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Runx2 Protein Represses Axin2 Expression in Osteoblasts and Is Required for Craniosynostosis in Axin2-deficient Mice*

Background: Runx2 and Axin2 are required for proper skeletal development. Results: Runx2 and Hdac3 repress Axin2 transcription in osteoblasts. Runx2 insufficiency prevents craniosynostosis in Axin2-deficient mice. Conclusion: A Runx2-Axin2 regulatory mechanism controls the pace of calvarial bone formation. Significance: The molecular and functional interplay between Runx2 and Axin2 controls the rate of cranial suture closure. Similar interactions may occur during skeletal development and carcinogenesis. Runx2 and Axin2 regulate craniofacial development and skeletal maintenance. Runx2 is essential for calvarial bone development, as Runx2 haploinsufficiency causes cleidocranial dysplasia. In contrast, Axin2-deficient mice develop craniosynostosis because of high β-catenin activity. Axin2 levels are elevated in Runx2−/− calvarial cells, and Runx2 represses transcription of Axin2 mRNA, suggesting a direct relationship between these factors in vivo. Here we demonstrate that Runx2 binds several regions of the Axin2 promoter and that Runx2-mediated repression of Axin2 transcription depends on Hdac3. To determine whether Runx2 contributes to the etiology of Axin2 deficiency-induced craniosynostosis, we generated Axin2−/−:Runx2+/− mice. These double mutant mice had longer skulls than Axin2−/− mice, indicating that Runx2 haploinsufficiency rescued the craniosynostosis phenotype of Axin2−/− mice. Together, these studies identify a key mechanistic pathway for regulating intramembranous bone development within the skull that involves Runx2- and Hdac3-mediated suppression of Axin2 to prevent the untimely closure of the calvarial sutures.

Runx2 and Axin2 regulate craniofacial development and skeletal maintenance. Runx2 is essential for calvarial bone development, as Runx2 haploinsufficiency causes cleidocranial dysplasia. In contrast, Axin2-deficient mice develop craniosynostosis because of high ␤-catenin activity. Axin2 levels are elevated in Runx2 ؊/؊ calvarial cells, and Runx2 represses transcription of Axin2 mRNA, suggesting a direct relationship between these factors in vivo. Here we demonstrate that Runx2 binds several regions of the Axin2 promoter and that Runx2-mediated repression of Axin2 transcription depends on Hdac3. To determine whether Runx2 contributes to the etiology of Axin2 deficiencyinduced craniosynostosis, we generated Axin2 ؊/؊ :Runx2 ؉/؊ mice. These double mutant mice had longer skulls than Axin2 ؊/؊ mice, indicating that Runx2 haploinsufficiency rescued the craniosynostosis phenotype of Axin2 ؊/؊ mice. Together, these studies identify a key mechanistic pathway for regulating intramembranous bone development within the skull that involves Runx2-and Hdac3-mediated suppression of Axin2 to prevent the untimely closure of the calvarial sutures.
Craniosynostosis is a common birth defect, appearing in ϳ1 of 2500 live births (1,2). This condition is characterized by premature fusion of one or more cranial sutures, which causes abnormal skull shapes and cognitive disabilities if surgical reconstructions are not performed (1,2). Considerable progress has been made in understanding the genetic causes of craniosynostosis, as mutations in MSX2, TWIST1/2, FGFR1/ 2/3, and EFNB2 are linked to various craniosynostosis syndromes (3)(4)(5). Despite these advances, the etiology of ϳ85% of non-syndromic cases remains unknown (3).
Mouse models recapitulate human craniosynostosis syndromes and provide substantial mechanistic insight into disease mechanisms. For example, Twist haploinsufficiency (6,7) and introduction of activating mutations in Fgfr1 (8) and Fgfr2 (9) faithfully mimic features of human craniosynostosis in developing mice. Other mouse models reveal new candidate genes that were not previously linked to human craniosynostosis syndromes. One such gene is Axin2 (10 -12), an intracellular negative feedback regulator of Wnt, Lrp5/6, and ␤-catenin signaling (10 -12, 14, 16). Axin2-deficient mice, created by insertion of LacZ into exon 2 (hereafter referred to as Axin2 Ϫ/Ϫ mice), feature up-regulated ␤-catenin signaling in the cranial sutures (10,12,16). These mice develop craniosynostosis because of enhanced mesenchymal progenitor cell proliferation in the cranial sutures and rapid differentiation of progenitor cells into matrix-producing osteoblasts (10 -12).
Skeletal development and bone mass maintenance are complex processes involving multiple levels of regulation by transcription factors and epigenetic modifications to control gene expression and coordinate activities of bone-residing cells. Runx2 is a master osteoblast transcription factor that is essential for proper bone development. Runx2-deficient mice die at birth because of a lack of skeletal ossification, and Runx2 haploinsufficient mice and humans develop an osteopenic phenotype and features of cleidocranial dysplasia, including delayed calvarial development (17)(18)(19). A 30% reduction in Runx2 levels is sufficient to induce a cleidocranial dysplasia phenotype in mice (19). Interestingly, although Runx2 insufficiency impairs cranial suture closure, increased Runx2 expression coincides with the increased osteoblast differentiation that causes premature suture closure in many models of craniosynostosis (20 -24), suggesting convergence on a key regulatory role for Runx2 in cranial suture biology and, in particular, in craniosynostosis etiology. This is supported by a recent finding of RUNX2 duplication in human metopic craniosynostosis (25) and studies showing that overexpression of Runx2 in the condensing calvarial mesenchyme causes craniosynostosis in mice (26).
The opposing cranial phenotypes of the Axin2-deficient (craniosynostosis) and Runx2 haploinsufficient (cranial dysplasia) mice suggest potential interaction between these two factors in vivo, an idea supported by our observations that Axin2 protein levels were up-regulated in Runx2-null osteoblasts and that Runx2 repressed Axin2 transcription (27). Runx2 represses transcription by recruiting histone deacetylases (Hdacs) 2 to gene promoters. We showed that Hdac3 binds to the N terminus of Runx2 and represses Runx2-dependent activation of Bglap (28). Although suppression of Hdac3 in committed osteoblasts accelerates differentiation (28), conditional deletion of Hdac3 in osteochondral progenitor cells (from Osterix-Cre:Hdac3 fl/fl conditional knockout mice, hereafter referred to as Hdac3 CKO Osx mice) causes calvarial, axial, and appendicular bone loss in both trabecular and cortical compartments because of reduced bone formation (29). Interestingly, Axin2 was one of the most highly expressed genes in Hdac3 CKO Osx calvaria as compared with wild-type littermates, suggesting that the deleterious osteoblast phenotype in these mice could be due, in part, to a suppression of Wnt/␤-catenin signaling via up-regulation of its inhibitor Axin2.
To determine whether Axin2, Runx2, and Hdac3 directly interact and are components of the same molecular pathway that regulates suture closure, we performed molecular experiments and crossed the Axin2 Ϫ/Ϫ and Runx2 ϩ/Ϫ mouse models to generate double mutant Axin2 Ϫ/Ϫ :Runx2 ϩ/Ϫ mice. These studies demonstrate that Runx2 haploinsufficiency rescues the craniosynostosis phenotype in Axin2 Ϫ/Ϫ mice and that Runx2 directly represses Axin2 expression in an Hdac3-dependent manner.
Transient Transfections, Adenoviral Transductions, and PCR-C2C12 cells were transfected with Lipofectamine (Invitrogen) in 6-well plates with 1 g of Runx2 expression plasmid (Runx2-I, P2 promoter-dependent isoform, beginning with the amino acids MRIPV) or 1 g of a control plasmid (pcDNA3, Invitrogen). RNA was harvested with TRIzol reagent (Invitrogen) 48 h after transfection and was reverse transcribed into cDNA using the SuperScript III first-strand synthesis system (Invitrogen). Expression levels of mRNAs for Runx2, Axin2, and Gapdh were measured by real-time PCR. Primer sequences were reported previously (32,33). Reactions were performed using 37.5 ng of cDNA/15 l with Bio-Rad iQ SYBR Green Supermix and the Bio-Rad MyiQ single color real-time PCR detection system. Transcript levels for each gene of interest were normalized to the reference gene Gapdh. Gene expression levels were quantified using the 2ˆ(-⌬⌬ Ct) method (17).
To observe the effects of reintroduction of Runx2 expression into Runx2 Ϫ/Ϫ cells, control and Runx2 adenoviruses were prepared as described previously (31). WT and Runx2 Ϫ/Ϫ cells were infected with each indicated virus at a multiplicity of infection of 100. RNA was harvested with TRIzol reagent (Invitrogen) 24 h after infection. RNA was reverse-transcribed, and gene expression levels for Runx2, Axin2, Hdac3, and Gapdh were quantified as described above.
ChIP-Seq-MC3T3-E1 cells cultured in osteogenic medium for 0, 9, or 28 days were washed with 37 ºC PBS and then fixed at room temperature with 1% formaldehyde for 8 min. Cells were lysed according to a published protocol (35). Nuclear lysates were sonicated to generate chromatin fragments ranging from 200 -600 bp. Chromatin was then precipitated with Runx2 antibody (M-70, Santa Cruz Biotechnology, Inc.) or control normal IgG followed by protein G Dynabeads (Invitrogen). Runx2-precipitated chromatin was washed, and cross-links were reversed overnight at 65 ºC. DNA samples were subsequently treated with RNase A followed by proteinase K at 55 ºC. DNA was recovered by phenol/chloroform extraction followed by ethanol precipitation and transformed into a sequencing library according to the Illumina DNA library preparation protocol. A fraction of DNA fragments with genomic inserts at 200 Ϯ 50 bp were sent for sequencing on an Illumina Genome Analyzer II operated by the Deep Sequencing Core Facility at the University of Massachusetts Medical School. Base calls and sequence reads were generated by Illumina CASAVA software (version 1.6), and short reads were aligned to the mouse mm9 genome using Bowtie aligner (version 0.12.7) (36). Values shown for Axin2 represent two biological replicates. Runx2enriched peaks were analyzed by model-based analysis of ChIP-Seq (MACS) (37).
Construction of Plasmids and Luciferase Reporter Assays-The 5.6-kbp Axin2-luciferase construct (32) containing the mouse Axin2 promoter, exon 1, and intron 1 was obtained from Addgene (plasmid no. 21275). To focus on the Axin2 promoter regions showing Runx2 association identified in ChIP and ChIP-Seq assays, the full-length Axin2-Luc construct (hereafter referred to as Axin2(FL)) was cut with restriction enzymes (Acc65I and HindIII) to generate smaller inserts. Deletion constructs were ligated into the pGL3 basic vector (Promega). Sitedirected mutagenesis of the four Runx2 consensus sites was performed with the QuikChange II site-directed mutagenesis kit (Agilent Technologies, catalog no. 200523). The effects of Runx2 on these constructs were determined using a pCMV-HA-Runx2 expression plasmid (Runx2 isoform 1, beginning with the amino acids MASNS) that has been described previously (27,28). Mutant pCMV-HA-MASNS(M182R) was generated using the site-directed mutagenesis kit described above and gene-specific oligonucleotides containing a mutation in the DNA binding site (5Ј-CGAAATGCCTCCGCTGTT AGGAA-AAACCAAGTAGCCAGG-3Ј and 5Ј-CCTG GCTACTTG-GTTTTTCCTAACAGCGGAGGCATTTCG-3Ј).
C2C12 cells were transfected with Lipofectamine (Invitrogen) in 12-well plates with 200 ng of each Axin2-luciferase promoter construct, as indicated, and pRL-null (10 ng). Runx2 expression plasmids (300 ng) were added as indicated (28). Following a 48-h incubation at 37°C, luciferase activity in 20 l of cell lysate was measured using the dual luciferase assay system (Promega) and a Glomax 96 microplate luminometer. All transfections were performed in triplicate, and data were normalized to the activity of Renilla luciferase.
Animal Studies-All animal research was conducted according to guidelines provided by the National Institutes of Health and the Institute of Laboratory Animal Resources National Research Council. The Mayo Clinic Institutional Animal Care and Use Committee approved all animal studies. Animals were housed in an accredited facility under a 12-hour light/dark cycle and were provided with water and food (PicoLab Rodent Diet 20, LabDiet) ad libitum.
Hdac3 CKO Osx Mice-Hdac3 CKO Osx , heterozygous and wild-type littermate mice were generated and genotyped as described previously (29). Calvarial explants were removed from 5-or 6-day-old mice, dissected free of soft tissues, and incubated in collagenase digestion medium before flash-freezing in liquid nitrogen, as reported previously (38). For mRNA analyses, bone explants were homogenized in TRIzol using a high-speed disperser (Ultra-Turrax T25, IKA). RNA was extracted and purified from the ground tissue with TRIzol reagent (Invitrogen) and was reverse-transcribed into cDNA and analyzed via real-time PCR as described above. For protein isolation, calvarial explants were crushed in liquid nitrogen with a mortar and pestle and were suspended in modified radioimmune precipitation assay buffer supplemented with protease inhibitors. Protein extracts from calvarial explants were sonicated and resolved by SDS-PAGE. Immunoblotting was performed with antibodies recognizing Axin2 (1:1000, Abcam, catalog no. Ab32197) and mSin3A (1:1000, Santa Cruz Biotechnology, Inc., K-20). mSin3A is a ubiquitous global transcriptional corepressor and used here as a loading control.
Cranial Morphology and Suture Histology in Axin2 Ϫ/Ϫ : Runx2 ϩ/Ϫ Mice-Cranial morphology was assessed via microcomputed tomography and image analysis software at 4 weeks of age, when Axin2 Ϫ/Ϫ mice demonstrate a significant reduction in skull length (15), and at 6 months of age, to assess skull morphology in adult mice. Mice were euthanized and skulls were fixed in 70% ethanol. Skulls were scanned in 70% ethanol on a CT35 scanner (Scanco Medical AG, Basserdorf, Switzerland) at 20 -30 m resolution (energy settings 70 kV, 114 A) and reconstructed with the software of the manufacturer using a threshold value of 150. Skull coronal length (coronal suture to nose tip) and width (measured at the distal aspect of the zygomatic process across the parietal bones) were quantified using image analysis software (Bioquant Osteo, Nashville, TN).
For analysis of suture histology, skulls were embedded in glycolmethacrylate. Micro-CT scans of the embedded skulls were used to precisely locate the posterior frontal suture (analogous to the human metopic suture), as this suture closes prematurely in Axin2-deficient mice and contributes to their craniosynostosis phenotype (12). Thin (5-m) sections were mounted and stained with Gomorri's trichrome (39) to highlight bone and connective tissues. Cell and bone morphology were examined at ϫ600 magnification.
BMSC Growth, Mineralization, and Gene Expression-BMSC cultures were chosen as a model to study the in vitro osteoblastic differentiation patterns of progenitor cells from mice of each genotype because these cultures can be performed on tissue explants from mice at 4 weeks of age (or older), permitting tissue harvests at time points that corresponded directly with ages of morphological analyses of the skull. BMSC isolated from 4-week-old mice were seeded in osteogenic medium and cultured for 28 days to promote calcified matrix production, then fixed in 10% neutral buffered formalin and stained with 2% Alizarin red. Calcified matrix production was quantified as the percentage of Alizarin red-positive area within each well with image analysis software (Bioquant Osteo). For gene expression studies, RNA was harvested with TRIzol reagent (Invitrogen) after 7, 14, or 21 days in osteogenic culture. RNA was reverse transcribed and expression levels of Runx2 or Bglap (genes associated with osteoblast maturation) were quantified as described above and previously (32).
Statistics-Statistics were performed with JMP 9.0 statistical analysis software (SAS Institute, Inc., Cary, NC). Data were compared between groups within each experiment with Student's t-tests (when only two groups were compared) or ANOVA (when three or more groups were compared) followed by Student's t post-hoc comparisons. A significance of p Ͻ 0.05 was used for all comparisons.

Runx2 Regulates Axin2 Expression in Osteoblasts-To
understand the genetic interplay between Axin2 and Runx2, we examined Axin2 transcripts using mRNA isolated from calvarial cells of Runx2 Ϫ/Ϫ E17.5 embryos (40). Axin2 mRNA levels were increased 5.5-fold in Runx2 Ϫ/Ϫ cells as compared with wild-type primary calvarial cells (Fig. 1A). The finding that Runx2 controls Axin2 mRNA corroborates our previous study showing that Axin2 protein levels are elevated in Runx2 Ϫ/Ϫ cells (27) and indicates that Runx2 controls Axin2 protein levels through its ability to affect transcription and mRNA accumulation.
We hypothesized that Axin2 was a direct transcriptional target of Runx2 and identified four Runx2 consensus binding sequences (i.e. ACCACA or TGTGGT (41)) in the Axin2 promoter (15), which were designated as putative Runx2 binding sites 1 to 4 (Fig. 1B). These sequences were tested for Runx2 binding using an EMSA with lysates from C2C12 cells overexpressing Runx2. Runx2 bound the first site with high affinity and the fourth site with moderate affinity (Fig. 1B). Weak in vitro binding was also suggested at sites 2 and 3 in the presence of a Runx2-specific antibody (␣-AML3) that caused a supershift. However, in ChIP assays with differentiated osteoblasts, Runx2 associated with the Axin2 promoter at all four sites, and, in particular, strongly bound at sites 2 and 3 (Fig. 1C). Association of Runx2 with regions of the Axin2 promoter containing Runx2 binding sites was enriched several fold as compared with a downstream region lacking a Runx2 binding site (Fig. 1D). These data indicate that Runx2 associates with the Axin2 promoter in differentiating osteoblasts.
To further examine Runx2's association with the Axin2 promoter during osteoblast differentiation, a ChIP sequencing analysis was performed on MC3T3 cells differentiated for 0, 9, or 28 days (Fig. 2A). Association of Runx2 with the four Runx2 consensus motifs characterized above (hereafter referred to as region 1) was confirmed. This unbiased and genomic approach revealed another region of the Axin2 promoter with relatively stronger Runx2 association in differentiating osteoblasts ( Fig.  2A). This region (hereafter referred to as region 2) of Runx2 association consisted of three strong peaks located within 2 kbp of the Axin2 transcription start site. The first two of these peaks (hereafter referred to as peak i and peak ii) were validated by ChIP-PCR (Fig. 2B). Notably, a consensus Runx2 binding motif was not present in these DNA sequences, despite the strong association of Runx2 with region 2 of the Axin2 promoter. Analysis of region 2 DNA sequences with the Transcription Element Search System (42) revealed potential binding elements for other factors known to interact with Runx2 (e.g. Lef/ Tcf transcription factors, C/EBP␤). However, the cofactors with which Runx2 interacts in Region 2 remain unknown at present.

Runx2 Represses Axin2 in Osteoblasts
Runx2 Represses Axin2 Transcription-To test the role of Runx2 in regulating Axin2 transcription, an Axin2-luciferase reporter construct containing the 5.6-kbp region described above (15) was cotransfected with Runx2 expression plasmids into C2C12 cells. Runx2 repressed the basal activity of the fulllength Axin2 promoter construct by 60 -70% (Fig. 2C), consistent with our previous observations. Runx2 also repressed luciferase reporters driven by either region 1 or region 2 of the Axin2 promoter (Fig. 2C).
We next sought to determine whether inducing Runx2 expression would suppress endogenous Axin2 levels. Overexpression of Runx2 in C2C12 cells reduced Axin2 transcript levels by ϳ50% as compared with cells transfected with a control plasmid (Fig. 2D). Moreover, upon induction of osteoblastic differentiation, Runx2 expression levels increased 3.5-fold in primary mouse bone marrow stromal cells and coincided with 80% reductions in endogenous Axin2 expression (Fig. 2E). Together, these data demonstrate that Runx2's suppression of Axin2 may play a role in the early stages of osteoblastic lineage commitment and differentiation.
To determine whether DNA binding was required for repression of Axin2 by Runx2, we expressed a mutant Runx2 protein that contains a point mutation in the Runt domain, M182R, that is linked to human cleidocranial dysplasia and prevents DNA binding (43,44). Runx2-M182R was less effective than Runx2 at repressing the FL and region 1 constructs and completely failed to repress region 2, confirming that the Runt domain contributes to the repression of Axin2 by Runx2 in both regions of the promoter and is required for repression in region 2 (Fig. 2C). Interestingly, Runx2 and Runx2-M182R repressed the Axin2-region 1 reporter construct even when the consensus Runx2 binding sites were mutated (Axin2 (mutReg1)-Luc), suggesting that repression of Axin2 by Runx2 may require the cooperation of other transcription factors.
Runx2 Represses Axin2 Transcription in an Hdac3-dependent Manner-We and others demonstrated previously a direct interaction between Hdac3 and Runx2 in osteoblasts (28,45). To test the role of Hdac3 in Runx2-directed repression of Axin2 transcription, we collected protein and mRNA from calvarial bones or osteoblasts of Hdac3 CKO Osx animals or control littermates. Axin2 mRNA was increased 3.8-fold in Hdac3 CKO Osx calvarial osteoblasts (Fig. 3A) and was elevated in calvarial explants (B) as compared with wild-type littermates. Axin2 protein levels were also elevated in Hdac3 CKO Osx cal-  FEBRUARY 22, 2013 • VOLUME 288 • NUMBER 8 varial explants (Fig. 3C). These experiments demonstrate that Runx2 and Hdac3 actively repress Axin2 expression.

Runx2 Represses Axin2 in Osteoblasts
In an attempt to rescue the high Axin2 expression levels in Runx2 Ϫ/Ϫ calvarial osteoblasts, we transduced WT and Runx2 Ϫ/Ϫ cells with either control or Runx2 adenovirus. As expected, Axin2 levels were up-regulated in Runx2 Ϫ/Ϫ cells infected with a control adenovirus. Unexpectedly, although the transduction was successful at increasing Runx2 expression levels, Axin2 levels remained high in Runx2 Ϫ/Ϫ cells treated with Runx2 adenovirus (Fig. 3D). Further analyses revealed that

Runx2 Represses Axin2 in Osteoblasts
Hdac3 levels were suppressed in Runx2 Ϫ/Ϫ cells and remained low in Runx2 Ϫ/Ϫ cells infected with Runx2 adenovirus (Fig.  3D), indicating that Hdac3 is essential to the mechanism by which Runx2 represses Axin2 transcription.
Runx2 Haploinsufficiency Rescues Several Aspects of the Craniosynostosis Phenotype of Axin2-deficient Mice-To determine whether Axin2 and Runx2 are components of the same molecular pathway in vivo, Runx2 ϩ/Ϫ and Axin2 Ϫ/Ϫ mice were crossed to generate double-mutant Axin2 Ϫ/Ϫ :Runx2 ϩ/Ϫ mice. We limited studies to four genotypes of interest (WT mice, Runx2 ϩ/Ϫ and Axin2 Ϫ/Ϫ single mutants, and double mutant Axin2 Ϫ/Ϫ :Runx2 ϩ/Ϫ mice) because Axin2 haploinsufficient mice do not present a craniosynostosis phenotype (12). Skull morphology was quantified from micro-CT reconstructions at 4 weeks and 6 months of age. At 4 weeks of age, Axin2 Ϫ/Ϫ single mutants demonstrated reduced skull length, measured from the coronal suture to nose tip (-17%), as compared with wildtype animals (17)(18)(19) (Fig. 4, A, C, and D), which is consistent with previous studies (12). Axin2 Ϫ/Ϫ mice also featured an increase in skull width across the parietal bones (Fig. 4B). Micro-CT and histological analyses revealed a near absence of the jugum limitans and fully fused posterior frontal suture in the Axin2 Ϫ/Ϫ mice at 4 weeks of age (Fig. 4, D and E), indicative of craniosynostosis. Skull lengths and widths were statistically similar in 4 week-old Runx2 ϩ/Ϫ mice as compared with wildtype mice. Interestingly, 4-week-old double mutant Axin2 Ϫ/Ϫ : Runx2 ϩ/Ϫ mice had skulls that were longer and narrower than Axin2 Ϫ/Ϫ mice. The average length of the skull was significantly increased by 12%, and skull width was significantly decreased by 5% in the Axin2 Ϫ/Ϫ :Runx2 ϩ/Ϫ mice as compared with the Axin2 Ϫ/Ϫ mice (Fig. 4, A-C). These measurements in the double mutant animals were statistically comparable with Runx2 ϩ/Ϫ and WT mice. Micro-CT and histological analyses revealed that the posterior frontal suture of the Axin2 Ϫ/Ϫ : Runx2 ϩ/Ϫ mice remained patent and morphologically mimicked that of the Runx2 ϩ/Ϫ mice (Fig. 4, D and E), providing a FIGURE 2. Runx2 binds and represses two distinct regions of the Axin2 promoter. A, chromatin immunoprecipitations were performed with a Runx2 antibody in MC3T3-E1 cells at various phases of osteoblastic differentiation: day 0 (undifferentiated/proliferation), day 9 (matrix deposition), and day 28 (matrix mineralization). Immunoprecipitated DNA was analyzed with ChIP-Seq. Region 1 contains the four consensus Runx2 binding motifs characterized in EMSA and ChIP assays. Three strong peaks of Runx2 association were located in region 2 despite the fact that this segment of the Axin2 promoter does not contain any consensus Runx2 binding motifs. B, real-time PCR results from ChIP assays within region 2. Association of Runx2 with DNA in peaks i and ii is enriched severalfold compared with a downstream region (ϳ1 kbp past peak ii) lacking Runx2 association in the ChIP-Seq assay (No ChIP-Seq association). Runx2 immunoprecipitations (IP) were performed with the D130-3 antibody (MBL). Isotype control IgG2b immunoprecipitations are negative controls. C, constructs encoding full-length Runx2 (Runx2-II, starting with the amino acids MASNS) or a mutated form of Runx2-II (Runx2-M182R) containing a point mutation in the runt domain that prevents Runx2 from binding DNA) were transfected into C2C12 cells along with the full-length Axin2-luciferase reporter construct (Axin2(FL)) or truncated constructs featuring region 1 or region 2 of the Axin2 promoter. Bars with different superscript letters are statistically different from one another. Luc, luciferase. D, Runx2 overexpression suppressed endogenous expression levels of Axin2 by ϳ50% as compared with control C2C12 cells transfected with a pcDNA3 plasmid (Control). *, p Ͻ 0.05; **, p Ͻ 0.08 versus control. SEM, standard error of the mean. E, primary mouse BMSC were cultured in osteogenic medium for 3 days to induce osteoblastic differentiation. Endogenous Runx2 and Axin2 expression were quantified and compared with undifferentiated (Day 0) cells. *, p Ͻ 0.05 versus day 0 cells.  FEBRUARY 22, 2013 • VOLUME 288 • NUMBER 8 mechanism by which the gross morphology of the craniosynostosis phenotype of the Axin2 Ϫ/Ϫ single mutant animals was rescued. Taken together, these data indicate that Runx2 is essential for Axin2-deficiency induced premature fusion of the posterior frontal suture. Of note, the jugum limitans was absent in the double mutant mice, suggesting that not all of the morphological features of Axin2 deficiency induced craniosynostosis are rescued by reducing Runx2 levels. At 6 months of age, Axin2 Ϫ/Ϫ mice retained the shortened skull morphology that was first observed at 4 weeks of age. Surprisingly, skull lengths also were shorter in both Runx2 ϩ/Ϫ and in Axin2 Ϫ/Ϫ :Runx2 ϩ/Ϫ as compared with WT mice, possibly as a consequence of the cleidocranial dysplasia phenotype in these mice.

Runx2 Represses Axin2 in Osteoblasts
Bone Marrow-derived Osteoblasts from Axin2 Ϫ/Ϫ :Runx2 ϩ/Ϫ Mice Produce a Mineralized Matrix but Show Decreased Osteoblastic Gene Expression-BMSCs were obtained from femurs and tibias of 4-week-old mice and cultured in osteogenic medium to investigate cellular differentiation patterns. Mineralization assays revealed a severe reduction in the calcified matrix produced by osteoblast-differentiated BMSC from Runx2 ϩ/Ϫ mice as compared with WT animals (Fig. 5A). Neither osteoblasts derived from Axin2 Ϫ/Ϫ nor Axin2 Ϫ/Ϫ : Runx2 ϩ/Ϫ mice were impaired in their ability to produce a calcified matrix (Fig. 5A). Gene profiling showed that osteoblasts derived from Axin2 Ϫ/Ϫ :Runx2 ϩ/Ϫ mice produced substantially less Runx2 and Bglap as compared with osteoblasts from Axin2 Ϫ/Ϫ single mutant mice and instead mimicked temporal gene expression patterns of cultures from WT mice (Fig. 5B). This result suggests that Runx2 insufficiency relieves the craniosynostosis phenotype in the Axin2 Ϫ/Ϫ mice by reducing osteoblastic gene expression to normal levels in progenitor cells.

DISCUSSION
Runx2 and Axin2 are key regulators of craniofacial development and skeletal maintenance with physiologically opposing functions. Runx2 is a positive master regulator of osteoblastspecific transcription that is required for commitment of mesenchymal progenitors to the osteoblast lineage and for osteoblastic differentiation (17,18,46). Axin2 is a negative intracellular feedback inhibitor of Wnt/␤-catenin signaling (15), a pathway that stimulates proliferation and survival of osteoblast progenitors (47) and facilitates the coupling of bone formation to bone resorption via regulation of osteoprotegerin (Opg) expression in osteoblasts (48). Our data demonstrate that the opposing activities of Runx2 and Axin2 are functionally linked in osteoblasts through a Runx2-and Hdac3-mediated transcriptional mechanism. Both Runx2 Ϫ/Ϫ and Hdac3 CKO Osx calvarial osteoblasts express more Axin2 than WT cells, which may be explained by a loss of the repressive effects of Runx2 and Hdac3 on Axin2 transcription. In vivo, introduction of Runx2 haploinsufficiency onto the Axin2 Ϫ/Ϫ background rescued the craniosynostosis phenotype of the Axin2deficient animals. In vitro studies with BMSCs cultured in osteogenic media suggest that the skull morphology rescue in Axin2 Ϫ/Ϫ :Runx2 ϩ/Ϫ mice may be due to reduced osteoblastic activity, as expression levels of characteristic osteoblastic genes were lower in cells from Axin2 Ϫ/Ϫ :Runx2 ϩ/Ϫ mice as compared with Axin2 Ϫ/Ϫ single mutants. On the basis of these data, we propose a model where Axin2 and Wnt/␤-catenin signaling are upstream of Runx2 in mesenchymal progenitor cells but Runx2 directly represses Axin2 transcription during the early stages of progenitor cell commitment to the osteoblastic lineage and to osteoblastic differentiation, reflecting developmental feedback regulation (Fig. 6). This model is compatible with the finding that Runx2 expression is induced by canonical Wnt signaling (49) and is supported by our data demonstrating that Runx2 represses Axin2 in a manner dependent on Hdac3 in osteoblasts. Our data are consistent with the idea that Runx2 plays a central role in the etiologies of several different forms of craniosynostosis. For example, Runx2 is downstream of the zinc finger transcription factor GLI3, another factor recently linked to craniosynostosis in mice. Gli3-deficient mice develop craniosynostosis in the interfrontal and lambdoid sutures, and removing one allele of Runx2 or increasing Twist1 expression (which subsequently represses Runx2) rescued this phenotype (22,50).
Although Axin2 has not yet been linked to a specific craniosynostosis syndrome in humans, other clinical observations support an inverse link between Runx2 and Axin2 in human craniofacial development. RUNX2 haploinsufficiency in humans causes the formation of supernumerary teeth (46), whereas AXIN2 mutations are associated with tooth agenesis in humans (51). Biological interaction between Axin2, Runx2, and Hdac3 may also factor into disease mechanisms in cancer. Inactivating mutations in AXIN2 are associated with an increased incidence of several types of cancer because of unchecked Wnt signaling (51)(52)(53)(54), whereas RUNX2 is highly expressed in breast and prostate cancers predisposed to skeletal metastasis (55,56), and HDAC3 is one of the most commonly up-regulated genes in human solid tumors (57). These observations are consistent with our data showing that Runx2 represses Axin2 in an Hdac3dependent manner in vivo and suggest that this interaction may have biological relevance with respect to modulating Wnt/␤catenin signaling in both skeletal development and cancer.
The inhibitory effects of Axin 2 on ␤-catenin signaling modulate bone formation in the cranial sutures and prevent premature suture closure (10 -12, 21). In mice, the posterior frontal suture (analogous to the human metopic suture) is the only cranial suture that fuses naturally (58). Axin2 expression in this suture diminishes over time prior to the onset of fusion (11,12) and is inversely correlated with the amount of activated ␤-catenin present in the suture mesenchyme (11). Constitutive activation of ␤-catenin in Axin2-expressing cells is sufficient to recapitulate the craniosynostosis phenotype of the Axin2-deficient mice (21). It is possible that Runx2 regulates suture fusion by repressing Axin2 expression to facilitate a temporal increase in ␤-catenin signaling that promotes bone formation (Fig. 6). Hdac3 also interacts with the canonical Wnt/␤-catenin signaling pathway in that suppression of Hdac3 prevents nuclear translocation of ␤-catenin (59). Canonical Wnt signaling was one of the most differentially regulated pathways in Hdac3 CKO Osx mice calvaria as compared with WT littermates, and several Wnt inhibitors (Axin2, Ror2, Sfrp4) were among the most up-regulated genes (29). Our data suggest that Hdac3 is essential for the mechanism by which Runx2 represses Axin2, as Axin2 levels were strongly transcribed in Hdac3-insufficient mice.
Runx2 haploinsufficient mice mimic features of human cleidocranial dysplasia in that they present with hypoplastic clavicles and delayed cranial suture development with persistent fontanels in the calvaria (17)(18)(19). Runx2 is a stimulated by canonical Wnt/␤-catenin signaling (49), and ␤-catenin signaling is enhanced by Axin2 deficiency (10, 12, 16). Our gene expression data from BMSC suggest that double mutant Axin2 Ϫ/Ϫ :Runx2 ϩ/Ϫ osteoblasts may trend toward expressing higher levels of Runx2 as compared with osteoblasts from Runx2 ϩ/Ϫ single mutants (Fig. 5B). Inhibition of Gsk-3␤ (the kinase that targets ␤-catenin for proteasomal degradation) also increases ␤-catenin signaling (60). Interestingly, mice heterozygous for both Runx2 and Gsk-3␤ had larger clavicles and improved calvarial development as compared with the Runx2 heterozygous single mutant animals (61). However, despite the important role of Gsk-3␤ in canonical Wnt signaling, the rescue of the Runx2 ϩ/Ϫ cleidocranial dysplasia phenotype in these mice was explained by reduction of the kinase activity of Gsk-3␤ toward Runx2 and not because of increases in Wnt signaling (61). Thus, it remains unclear whether increasing ␤-catenin signaling could rescue the cleidocranial dysplasia phenotype of the Runx2 heterozygous animals. Future studies will address whether osteoblasts in the Axin2 Ϫ/Ϫ :Runx2 ϩ/Ϫ mice demonstrate increased ␤-catenin signaling and whether this increase could be sufficient to rescue the cleidocranial dysplasia phenotype of the Runx2 ϩ/Ϫ mice.
In conclusion, a novel relationship exists between Runx2, Axin2, and Hdac3 that regulates osteoblast activity in the cranial sutures. Runx2 binds and represses Axin2 expression in an Hdac3-dependent manner in osteoblasts. As a consequence of this relationship, reducing levels of either Runx2 or Hdac3 increases Axin2 expression. Runx2 is essential for Axin2 deficiency-induced craniosynostosis, as introduction of Runx2 haploinsufficiency into the Axin2 Ϫ/Ϫ background rescues the craniosynostosis phenotype of the Axin2 Ϫ/Ϫ single mutant mice. These data shed light on new mechanisms involved in cranial suture biology and may have further-reaching implications in the fields of skeletal and cancer biology. Additional studies will be required to determine the role of ␤-catenin signaling in this model and to quantify the effects of introducing Axin2 deficiency into the Runx2 ϩ/Ϫ background with respect to measures of the cleidocranial dysplasia phenotype of Runx2 haploinsufficient mice.


extfieldsofstudy: Chemistry Medicine

text:
Whole-Body Vibrations Do Not Elevate the Angiogenic Stimulus when Applied during Resistance Exercise

Knowledge about biological factors involved in exercise-induced angiogenesis is to date still scanty. The present study aimed to investigate the angiogenic stimulus of resistance exercise with and without superimposed whole-body vibrations. Responses to the exercise regimen before and after a 6-week training intervention were investigated in twenty-six healthy male subjects. Serum was collected at the initial and final exercise sessions and circulating levels of matrix metalloproteinases (MMP) -2 and -9, Vascular Endothelial Growth Factor (VEGF) and endostatin were determined via ELISA. Furthermore, we studied the proliferative effect of serum-treated human umbilical vein endothelial cells in vitro via BrdU-incorporation assay. It was found that circulating MMP-2, MMP-9, VEGF and endostatin levels were significantly elevated (P<0.001) from resting levels after both exercise interventions, with higher post-exercise VEGF concentrations in the resistance exercise (RE) group compared to the resistive vibration exercise (RVE) group. Moreover, RE provoked increased endothelial cell proliferation in vitro and higher post-exercise circulating endostatin concentrations after 6 weeks of training. These effects were elusive in the RVE group. The present findings suggest that resistance exercise leads to a transient rise in circulating angiogenic factors and superimposing vibrations to this exercise type might not further trigger a potential signaling of angiogenic stimulation in skeletal muscle.

Introduction
There is growing evidence that physical activity is a potent stimulator of angiogenesis in skeletal and cardiac muscle [1]. Endurance training is thought to increase capillarity in skeletal muscle [2], whereas high resistance training has been shown to decrease capillary density [3], most likely as a result of fibre hypertrophy with insufficient angiogenesis. Knowledge about the exact mechanisms of blood vessel growth is to date still scanty. In the current models of sprouting angiogenesis, capillary formation involves two essential steps, namely (i) degradation of the extracellular matrix (ECM) surrounding the capillary and (ii) activation, migration and proliferation of capillary endothelial cells [4]. ECM breakdown is mediated by a family of zinc-and calciumdependent enzymes, the matrix metalloproteinases (MMP) [5]. The proteases MT1-MMP, MMP-2 and -9 seem to play a crucial role in the formation of new capillaries in skeletal muscle [6] and previous studies reveal that their serum concentrations are significantly elevated after endurance exercise [7]. Furthermore, members of the MMP-family are known to release endostatin by proteolytic cleavage of the C-terminal NC1 domain of Collagen XVIII [8]. To date, the role of endostatin in the angiogenic process is not clear due to its complex signaling functions. As both pro-angiogenic [9] and anti-angiogenic [10] characteristics have been described for endostatin, it has been considered to function as an angiogenic modulator [11]. Endostatin seems to play a crucial role in exercise-induced angiogenesis, as serum concentrations were acutely elevated after endurance exercise [12,13]. However, other studies have reported decreased serum concentrations of endostatin as an adaptation to long-term endurance training [7].
Endothelial cell activation, migration, and proliferation is mediated by Vascular Endothelial Growth Factor (VEGF), a potent endothelial cell mitogen [14]. VEGF has been shown to be activated upon elevated shear stress perturbation [15], muscle stretch [16] and hypoxia [17]. Additionally, VEGF has been reported to be essential for exercise-induced angiogenesis in skeletal muscle [18]. The findings of a previous study evaluating the effects of endurance exercise with and without whole-body vibrations revealed that circulating VEGF was specifically increased in the group where vibrations were superimposed to the exercise stimulus [13].Of note, it has been suggested that the mechanical stimulus of whole-body vibration (WBV) increases shear stress at the walls of blood vessels [19], leads to increases in blood flow velocity after vibration termination [20] and can elicit muscle de-oxygenation [21]. Based on the finding that shear stress and hypoxia are able to induce angiogenesis [4], we hypothesized that the superposition of a vibration stimulus to resistance exercise would add a pro-angiogenic stimulus to the exercise. It would be desirable to find a novel training mode that concurrently increases muscle strength and induces capillary growth to optimize the flux of oxygen and nutrients to the muscle and thus improve muscular performance. In order to investigate the pro-angiogenic stimulus of the exercises, we determined serum concentrations of the angiogenic factors MMP-2, MMP-9, VEGF and endostatin at rest and in response to resistance exercise and resistive vibration exercise. Additionally, we performed in vitro assays to evaluate the proliferative property of exercise-serum treated endothelial cells.

Ethics statement
Twenty-six healthy, recreationally active male subjects (2660.8 years) were included into the study after providing a written informed consent. The study was conducted in compliance with the Declaration of Helsinki following approval by the Ethics Committee of the Northern Rhine medical association (Ä rztekammer Nordrhein) in Düsseldorf (application no. 2010-174).

Study design and subject characteristics
The present EVE study (''molecular and functional Effects of Vibration Exercise'') was conducted in a stratified, randomized two-group parallel design. A detailed description of the exercises and study design has been published elsewhere [22]. Any competitive sports, participation in strength training during the past six months, smoking, diabetes as well as any current medication were considered as exclusion criteria. Subjects were stratified into two matched groups according to their maximum jumping height, forming two groups with comparable neuromuscular fitness [23]. A coin was then tossed to randomly assign the groups to one of the two training interventions: resistance exercise or resistive vibration exercise. The subjects anthropometric data at baseline are given in Table 1, and no statistically significant group difference was found (P.0.11).

Training design
The present study was designed to compare acute and longterm effects of two training interventions: resistance exercise (RE) and resistive vibration exercise (RVE). Participants trained 2-3 times per week for six weeks (completing 16 exercise sessions), with each session lasting 9min. Participants trained with weights on a guided barbell (PTS Dual action Smith, Hoist, U.S.A). The individual training load was set at 80% of their One-Repetition-Maximum (1-RM), which was determined according to the method described by Baechle and Earle [24].
The exercise consisted of squats (with each 2 sec. eccentric and 2 sec. concentric phase) and heel raises (with each 1 sec. eccentric and 1 sec. concentric phase), divided by a 1-min break. Movement rhythm was guided by a metronome. Each exercise session consisted of a warm-up composed of two sets with each 10 squats and 15 heel raises with the unloaded barbell (15 kg) as training weight. The actual exercise was carried out in three sets: first and second sets were composed of 8 squats ( = 32 sec. per set) and 12 calf raises ( = 24 sec. per set) and in the third set, maximum number of repetitions for squats and calf raises were performed. The subjects in the RVE group performed the resistance exercise protocol with simultaneous side-alternating whole-body vibrations (GalileoH Fitness, Novotech, Germany) with a 6 mm peak-to peak displacement, whereas subjects in the RE group trained with the same setting, without superimposed vibrations.
The training followed an incremental training design with regards to weight and vibration frequency. Training weights were increased over time according to the subjects' individual training progressions, as described previously [22]. In brief, the number of squats in the 3 rd set was used as a reference to re-determine the subjects' individual 80% of the 1-RM for the following training, using the method described by Baechle and Earle [24]. Training weights in the RE group increased from 75.261.8 kg during the initial exercise to 130.265.1 kg during the final exercise. Weight increase was significantly smaller the RVE group, which increased from 81.562.1 kg during the initial exercise to 110.264.4 kg during the final exercise. Training weight increase was hampered by training with vibration frequencies above 35 Hz, as discussed in the methodological paper on the training design previously published [22].
Vibration frequencies were increased from 20 Hz in the first week to 40 Hz during the last two weeks with 5-Hz weekly increments. The reason for the increase in vibration frequency was that we aimed to test physiological responses when exercising at 40 Hz side-alternating WBV, which to the best of our knowledge has not been tested in any other study. Pilot testing revealed that resistance exercise with 40 Hz side-alternating WBV is more challenging for people not accustomed to WBV, suggesting that it could potentially elicit greater effects than lower vibration frequencies, but also that one must envision problems when embarking directly on such high a frequency. Thus, in order to prevent problem-related drop-out from the RVE group and thus a study bias, we decided to initially set the vibration frequency to 20 Hz and to gradually increase the vibration frequency to 40 Hz.

Serum Collection
Venous blood samples were collected at the initial and final exercise sessions of the 6-week training intervention as illustrated in Figure 1. On that day, subjects had a standardised breakfast (two wheat bread rolls with butter and jam) two hours before exercise. Blood was collected one hour prior to exercise (Rest) and +2 min, +5 min, +15 min, +35 min and +75 min after exercise through a short catheter into serum monovettes (Sarstedt, Nümbrecht, Germany) from the cephalic vein, allowed to clot for 10 minutes, centrifuged at 3000 rpm at 4uC (Heraeus Multifuge 1S-R, Thermo Scientific, Waltham, MA, USA), distributed into small tubes and immediately frozen at 220uC until analysis.

Cell lines and culture conditions
Human Umbilical Vein Endothelial Cells (HUVEC, #C-12200, PromoCell, Heidelberg, Germany) were cultured at 37uC and 5% CO 2 in basal medium with added growth supplements (Endothelial Cell Growth Medium KIT, #C-22110, PromoCell, Heidelberg, Germany). Prior to incubation with human serum and 5-Bromo-2-Deoxyuridine (BrdU), cells were split into 96-well plates (DetachKit, #C-41210, PromoCell, Heidelberg, Germany) and cultured in starvation medium (i.e. basal medium with only 0.5% Fetal Calf Serum as growth supplement) for 24 hours. BrdU incubation was performed in conditioned medium (i.e. basal medium containing 2% of human serum providing growth and proliferation factors). Sera obtained from pre-and post-exercise (Rest, +2 min and +75 min post) at each initial and final exercise sessions were used for generating the conditioned medium, see Figure 1.

Statistical Analyses
Statistical analyses were performed using STATISTICA 10 for Windows (Statsoft, Tulsa, Oklahoma, USA, 1984-2010). The effect of either resistance exercise (RE) or resistive vibration exercise (RVE) on serum concentrations of the angiogenic factors MMP-2, MMP-9, VEGF and endostatin was determined via repeated measures ANOVA with time (Rest vs.+2 min,+5min,+15 min,+35 min, +75 min after exercise) and training status (initial vs. final exercise session) as factors. BrdU incorporation data were normalised to fold increases from resting levels (i.e. absorption of cells incubated with serum derived +2 min and +75 min after exercise divided by absorption of cells incubated with serum at Rest). A repeated ANOVA was performed with time (+2 min vs.+75 min) and training status (initial vs. final exercise) as factors. Tukey's test was used for post-hoc testing. Values are given as means 6 standard error of means (SEM). Statistical significance level was set at P,0.05.

Resting levels
Resting levels of the circulating angiogenic factors MMP-9, VEGF and endostatin were comparable before and after the 6week training intervention (P.0.19) and there were no significant differences in resting levels between the two groups (P.0.68), as shown in Table 2. Resting levels of MMP-2 measured at the final exercise session differed between groups with the RVE group depicting higher values than the RE group (RVE: 193.068.71 ng/mL vs. RE: 172.068.5 ng/mL, P,0.001), which had not been the case at the initial exercise session (P = 0.37).

Effect of Resistance Exercise upon angiogenic factors
MMP-2, MMP-9, VEGF and endostatin were all significantly increased from resting levels after both resistance exercise and resistive vibration exercise (time effect: P,0.001) and all factors depicted maximum concentrations two minutes after exercise termination. In the following, relative increases from resting levels are given for the maximum concentrations that were measured at the time point +2 min.

MMP-2
Acute effects. In the RE group, MMP-2 levels were increased from resting levels by 862% P = 0.001) two minutes after the initial exercise and decreased by 561% (P = 0.035) at the time point +75 min. In the RVE group, on the contrary, MMP-2 levels were not significantly elevated from resting levels after the initial exercise (P = 0.9), and were decreased by 862% (P = 0.01) at the time point+75 min ( Fig. 2A). There were no significant differences between RE and RVE groups at the initial exercise (P = 0.99).
Long-term effects. In the RE group, there were no significant differences in the time courses when comparing initial and final exercise sessions (P = 0.99) as depicted in Fig. 2B(i). At the final exercise of the RVE group, however, the MMP-2 levels were generally elevated over the time course of the initial exercise (time*intervention effect: P = 0.049), see Figure 2B(ii). Post-Hoc testing revealed that MMP-2 concentrations were significantly higher at the time points +2 min (P = 0.028), +15 min (P = 0.019) and +75 min (P = 0.015) in the RVE group compared to the same time point at the initial exercise. While MMP-2 was not elevated from resting levels in the RVE group after the initial exercise of the 6-week training intervention, MMP-2 concentrations were significantly elevated by 862% (P = 0.02) two minutes after the final exercise. Due to the RVE-specific increases in MMP-2 concentrations, clear group differences were apparent at the final exercise session with the RVE group depicting significantly higher MMP-2 concentrations compared to the RE group at rest and after exercise (RE vs. RVE: P,0.01).

MMP-9
Acute effects. MMP-9 was elevated from resting levels 2-15 min after exercise (time effect: P,0.001). The MMP-9 increase after the initial exercise accounted for 71619% in the RE group and 74616% in the RVE group with no significant differences between groups (RE vs. RVE: initial exercise: P = 0.439; final exercise: P = 0.35), see Fig. 3A.
Long-term effects. There was no effect of the 6-week training intervention upon the acute MMP-9 response in serum (initial vs. final exercise: RE: P = 0.44; RVE: P = 0.98), see Figure 3B.

Endostatin
Acute effects. Serum levels of endostatin were increased from resting levels 2-15 min after both RE and RVE (time effect: P,0.001). After the initial training, endostatin levels were elevated by 1763% in the RE group and by 2264% in the RVE group with no significant differences between groups (P = 0.85), see Figure 4A.
Long-term effects. After the final exercise, endostatin concentrations in the RE group were uniformly greater than concentrations after the initial exercise (time * intervention effect: P,0.001, see Figure 4B(i). This long-term effect was not seen in the RVE group (time * intervention effect: P = 0.991), see Figure 4B(ii).

VEGF
Acute effects. In the RE group, VEGF was elevated from resting levels 2-15 min after the initial exercise (time effect: P,0.001). In the RVE group, the response differed as this group showed elevated VEGF concentrations only at the time point +2 min (time effect: P,0.001). VEGF concentrations were significantly higher in the RE group with a 41616% increase from resting levels compared to the RVE group, which showed a 3367% increase at the time point +2 min (P = 0.014). Significantly higher VEGF concentrations in the RE group compared to the RVE were also detected at the remaining time points 5-75 min after exercise termination (P-values between 0.02 and 0.004), see Figure 5A.
Long-term effects. There were nonsignificant changes in the responses to the exercises after 6 weeks of training, (initial vs. final exercise: RE: P = 0.520; RVE: P = 0.814, see Figure 5B) and VEGF concentrations after the final exercise were also higher in the RE group compared to the RVE group (RE vs. RVE: Pvalues between 0.01 and 0.005).

Endothelial Cell Proliferation
We used the human serum derived at rest and +2 min and +75 min after exercise to test the proliferative effect upon human umbilical vein endothelial cells (HUVEC) in vitro. These time points were suitable as the angiogenic factors measured via ELISA depicted maximum serum concentrations +2 min after exercise termination and concentrations were back at resting levels at the time point+75 min. Absorption data detecting BrdU incorporation were normalized to fold increases from resting levels.
Endothelial cells incubated with serum derived at +2 min after resistance exercise showed an increased proliferation compared to cells incubated with +75 min serum (time effect: P = 0.0171). This effect was not seen in the RVE group (time effect: P = 0.295). EC proliferation did not differ between cells treated with serum derived after the initial or final exercises in neither group (RE: P = 0.94; RVE: P = 0.91) and no significant differences between the groups were found (P = 0.122), see Figure 6.

Discussion
To test our hypothesis that superimposing whole-body vibrations to resistance exercise would add a pro-angiogenic stimulus to the training, we evaluated serum concentrations of angiogenic

MMPs
Degradation of the vascular basement membrane is an initial step in angiogenic sprout formation and allows endothelial cells to migrate into the interstitial matrix in order to form a premature capillary lumen. Matrix metalloproteinases are crucial for extracellular matrix degradation and are thought to be essential for physiological angiogenesis [25]. MMPs have furthermore been implicated in the release and bioavailability of growth factors [26,27] and play a role in initiating endothelial cell migration and proliferation [28]. Our data show elevated serum MMP-2 levels two minutes after resistance exercise. In the group where wholebody vibrations were superimposed to the exercise, MMP-2 was not elevated after an initial exercise bout but showed an adaptation to long-term training; after 6 weeks of RVE, MMP-2 was elevated above resting levels and concentrations were higher (at rest and post-exercise) compared to the group that had trained without vibrations. This might be a compensatory adaptation to the initial lack of MMP-2. Beyond initiating capillary growth, MMP-2 has furthermore been shown to increase the bioavailability of insulin-like growth factor (IGF) through proteolysis of the IGF binding protein [26,27]. If this is the case, the observed increases in circulating levels of MMP-2 after six weeks of RVE might reflect an increased IGF-associated anabolic stimulation.
The presented MMP-9 data showed a prolonged increase compared to MMP-2 and MMP-9 was elevated from resting levels until 15 min after both exercise regimes with no detectable longterm effect. A previous study measuring serum MMP-9 concen- tration pre and post eight weeks of resistance training vs. eight weeks of callisthenic training report an increase in the acute MMP-9 response after eight weeks of training only in the callisthenic group [29]. A study on downhill running showed no acute changes in serum MMP-2 but increased serum MMP-9 levels immediately after exercise [30]. Running step tests increased plasma MMP-2 one hour post-exercise whereas plasma MMP-9 was increased immediately after exercise with decreased resting MMP-9 levels after six months of training [7].
Together with the present findings, these data suggest that MMP-responses to acute and long-term training interventions highly depend on workload, volume and contraction form of the exercise. The exposure to different mechanical stimuli seems to foster extracellular matrix remodeling in divergent ways, revealing a potential role of MMPs in initiating training-specific muscle adaptations. A limitation of the procedure is that the available antibodies do not differentiate between the active and pro-enzyme forms of MMPs and we did not measure serum concentrations of tissue inhibitors of metalloproteinases. Therefore, increased MMP-2 and MMP-9 immunoreactivity does not necessarily indicate an increased enzymatic activity.

Endostatin
Our data show that circulating endostatin was elevated from resting levels 2-15 min after a bout of resistance exercise with no additional effect of superimposed vibrations. Previous studies report prolonged elevations of circulating endostatin compared to the time curves we observed: elevations in plasma from 1 h [31] until 6 h post exercise [12] have been reported after endurance exercise. After 90 min of cycling exercise, Suhr and colleagues [13] found endostatin to be elevated in plasma 0-60 min after exercise termination and superimposing vibrations to this exercise type shortened the elevation from baseline levels to 0 min after exercise, which is an effect of superimposed vibration we did not observe in the present study.
Although we did not see any long-term adaptations in basal endostatin levels, as previously reported for endurance training [7], the response was altered after the 6-week training intervention. Endostatin concentrations in serum were acutely higher after 6 weeks of training and this adaptation was specific for the RE group. Thus, superimposed vibrations seemed to inhibit this biological adaptation to long-term training. Due to endostatin's complex signaling functions, it is not a simple task to interpret the physiological impact of elevated endostatin concentrations after exercise. Initially, endostatin was described as an anti-angiogenic protein [10] capable of inducing apoptotic signals in endothelial cells [32] and to inhibit EC migration, -proliferation and tube formation [33]. Conversely, it was later shown that endostatin has both pro-and anti-angiogenic functions depending on its concentration and the proliferation status of endothelial cells [11]: towards the running opinion, Schmidt and colleagues [11] showed that endostatin concentrations of 50 ng/mL induced EC proliferation and migration with no induction of apoptosis; whereas concentrations of 1000 ng/mL and above had the contrary effect. Based on these data, the endostatin concentrations we reported in the present study (90-140 ng/mL) lie close to the concentrations that were considered as a pro-angiogenic range. Thus, the observed increase in endostatin response after 6 weeks of training (RE only) might reflect a pro-angiogenic long-term training adaptation, which is inhibited by superimposed vibrations.
The acutely elevated endostatin levels seem to have a critical function during exercise. As recently demonstrated by our group, endostatin induces the release of the vasodilator NO in endothelial cells [34]. The acute exercise-dependent endostatin release therefore seems to be essential to activate signaling pathways that result in peripheral vasodilation and consequently improves oxygen delivery to working skeletal muscles to maintain the muscle performance capacity.

VEGF
The process of endothelial cell proliferation is mediated mainly by Vascular Endothelial Growth Factor (VEGF), a potent endothelial cell mitogen [14]. Exercise leads to increases of VEGF protein in muscle tissue [31] and VEGF has shown to be essential for exercise-induced angiogenesis in skeletal muscle [18]. VEGF serum concentrations were shown to be decreased [12,31] or elevated [35] after endurance-type exercise. Our data are to our knowledge the first that reveal acute increases of circulating VEGF immediately after resistance-type exercise. We could show that VEGF was elevated in serum 2-15 minutes after resistance exercise, whereas superposition of vibrations to the exercise shortened this response to only two minutes after exercise and provoked significantly lower VEGF concentrations compared to the group that trained without vibrations. As we did not measure VEGF expression in muscle tissue, this finding gives rise to multiple possible explanations. First, decreased circulating VEGF could indicate that more VEGF is still held and active in the tissue and has not been washed out into the blood. Second, reduced circulating VEGF upon vibration exposure could indicate that whole-body vibrations in some way prevented VEGF secretion or release in muscle tissue, which would indicate that superimposing vibrations would not be beneficial for a potential activation of angiogenic signaling in skeletal muscle. Third, VEGF is produced in many cell types and the increased circulating VEGF might also derive from a systemic exercise effect which is not related to muscle tissue and could indicate enhanced endothelial regenera- tion, which would reflect a beneficial effect of resistance exercise that was inhibited by superimposed vibrations.
In a previous study in our lab, the effect of high-intensity cycling exercise with and without whole-body vibrations was evaluated and this study revealed contrary results considering vibration exposure: plasma VEGF levels were only increased in the group where vibrations were superimposed to the exercise stimulus [13]. As previous studies reveal that WBV increase the shear stress in blood vessels [19], Suhr and colleagues concluded that vibrationinduced increases in shear stress-stimulated VEGF release as described by Milkiewicz and colleagues [15]. This explanation does not seem to be applicable in the present study, as our data reveal the contrary, i.e. reduced VEGF upon vibration exposure. Thus, whole-body vibration stimulation seems to have differential effects according to the mode it is applied. In the case of endurance cycling exercise, superimposed vibrations might be beneficial for promoting angiogenesis (reflected by increases in VEGF), whereas our data reveal that the contrary seems to be the case for resistance exercise. As exercise times in the aforementioned study (90 min) were much longer compared to the present study (9 min), it might well be that the initial effects of the exercises are comparable but the measured VEGF kinetics may differ due to the time shift in the measurements.
It is well known that levels of angiogenic markers differ according to the type of blood product in which they were measured (serum vs. plasma). Previous studies were inconsistent in the type of blood product used and this might contribute to discrepancies between studies.

Endothelial cell proliferation
One limitation of measuring angiogenic markers in serum is that their site of action resides within the muscle tissue itself and we determine merely the 'wash-out' in serum. Consequently, we sought to investigate whether and in which manner elevated serum concentrations would possibly influence endothelial cells in vitro, because this model is well-established to test general defined reactions of endothelial cells in vitro that might reflect in vivo situations.
As all factors showed maximum concentrations +2 min after exercise and were back at resting levels +75 min after exercise, we chose to treat human umbilical vein endothelial cells (HUVEC) with serum derived from these time points. We found that endothelial cells incubated with serum derived +2 min after RE showed increased proliferation compared to cells incubated with serum derived+75 min after exercise. This effect was not seen in the RVE group. VEGF was the only angiogenic factor that showed group-specific differences after exercise (see Figure 5A). VEGF serum concentrations were higher +2 min after RE ([3526104 pg/mL] after initial-and [3696107 pg/mL] after final exercise) compared to +2 min after RVE ([280650 pg/mL] after initial-and [268643 pg/mL] after final exercise), which may be an explanation for the group-specific differences in cell proliferation. The recommended VEGF concentration for HU-VEC culture is 500 pg/mL (Endothelial Cell Growth Medium KIT, #C-22110, PromoCell, Heidelberg, Germany), which lie close to the VEGF concentrations we measured in the RE group. However, there are various additional factors that were not measured in the present study that, however, could have influenced HUVEC proliferation, i.e. basic Fibroblast Growth Factor [36], epidermal growth factor (EGF) or heparin-binding EGF-like growth factor [37].
Thus, our data, with certain limitations, reveal that superimposed whole-body vibrations to resistance exercise leads to decreased endothelial cell proliferation, probably due to decreased release or expression of VEGF. Considering long-term adaptations, we did not find any differences in HUVEC proliferation when comparing initial and final exercise sessions. Despite acutely higher endostatin levels during the final exercise in the RE group and higher MMP-2 concentrations in the RVE group, these effects were not reflected by increased cell proliferation during the final exercise.

Comparison of Time curves
When comparing the time curves of MMP-9 with VEGF and endostatin, it seems that the exercise-induced increase of MMP-9 is paralleled by VEGF and endostatin. First, all factors were increased 2-15 min after exercise and second, all three factors show increased mean concentrations after 6 weeks of training (although only significant for endostatin), see Figure 3B(i), 4B(i) and 5B(i). Conversely, the factor MMP-2 showed different kinetics as it was elevated only for two minutes after exercise and the longterm adaptation that was seen for MMP-2 in the RVE group was specific for MMP-2 and did not affect any of the other factors. In sum, these observations indicate that MMP-9, VEGF and endostatin seem to be interdependent, whereas MMP-2 seems to be differentially regulated. Our data are in line with previous observations in cell culture which showed that MMP's are capable of inducing VEGF release [38]. Moreover, the presented data confirm a previous finding in which the authors described that MMP-9 was more prone to release VEGF compared to MMP-2 in vitro and that that MMP-2 regulation occurred independently of VEGF signaling [28]. The parallel increase of MMP-9 and endostatin confirms that endostatin is proteolytically released by MMP's, as described previously [8] and our data hint to MMP-9 playing a larger part in this release compared to MMP-2, at least after bouts of resistance exercise.
In summary, our data show that RE leads to transient increases in circulating pro-angiogenic markers and furthermore, endothelial cell proliferation in vitro is increased by factors in serum obtained acutely after RE. Superimposing vibrations to resistance exercise decreases post-exercise circulating VEGF concentrations, which supposedly results in reduced endothelial cell proliferation in vitro. Six weeks of RE increased endostatin concentrations acutely after exercise, which is considered as a pro-angiogenic adaptation which was prevented by training with superimposed vibrations. In other words, the presented data suggest that superimposing a vibrations stimulus to resistance exercise might not be beneficial for triggering angiogenic-inducing signaling pathways in skeletal muscle.


extfieldsofstudy: Biology Medicine

s2fieldsofstudy: Biology Medicine Chemistry

text:
Histone Demethylase Retinoblastoma Binding Protein 2 is Overexpressed in Hepatocellular Carcinoma and Negatively Regulated by hsa-miR-212

Background The H3K4 demethylase retinoblastoma binding protein 2 (RBP2) is involved in the pathogenesis of gastric cancer, but its role and regulation in hepatocellular carcinoma (HCC) is unknown. We determined the function of RBP2 and its regulation in HCC in vitro and in human tissues. Methods We analyzed gene expression in 20 specimens each of human HCC and normal liver tissue by quantitative real-time PCR and immunohistochemistry. Proliferation was analyzed by foci formation and senescence by β-galactosidase staining. Promoter activity was detected by luciferase reporter assay. Results The expression of RBP2 was stronger in cancerous than non-cancerous tissues, but that of its binding microRNA, Homo sapiens miR-212 (hsa-miR-212), showed an opposite pattern. SiRNA knockdown of RBP2 significantly upregulated cyclin-dependent kinase inhibitors (CDKIs), with suppression of HCC cell proliferation and induction of senescence. Overexpression of hsa-miR-212 suppressed RBP2 expression, with inhibited cell proliferation and induced cellular senescence, which coincided with upregulated CDKIs; with low hsa-miR-212 expression, CDKIs were downregulated in HCC tissue. Inhibition of hsa-miR-212 expression upregulated RBP2 expression. Luciferase reporter assay detected the direct binding of hsa-miR-212 to the RBP2 3′ UTR. Conclusions RBP2 is overexpressed in HCC and negatively regulated by hsa-miR-212. The hsa-miR-212–RBP2–CDKI pathway may be important in the pathogenesis of HCC.

Introduction
Hepatocellular carcinoma (HCC) is one of the most common tumors in China [1][2][3]. Its early diagnosis is difficult. HCC can be diagnosed effectively in some cases with serological assessment [4][5][6], but the molecular mechanism of cell proliferation and effective biological markers for diagnosis are unclear.
We found RBP2 overexpressed in gastric cancer, and its inhibition could trigger cancer-cell senescence [16]. We also found RBP2 overexpressed in HCC. RBP2 was significantly underexpressed in normal liver tissue but strongly expressed in cancerous liver tissues, so RBP2 may take part in the development of HCC. Furthermore, siRNA inhibition of RBP2 expression could decrease the proliferation and induce senescence of high-differentiated HepG-2 cells and low-differentiated SMMC-7721 cells, 2 representative HCC cell lines, but the mechanism of RBP2 overexpression has not been elucidated.
MicroRNAs (miRNAs), non-coding 20-to 23-nt RNAs, play a key role in gene expression in different kinds of cancer [17][18]. MiRNAs can bind to the seed sequence of the 39 UTR of target genes and negatively regulate their expression in cell proliferation, differentiation, senescence, and apoptosis, for example. MiRNAs also participate in cancer metastasis and cancerous angiogenesis by regulating specific genes. Homo sapiens miRNA-212 (hsa-miR-212) has 2 sites binding in the 39 UTR of RBP2. It is downregulated and suppresses methyl-CpG-binding protein MeCP2 in human gastric cancer [19]. Downregulation of hsa-miR-212 increasing the expression of HB-EGF may be involved in cetuximab resistance in head and neck squamous cell carcinoma [20].
In this research, we investigated the involvement of RBP2 in HCC and the miRNA implicated in its regulation in human HCC. RBP2 inhibition and hsa-miR-212 overexpression may lead to cell proliferation arrest and cellular senescence by depressing cyclindependent kinase inhibitors (CDKIs; p16 ink4a , p21 CIP2 and p27 kip1 ). RBP2 and hsa-miR-212 may be biomarkers for diagnosis of HCC and therapeutic targets in its treatment.

Materials and Methods
The ethics committee of Shandong University School of Medicine approved this study and all participants provided written informed consent.

Clinical Samples
We obtained 20 samples of HCC from patients at Bengbu Medical University, AnHui Province, China, immediately after surgery and stored them in formalin. The diagnosis of HCC was confirmed by histological examination; no patient had undergone chemotherapy or radiotherapy before surgery. The general characteristics of patients are in Table S1. We excluded patients with a history of alcohol abuse.

Other Materials and Methods
Immunohistochemistry, cell culture [21][22][23] and siRNA interference, RNA extraction, RT-PCR and real-time PCR (primers are in Table 1), plasmid transfection and luciferase reporter gene assay (primers and methods are in Table 2), protein extraction and western blot assay, clonal formation assay and senescence-associated b-galactosidase (SA-b-Gal) staining are described in Data S1.

Statistical Analysis
Quantitative data are expressed as mean6SD or SEM. Student's t test was used to analyze the differences between groups. P,0.05 was considered statistically significant.

Results
RBP2 is overexpressed and hsa-miR-212 underexpressed in human HCC.
Immunohistochemical staining revealed higher RBP2 protein expression in HCC than normal liver tissue ( Figure 1A), and the mRNA level of RBP2 was higher in HCC than normal tissue (P,0.001; Figure 1C) [24]. RBP2 was highly expressed in most HCC tissues (75%), whereas most normal tissues (70%) showed low RBP2 expression ( Figure 1B). Overexpression of RBP2 at the mRNA and protein levels in HCC tissue suggested that RBP2 might participate in the initiation and development of HCC.
Hsa-miR-212, the predicted miRNA that could bind to the 39 UTR of RBP2 ( Figure 1D), may regulate its expression in these clinical samples. Hsa-miR-212 level was significantly lower in HCC than normal tissue (P,0.001; Figure 1E), which suggested a negative regulation between RBP2 and hsa-miR-212 in HCC. RBP2 and miR212 levels were negatively correlated (R 2 = 0.7941, Figure 1F).

Inhibition of RBP2 Expression in HCC Cells Repressed Cell Proliferation and Induced Cell Senescence by Upregulating CDKIs
Next, we investigated the mechanism by which RBP2 participates in hepatic oncogenesis in 2 hepatic cancer cell lines, HepG-2 and SMMC-7721 cells. The mRNA level of RBP2 was decreased with RBP2 siRNA knockdown, by about 80% and 72% in HepG-2 and SMMC-7721 cells, respectively (Figure 2A; Figure S1A), and RBP2 protein level was decreased ( Figure 2A). Table 1. PCR primer sequences used in the study. At the same time, H3K4me2 and H3K4me3 expression was significantly increased with RBP2 depletion ( Figure S2C). The expression of known targets of RBP2, namely, the CDKIs p16 ink4a , p21 CIP2 and p27 kip1 , was markedly increased with RBP2 siRNA knockdown ( Figure 2B; Figure S2A). As a result, the proliferation of HCC cells was sharply restricted ( Figure 2C).

Genes Primers
Cell senescence was induced with RBP2 siRNA knockdown for 72 hr in cancer cells ( Figure 2D). Therefore, CDKI expression could be regulated by RBP2, and knocking down RBP2 expression could repress cell proliferation and induce cell senescence.

Overexpression of RBP2 Blocked CDKI Expression in vitro
RBP2 mRNA and protein expression was induced with an overexpression plasmid ( Figure 3A; Figure S1B) and the expression of the CDKIs p16 ink4a , p21 CIP2 and p27 kip1 was downregulated ( Figure 3B and C; Figure S2B).
Expression of the CDKIs p21 CIP2 and p27 kip1 was Directly Regulated by RBP2 To determine whether p21 CIP2 and p27 kip1 were directly regulated by RBP2, siRNA knockdown of RBP2 expression greatly increased luciferase activity of p21 CIP2 and slightly but significantly increased activity of p27 kip1 in HepG-2 cells ( Figure 3D); ChIP assay confirmed the results ( Figure 3E). Therefore, RBP2 could directly bind to p21 CIP2 and p27 kip1 promoters and suppress their expression.
HepG-2 cells were injected into nude mice and tumors were formed ( Figure S1F). P21 CIP2 and p27 kip1 co-located with RBP2 on immunohistochemical staining of serial tissue sections of tumors: with RBP2 overexpression, the expression of p21 CIP2 and p27 kip1 was relatively low ( Figure 3F). Thus, RBP2 regulated the expression of CDKIs and affected cell proliferation, thereby contributing to the development of HCC in humans.

RBP2 was Negatively Regulated by hsa-miR-212
We confirmed that RBP2 was overexpressed in HCC and investigated the mechanism of the increased RBP2 expression. We found that RBP2 is a target of hsa-miR-212, and hsa-miR-212 could bind with 2 regions of the 39 UTR of RBP2 ( Figure 1D). With hsa-miR-212 overexpression in HepG-2 and SMMC-7721 cells, hsa-miR-212 expression was significantly increased, which confirmed the success of hsa-miR-212 overexpression ( Figure 4A). RBP2 expression was markedly repressed with hsa-miR-212 overexpression ( Figure 4B and C; Figure S1C). The expression of p16 ink4a and p27 kip1 , targets of RBP2, was increased in the 2 cell lines ( Figure 4D), which agreed with previous findings. Accordingly, cell proliferation was repressed and cell senescence induced with hsa-miR-212 overexpression ( Figure 4E and F). To further confirm that RBP2 was negatively regulated by hsa-miR-212, transfection of hsa-miR-212 inhibitor plasmid in the 2 cell lines decreased the expression of hsa-miR-212 ( Figure 5A) and increased that of RBP2 ( Figure 5A and B; Figure S1D). Accordingly, the expression of p16 ink4a and p27 kip1 was decreased with hsa-miR-212 inhibition ( Figure 5C).
To ascertain whether hsa-miR-212 directly regulated the expression of RBP2, use of an RBP2 39-UTR reporter vector significantly decreased luciferase activity with hsa-miR-212 overexpression in SMMC-7721 cells. However, co-transfection of hsa-miR-212 with the 2 mutants of the RBP2 39 UTR reporter vector (mutation of the 2 binding sites of RBP2, Table 2) reversed the luciferase activity ( Figure 5D). Thus, RBP2 was a direct target of hsa-miR-212.

Discussion
RBP2 is involved in the pathogenesis of gastric cancer, but its role and regulation in HCC was unknown. We determined the function of RBP2 and its regulation in HCC in vivo and in vitro. The expression of RBP2 was stronger in cancerous than noncancerous human tissues, but that of its binding miRNA, hsa-miR-212, was reversed. SiRNA knockdown of RBP2 significantly upregulated CDKIs, with suppression of HCC cell proliferation and induction of cell senescence. Overexpression of hsa-miR-212 had a similar effect. Conversely, inhibition of hsa-miR-212 expression upregulated RBP2 expression. We found that RBP2 is overexpressed in HCC and negatively regulated by hsa-miR-212. The hsa-miR-212-RBP2-CDKI pathway may be important in the pathogenesis of HCC.
Epigenetic modulation participates in various biological processes by affecting chromatin stability and transcriptional repression or activation of target genes. Epigenetic molecules may play Table 2. Primer sequences for construction of wild type (WT) and mutants (MUT1 and 2) of the 39 UTR of retinoblastoma binding protein 2 (RBP2) by the overlap extension PCR method.

Category Primers
Sequences      an important role in many types of tumorigenesis [25]. An important histone demethylase family, the JARID1 protein family, has been investigated widely in the biological process of higher organisms [18]. JARID1B/RBP2-H1/KDM5B was found to be a transcriptional regulator with tumor-suppressive potential in melamoma cells [26] and was used as a biomarker for a small subpopulation of slow-cycling melanoma cells [27]. The von Hippel-Lindau tumor-suppressor protein regulates gene expression and tumor growth via the histone demethylase JARID1C/ KDM5C/SMCX in clear-cell renal cell carcinoma [28]. The expression pattern of JARID1D/KDM5D/SMCY significantly differed in meningiomas of male and female patients and contributes to the female predominance of this tumor [29]. RBP2, a member of the JARID family protein, also named JARID1A or KDM5A, has histone demethylase activity that could demethylate H3K4me3 and H3K4me2 significantly [8,12]. Thus, RBP2 may affect the stability of chromatin through histone modulation. We first investigated the expression of RBP2 in human HCC and found the same expression as in gastric cancer. Recently, upregulation of RBP2/KDM5A expression was seen as characteristic of a drug-tolerant cancer cell subpopulation, cancer stem cells [15], which highlights the role of RBP2 in tumors and suggests its role in the initiation and development of HCC, but its mechanism in cancer was not eluciated.
MiRNAs affect tumorigenesis [30][31][32]. They can negatively regulate oncogenes and play a role as tumour suppressor genes but can also target tumour-suppressor genes and play a role as oncogenes. MiR-21 is overexpressed in most types of cancer [33][34] and Let-7 has an exclusively strong effect in lung cancer development [35]. Here, we found that RBP2 is a direct target of hsa-miR-212, and overexpressed RBP2 in HCC tissues may result from downregulated hsa-miR-212 expression. Hsa-miR-212 has been widely investigated [36][37][38][39]. We found that Hsa-miR-212 directly binds to the 39 UTR of RBP2 and suppressed its expression. The suppressed RBP2 expression led to inhibited proliferation and induced senescence of hepatocellular cells by upregulating CDKIs. So, the low expression of hsa-miR-212 in HCC may result in overexpression of RBP2 and downregulation of RBP2 targets such as CDKIs, accompanied by cell proliferation and blocking cellular senescence in HCC, which contributes to its pathogenesis. Here, we provide more evidence of epigenetic molecule regulation by miRNAs.
However, the mechanism by which hsa-miR-212 is downregulated in HCC is not known. Infection with hepatitis B virus (HBV) may play a role in this process because HCC can result from HBV infection. As well, downregulation of hsa-miR-212 may be due to hypermethylation of DNA in the hsa-miR-212 promoter because downregulation of hsa-miR-212 expression is through DNA hypermethylation in human gastric cancer [40]. Recently, miR-212 expression was found to be deregulated in lung cancer [41]. However, other mechanisms may lead to the suppression of hsa-miR-212 expression in HCC. Also, downregulated hsa-miR-212 expression may not be the only mechanism leading to the overexpression of RBP2. We found that depletion of one miRNA, hsa-miR-212, antagonized the suppression of RBP2, which led to RBP2 overexpression in HCC, but further investigation is needed to elucidate the reason for the overexpressed RBP2 in HCC. Induced epigenetic molecules could also regulate the expression of miRNAs [42][43][44], but we found no evidence of induced RBP2 affecting hsa-miR-212 expression because siRNA knockdown of RBP2 did not change the expression of hsa-miR-212 ( Figure  S1E).
In conclusion, our findings suggest that overexpressed RBP2 in HCC resulted from downregulated hsa-miR-212 expression and that overexpressed RBP2 could promote cell proliferation and suppress cell senescence by downregulating p16 ink4a , p21 CIP2 and p27 kip1 . The hsa-miR-212-RBP2-CDKI pathway in HCC may contribute to its initiation and development. Low expression of hsa-miR-212 and overexpression of RBP2 may be prognostic markers for HCC. As well, RBP2 and hsa-miR-212 expression may be a therapeutic target for HCC.

Supporting Information
Data S1 Supporting Materials and Methods.