRC) sequencing facility (Saskatoon, SK, Canada) and found to match perfectly with the promoter region of Htra1 (M. musculus|chr7|138078749-138079949). Three RUNX2 putative binding sites in the 0.4 kb Htra1 promoter (-399 to +72) at -252 bp (Mut1), -84 bp (Mut2) and +11 bp (Mut3) were mutated alone or in combination using QuikChange Lightning kit (Agilent Technologies) with the primer pairs listed in Table 2.
Luciferase assay
Dual luciferase reporter assay (Promega) was carried out as previously described [44]. Briefly, 1x104 primary mesenchymal progenitor cells were seeded into the white opaque 96-well plate. The cells were transfected with 100 ng of pGL3 firefly luciferase vector containing different fragments of the Htra1 promoter constructs, 100 ng of either the empty vector (pCMV6-myc-ddk) or the Runx2 expression vector; 40 ng of pRL-CMV Renilla luciferase vector was used as the transfection control. After 48 h, cells were lysed with the passive lysis buffer and then firefly luciferase assay reagent II was added to measure the firefly luciferase activity. This was followed by the addition of Stop & Glo reagent and the Renilla luciferase activity was measured. The relative luciferase activity was calculated by normalizing the firefly luciferase activity values to the respective Renilla luciferase values.
Statistical analyses
Statistical analyses were carried out using unpaired two-tailed T-test in the case of two groups. One-way ANOVA with Bonferroni multiple comparison test was used to analyze data with more than two groups, two-way ANOVA was carried out to analyze data with two variables in the study and a p-value of <0.05 was considered significant. Analyses are from at least three independent experiments and the respective number of biological replicates are mentioned in the figure legends.
Results
HTRA1 is expressed in developing bones
To establish a relationship between Htra1 and developing mouse craniofacial bones, the spatial expression of the HTRA1 protein was examined using immunohistochemistry (IHC). IHC staining revealed that HTRA1 was mainly expressed in ossifying regions of the maxilla and mandible in wild-type embryos at E16.5 (Fig. 1A, E). Osteoblast markers RUNX2 and SP7 (formerly called OSX) exhibited similar expression patterns as HTRA1 in the developing maxilla (Fig. 1B, C) and mandible (Fig. 1F, G) region. Negative control IgG staining revealed the specificity of the HTRA1, RUNX2 and SP7 antibodies (Fig. 1D, H). The expression pattern of HTRA1 in ossifying craniofacial regions of the mouse embryo suggests that Htra1 might play a role in osteoblast differentiation of primary progenitor cells of craniofacial mesenchyme.
Htra1 promotes mineralized matrix production and osteogenic gene expression in primary mesenchymal progenitor cells
To test the nature of the primary mesenchymal progenitor cells to form mature osteoblasts, we subjected these cells to osteogenic differentiation for up to 21 days and assessed matrix mineralization using ARS staining at day 15 and day 21. Mesenchymal progenitor cells grown in growth media did not develop any mineralized nodules at day 15 or day 21 (Supplementary Fig. 1A, B – for all supplemental material see www.cellphysiolbiochem.com), while cells differentiated with osteogenic media exhibited matrix deposition and mineralization at day 15 and day 21 (Supplementary Fig. 1C, D). To investigate if Htra1 plays a role in osteoblast differentiation, we overexpressed Htra1 in primary mesenchymal progenitor cells for up to 21 days. A vector containing an inactive proteolytic mutant of Htra1 (Htra1 (mut)) [41] generated by site-directed mutagenesis and an empty vector were used as controls. At day (d)8, ALP staining did not reveal a considerable difference among primary mesenchymal progenitor cells transfected with either the empty vector, the Htra1 vector, or the Htra1 (mut) vector (Fig. 2A, B, C and M). However, overexpression of Htra1 resulted in a clear increase in ALP staining at d15, compared to the empty vector- and Htra1 (mut)-transfected cells (Fig. 2D, E, F and N). In addition, ARS staining reflected increased mineralized matrix in Htra1-overexpressed cells, compared to cells treated with empty vector and Htra1 (mut) at both d15 (Fig. 2G, H, I) and d21 (Fig. 2J, K, L). Quantitation of the ARS-bound matrix revealed a significant increase in the amount of bone matrix in Htra1-overexpressed cells at d15 and d21, compared to both empty vector- and Htra1 (mut)-transfected cells (Fig. 2O, P). In fact, the Htra1 (mut)-transfected cells produced levels of mineralized matrix that were indistinguishable from empty vector-transfected cells. These results demonstrate that Htra1 can increase the production of mineralized bone matrix in primary mesenchymal progenitor cells, and that Htra1 function in this process could be inhibited by mutating the protease domain.
To elucidate a potential signalling pathway stimulated by Htra1 during osteoblast differentiation of mesenchymal progenitor cells, Htra1-responsive genes were identified using an ossification and bone remodelling real-time PCR array. Several genes, such as Fgf9, Ibsp, Vegfa, Col2a1, and Mmp9, were upregulated significantly by 1.98, 2.97, 1.52, 2.33, and 4.04-fold, respectively, in Htra1-overexpressed cells at d7, compared to empty vector-transfected cells (Fig. 3A). Additional osteogenesis-associated genes, such as Fgfr3 and Estrogen receptor beta (Esr2) were upregulated, although not to the level of statistical significance. In contrast, Htra1 downregulated inhibitors of osteogenesis, such as Sost, Calca, and Estrogen receptor alpha (Esr1) 0.23, 0.60, and 0.51-fold, respectively, although Calca reduction was not statistically significant (Fig. 3A). Upregulation of the osteoblast markers Sp7 and Alpl by Htra1 was demonstrated using qPCR. Compared to empty vector-transfected cells, overexpression of Htra1 significantly upregulated Sp7 (1.7-fold at d7 and 1.37-fold at d15) and Alpl (2.1-fold at both d7 and d15) (Fig. 3B, C). In contrast, the Htra1 (mut) vector failed to upregulate the expression of Sp7 and Alpl at d7 and d15 (Fig. 3B, C). These data indicated that Htra1 regulates several osteogenesis-related genes to promote osteoblast differentiation and that mutation of the protease domain of HTRA1 blocks this activity.
RUNX2 interacts with the Htra1 promoter to induce Htra1 expression during osteoblast differentiation
Although Htra1 overexpression upregulated some osteoblast markers, such as Alpl and Sp7, Runx2 was not altered (data not shown). Since RUNX2 and HTRA1 are co-expressed during osteogenesis in vivo (Fig. 1), Runx2 might be upstream of Htra1 during osteoblast differentiation. To investigate this, Runx2 was overexpressed in primary mesenchymal progenitor cells for eight days (Fig. 4A). Overexpression of Runx2 increased the expression of Htra1 to the same extent as Sp7 (1.51-fold; Fig. 4B, C), a known transcriptional target of Runx2 [9, 45]. In addition, the upregulation of Runx2 resulted in a corresponding increase in RUNX2 (1.91-fold; Fig. 4D, E; Supplementary Fig. 2A) and HTRA1 (2.23-fold; Fig. 4F, G; Supplementary Fig. 2B) protein levels, compared to cells transfected with empty vector. These data show that Runx2 can induce Htra1 and HTRA1 expression during osteogenic differentiation. Supporting the idea that RUNX2 regulates Htra1 directly, several putative RUNX2 binding sites were identified in the genomic region 1kb upstream of the Htra1 transcriptional start site (Fig. 5A). To examine if RUNX2 binds to this proximal Htra1 promoter, streptavidin pull-downs were carried out using biotinylated promoter regions of Htra1 on lysates of d5 differentiating cells. Indeed, RUNX2 protein interacted with both 1 kb (-994 bp to +72 bp) and 0.4 kb (-399 bp to +72 bp) Htra1 promoter regions (Fig. 5B; Supplementary Fig. 3A, B). In addition, ChIP analyses confirmed that RUNX2 interacts with the proximal 0.4 kb Htra1 promoter during osteogenic differentiation (Fig. 5C).
To identify more discrete regions of the Htra1 promoter through which Runx2 could promote Htra1 expression, five different regions of the Htra1 promoter were cloned into a luciferase reporter vector: (i) -994 to +72 bp; (ii) -399 to +72 bp; (iii) -811 to -439 bp; (iv) -295 to +72 bp; and (v) -199 to +72 (Fig. 6A). Constructs were transfected individually into primary mesenchymal progenitor cells along with either a Runx2 overexpression or an empty vector. Dual luciferase reporter assay was carried out 48 h after transfection. Overexpression of Runx2 significantly increased luciferase activity of all the Htra1 promoter constructs except the -811 to -439 bp construct (Fig. 6B). Specifically, Runx2 increased expression of the proximal Htra1 promoter fragments -994 to +72 bp, -295 to +72 bp, and -199 to +72 bp by 146%, 145%, and 134%, respectively, with the -399 to +72 bp fragment showing the greatest increase in expression to 192% compared to control (empty vector) levels. These results highlighted the importance of Htra1 promoter fragments that contained putative RUNX2 binding site(s) between -399 to +72 bp in order for Runx2 to induce Htra1 expression during osteoblast differentiation of mesenchymal cells.
To identify critical RUNX2 binding sites for Htra1 promoter activity, putative RUNX2 binding sites on the -399 to +72 bp proximal Htra1 promoter fragment were mutated and Runx2-mediated expression was analyzed. The specific RUNX2 binding sites mutated were at -252 bp (Mut1), -84 bp (Mut2), and +11 bp (Mut3), alone or in combination (Fig. 7A). Overexpression of Runx2 increased the Htra1 promoter activity of wild-type (-399 to +72 bp) and Mut3 (+11 bp) to 165% and 151%, respectively (Fig. 7B), suggesting that the putative RUNX2 binding site at +11 bp was not required for Runx2 activation. However, Mut1 (-252 bp) and Mut2 (-84 bp) constructs, alone or in combination, blocked Runx2-mediated promoter activity of Htra1 (Fig. 6B), suggesting that both of these sites are required for Runx2 activation. Specifically, the Runx2-mediated expression levels of the Mut1 and Mut2 proximal Htra1 promoter regions were 95% and 104%, respectively, of control values (Fig. 7B). Combined proximal Htra1 promoter region mutations (Mut1*Mut2) also failed to increase expression by Runx2 (75%). To confirm that binding of RUNX2 to the -252 bp and -84 bp sites is required for Runx2 activation of the Htra1 promoter, streptavidin pull-down assays were performed using either the biotinylated wild-type Htra1 promoter probes (-399 to +72 bp) or the Mut1, Mut2, or Mut3 probes of the same region. Western blot analysis showed that RUNX2 was able to bind to the wild-type and Mut3 probes in differentiating osteoblasts (Fig. 7C; Supplementary Fig. 3C). In contrast, RUNX2 did not bind to Mut1 or Mut2 probes (Fig. 7C). These data showed that Runx2 induces Htra1 expression during osteoblast differentiation via RUNX2 binding to sites -252 bp and -84 bp at the proximal Htra1 promoter.
Discussion
Despite considerable knowledge of the signalling mechanisms governing osteoblast differentiation, several interacting partners and their role in osteogenesis remain obscure. In this study, we demonstrated that the serine protease Htra1 promotes osteoblast differentiation of primary mesenchymal progenitor cells. In addition, we revealed a specific molecular interaction between RUNX2 and Htra1 during osteoblast differentiation, delineating that RUNX2 binds to the proximal Htra1 promoter to activate Htra1 expression. Our data significantly improve the understanding of the role of Htra1 in osteoblast differentiation and unravel a novel interaction between Htra1 and Runx2.
Conflicting reports on whether Htra1 promotes or inhibits osteoblast differentiation might relate to the specific progenitor cell or the culture conditions in which an osteogenic role for Htra1 was evaluated. Here, the co-expression of HTRA1 and RUNX2 in developing craniofacial bones led to the hypothesis that Runx2-mediated Htra1 expression regulates osteoblast differentiation of craniofacial mesenchymal progenitor cells. Supporting this hypothesis, Htra1 overexpression in primary craniofacial mesenchymal progenitor cells enhanced osteogenic marker expression and secretion of mineralized bone matrix. As such, these findings are consistent with the reported osteoblast-promoting role of HTRA1 in human bone marrow-derived stem cells [20], mouse adipose-derived stromal cells [22], and periodontal ligament cells [46]. Conversely, Htra1 was reported to be an inhibitor of matrix mineralization in murine 2T3 and KusaO cell lines [23, 24]. These cell lines, however, were transformed with SV40, which significantly reduces the expression of Htra1 [47], thus complicating analyses of the role of Htra1 in osteoblast differentiation [20, 48]. In addition, these studies investigated the role of Htra1 after stimulation with BMP2 [23, 24]. Even though the addition of BMP2 can upregulate Htra1 in C2C12 preosteoblasts [49], a potential role for Htra1 in osteoblast differentiation might have been overwhelmed by BMP pathway stimulation, a dominant inducer of osteogenesis. Here, we have used a gain of function approach to show that Htra1 induces osteogenic differentiation of the mesenchymal progenitor cells. A caveat from the current study is the lack of complementary loss of function approach to check if silencing Htra1 inhibits osteogenic differentiation. However, it is known from the literature that silencing of Runx2 during osteogenic differentiation using Runx2 shRNA downregulates Htra1 expression, in addition to several known RUNX2-responsive genes such as AlpI, Mmp13 and Ibsp [11]. Htra1 deficient mice exhibit no obvious alteration in bone phenotype in vivo [48], but other HTRA members might compensate for Htra1 function. HTRA3, which shares the highest homology with HTRA1, is also expressed during bone formation [18, 48]. The role of Htra3 in osteoblast differentiation is unknown, but Htra1 and Htra3 could be functionally redundant in mediating osteogenesis of mesenchymal stem cells.
Regarding the influence of RUNX2 in regulating the expression of other Htra family members, Wu et al. (2014) show that Htra1 is the only Htra family member in the list of RUNX2-responsive genes during osteogenic differentiation in an unbiased microarray approach [11]. In addition, among the Htra family members, Htra1 alone is noted as RUNX2-responsive in bone metastatic prostate cancer cells [50]. RUNX2 does bind to the genomic regions of Htra1, Htra3, and Htra4 during osteogenic differentiation, but RUNX2 only binds to the promoter region of Htra1, while it binds to introns or exons for Htra3 and Htra4 [11]. In fact, most RUNX2-responsive genes exhibit binding sites in the promoter region, rather than introns or exons [6, 8-9, 11]. In this study, we delineated the molecular interactions of RUNX2 at two specific sites in the Htra1 promoter that seem to regulate Htra1 expression during osteoblast differentiation. Other members of the Htra family, including Htra3 and Htra4, do not seem to be RUNX2-responsive, but might function redundantly through different genetic networks that impact bone differentiation. Further work is needed to decipher the role and mechanism involving these Htra family members in an osteoblast-specific compound mutant mice model.
Through what downstream pathways might Htra1 promote osteoblast differentiation of mesenchymal progenitor cells? Osteogenic gene expression analysis performed here revealed that Htra1 upregulated the FGF signalling genes Fgf9 and Fgfr3, and FGF signalling promotes osteogenesis [51]. In addition, HTRA1 proteolytic cleavage of ECM proteins such as Biglycan, Syndecan-4, and Glypican-4 promotes long-range FGF signalling in Xenopus embryos [28]. In turn, FGF signalling is necessary and sufficient for Htra1 expression in chick facial and forelimb mesenchyme [31]. HTRA1 induces adipogenesis via MAP kinase-dependent MMP production in human mesenchymal stem cells [21]. In our study, the induction of Mmp9 expression by Htra1 might be related to the demonstrated upregulation of MMPs by HTRA1 proteolytic cleavage of ECM proteins, such as Fibronectin [21, 27] [25]. Therefore, a FGF-Runx2-Htra1-MMP signalling network could be vital in osteoblast differentiation (Fig. 7D).
Other genes identified here to be downstream of Htra1 during osteoblast differentiation have known osteogenic roles. IBSP expression increased after HTRA1 overexpression in human bone marrow-derived mesenchymal stem cells [20]. HTRA1 and IBSP co-localize during bone regeneration in osteo-induced spheroid cultures, where HTRA1 is hypothesized to fragment IBSP for proper matrix mineralization [20]. This might provide a mechanism for Htra1-induced matrix mineralization in primary craniofacial mesenchymal cells reported here. Also, we found that Htra1 overexpression downregulated Sost, Calca, and Esr1. Sost is an antagonist of WNT signalling and is a negative regulator of bone formation [52], and SOST was previously reported to be inhibited by HTRA1 during osteoblast differentiation of human bone marrow-derived mesenchymal stem cells [20].
Regarding critical upstream regulators of Htra1 during osteoblast differentiation, we showed that RUNX2 is recruited to the proximal Htra1 promoter to activate Htra1 expression. Runx2 plays a critical role in chondrocyte maturation and bone formation [1]. RUNX2 binds to RUNT binding site RCCRC(A/T) [10] in the promoter regions of downstream genes to drive osteoblast-specific expression [5, 53, 54]. Several targets of RUNX2 during osteogenesis are not yet functionally characterized [11]. Recently, ChIP-seq analysis of RUNX2 during osteoblast differentiation identified several putative target genes, including Htra1 [11]. Since false-positives from ChIP-seq analysis are generally unavoidable, characterization of putative downstream targets is essential. In our study, RUNX2 was demonstrated by multiple binding and reporter assays to interact with the proximal -399 to +72 bp region of the Htra1 promoter to promote Htra1 expression during osteoblast differentiation. Furthermore, mutation analyses revealed two RUNX2 binding sites (-252 bp and -84 bp) required for the activation of Htra1 osteogenic expression. The proximity of these binding sites to the transcription start site is comparable to the promoter interaction of RUNX2 with many other target genes, such as Col10a1 [55], Ibsp [56], and Mmp9 [57]. As such, the data here uncovers a previously unknown functional interaction between RUNX2 and Htra1.
The functional interaction of RUNX2 and Htra1 might be important in several pathological conditions, including cancer. Runx2 is a negative regulator of cell proliferation, and suppression of Runx2 expression is linked to tumorigenesis [58, 59]. Moreover, forced expression of Runx2 inhibits osteosarcoma cell proliferation [60]. Htra1 is also a tumor suppressor gene, inhibiting cancer cell proliferation [61, 62]. Therefore, considering the role of Htra1 as a tumor suppressor and modulator of MMP expression, the regulation of Htra1 by Runx2 identified here in osteoblasts might have physiological and pathological relevance in other cell lineages. Further studies are needed to address the interaction of RUNX2 and Htra1 in other systems, such as tumorigenesis.
Conclusion
In conclusion, our data show that Htra1 is a novel direct downstream target of RUNX2, which binds to the Htra1 promoter at -252 bp and -84 bp, and Htra1 promotes osteoblast differentiation of primary mesenchymal progenitor cells.
This work was supported by Discovery Grant 171317-2012 to AJN from the Natural Sciences and Engineering Research Council of Canada (NSERC) and, in part, by a Subtelny Orthodontic Clinical Research Grant from American Cleft Palate-Craniofacial Association (ACPA-CPF), USA to PPRI and a Canadian Institute for Health Research grant 148683 to BFE.
PPRI was a recipient of the Apotex Graduate Scholarship Award from the College of Pharmacy and Nutrition, University of Saskatchewan; Graduate Research fellowship and Saskatchewan Innovation and Opportunity Scholarships from the College of Graduate Studies and Research, University of Saskatchewan. MPT was a recipient of the Dean’s Graduate Scholarship, University of Saskatchewan and an endMS Doctoral Studentship award from the Multiple Sclerosis Society of Canada (MSSC). The authors acknowledge Larhonda Sobchishin for providing technical expertise and Rongqi Liu and Nasim Rostampour for technical assistance.
Author contributions
PPRI, BFE and AJN conceived the study. AJN coordinated the study. PPRI designed, performed all the experiments, analyzed the data and wrote the manuscript. PPRI and MPT performed the luciferase assay and the osteogenesis array experiments. BFE and MPT proofed and revised the manuscript for critical content and interpretation of data and approved the final version for submission.
This manuscript is dedicated to the memory of Dr. Adil J. Nazarali, who passed away before the submission of the manuscript.
Disclosure Statement
The authors declare no conflicts of interest.
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