Original Article

microRNA-103a functions as a mechno -sensitive microRNA to inhibit bone formation through targeting Runx2 † Bin Zuo,1*JunFeng Zhu,1*Jiao Li,2ChuanDong Wang,2XiaoYing Zhao,2GuiQuan Cai,1 Zheng Li,1Jianping Peng,1Peng Wang,1Chao Shen,1Yan Huang, 2Jiake Xu3,XiaoLing Zhang,2‡XiaoDong Chen1‡ 1

Department of Orthopedic Surgery, Xinhua Hospital, Shanghai JiaoTong University School of

Medicine (SJTUSM), Shanghai, China 2 The Key Laboratory of Stem Cell Biology, Institute of Health Sciences, Shanghai Institutes for Biological Sciences (SIBS), Chinese Academy of Sciences (CAS) & Shanghai JiaoTong University School of Medicine (SJTUSM), Shanghai, China 3

School of Pathology and Laboratory Medicine, The University of Western Australia Perth, WA 6009,

Australia *

These authors contributed equally

‡

Author for correspondence ([email protected] and [email protected])

†

This article has been accepted for publication and undergone full peer review but has not been through the copyediting, typesetting, pagination and proofreading process, which may lead to differences between this version and the Version of Record. Please cite this article as doi: [10.1002/jbmr.2352]

Additional Supporting Information may be found in the online version of this article. Initial Date Submitted January 24, 2014; Date Revision Submitted August 4, 2014; Date Final Disposition Set August 20, 2014

Journal of Bone and Mineral Research © 2014 American Society for Bone and Mineral Research DOI 10.1002/jbmr.2352

Abstract Emerging evidence indicates that microRNAs (miRNAs) play essential roles in regulating osteoblastogenesis and bone formation. However, the role of miRNA in osteoblast mechanotransduction remains to be defined. In this study, we aimed to investigate if miRNAs regulate mechanical stimulation-triggered osteoblast differentiation and bone formation through modulation of Runx2, the master transcription factor for osteogenesis. We first investigated the role of mechanical loading both in a mouse model and in an osteoblasts culture system and the outcomes clearly demonstrated that mechanical stimuli can regulate osteogenesis and bone formation both in vivo and in vitro. Using bioinformatic analyses and subsequently confirmed by quantitative Real-Time PCR (qRT-PCR), we found that multiple miRNAs were responding to in vitro mechanical stimulation that potentially target Runx2. Among which miR-103a was fully characterized. miR-103a and its host gene PANK3 were both downregulated during cyclic mechanical stretch-induced (CMS) osteoblast differentiation, whereas Runx2 protein expression was upregulated. Overexpression of miR-103a significantly decreased and inhibition of miR -103a increased Runx2 protein level, suggesting that miR-103a acts as an endogenous attenuator of Runx2 in osteoblasts. Mutation of putative miR -103a binding sites in Runx2 mRNA abolishes miR-103a-mediated repression of the Runx2 3'UTR luciferase reporter activity, suggesting that miR -103a binds to Runx2 3'UTR. Osteoblast marker genes profiling and osteogenic phenotype assays demonstrated that miR -103a negatively correlates with CMS-induced osteogenesis. Further, the perturbation of miR-103a also has a significant effect on osteoblast activity and matrix mineralization. More importantly, we found an inhibitory role of miR-103a in regulating bone formation in hindlimb unloading mice, and pretreatment with antagomir-103a partly rescued the osteoporosis caused by mechanical unloading. Taken together, our data suggest that miR-103a is the first identified mechano-sensitive miRNA that regulates osteoblast differentiation via directly targeting Runx2, and therapeutic inhibition of miR-103a may be an efficient anabolic strategy for skeletal disorders caused by pathological mechanical loading. KEY WORDS: CYCLIC MECHANICAL STRETCH; MICRORNAS; RUNX2; OSTEOBLAST; MECHANOTRANSDUCTION

Introduction Mechanical loading has been previously reported to play an essential role in modulating bone remodeling and homeostasis through mechanosignal transduction pathways. (1-4) Bone is a specifically designed sensory organ consisting of osteocytes and osteoblasts to respond to and adapt to changes in mechanical loads.(5-8) Osteocytes contribute to bone homeostasis by sensing and converting mechanical stress to biological signals, hence, the major effect cell in bone turnover (osteoblasts and osteoclasts) are regulated locally on the bone surface. (1,6,9-10) Osteoblast differentiation is triggered by mechanical stimulation, which induces the secretion of hormones and growth factors, thus affecting the differentiation and proliferation potential of osteoblasts.(11-13) Clinically, decreases in mechanical loading (underuse) owing to extended bed rest (such as long-duration be d rest patients) or exposure to microgravity (such as astronauts) may result in significant bone loss at weight bearing bones and subsequently rapid progression of osteoporosis,(14 -18) whereas increases in mechanical loading (overloading) may lead to stress fractures or fatigue fracture which predominantly occurring in athletes. (19-21) microRNAs (miRNAs) are a family of short, single-stranded noncoding RNA molecules involved in numerous biological processes. Functioning at the post-transcriptional level, miRNAs repress gene expression via degradation or translational inhibition of their target mRNAs by binding to the 3'-untranslated region (3'UTR) of mRNA.(22-25) It is estimated that miR NAs regulate ~30% of human protein-coding genes, demonstrating the essential role of miRNAs in controlling gene expression. (26-27) Multiple miRNAs have been identified to regulate the complex osteoblast-specific marker genes expression and osteogenesis in vitro .(28-32) However, the effects of mechanical stimuli on miRNAs expression and, if any, their functional roles in mechanotransduction in osteoblasts have not been well characterized and are, therefore, particularly interesting to be elucidated . In this study we screened for mechano-sensitive miRNAs in CMS-induced osteoblast differentiation and identified that miR-103a was negatively correlated with osteoblast differentiation and bone formation in response to cyclic mechanical loading.

Specifically, miR-103a participates in the inhibition of osteoblast differentiation and bone formation by directly targeting the 3'UTR of Runx2, the master regulator of osteoblast differentiation. (33-34) Our findings further demonstrate that therapeutic inhibition of miR -103a may partly rescue the osteoporosis caused by mechanical unloading in hindlimb unloading mice. (35 -36) This study may provide a novel mechanism and potential therapeutic target for skeletal disorders caused by pathological mechanical loading.

Materials and Methods Hindlimb-unloading mice 6-month-old male C57BL/6J mice were purchased from Shanghai SLAC Laboratory Animal Co. Ltd, and were individually caged under standard conditions (12 hours light/12 hours dark cycle, 21°C controlled temperature). The animals were suspended from the hindlimb for a period of 28 d. The hindlimb suspension procedure described by Morey-Holton and Globus was adopted for this study. (35-36) Briefly, a strip of medical adhesive tape (15 cm × 0.5 cm) was applied along the proximal one -third of the tail, which was suspended by passing the tape through a swivel that was attached to a metal bar on the top of the cage. This allowed the forelimbs to have contact with the grid floor and allowed the animals to move around the cage for free access to food and water. The suspension height was adjusted to prevent the hindlimbs from touching any supporting surface while maintaining a suspension angle of approximately 30°. The animal's overall appearance, drinking and eating habits, and tail were monitored three times per day. The distal tip of the tail was examined to ensure that the procedure did not occlude blood flow to the tail (i.e., the tail remained pink). After euthanasia, bilateral femurs and tibiae were dissected and processed for microCT examination, bone histomorphometric analysis and quantitative Real-time PCR (qRT-PCR) analysis. All the experimental procedures were approved by the Committees of Animal Ethics and Experimental Safety of Chinese Academy of Sciences.

Cell culture , Transfection The hFOB 1.19 cell line was purchased from the Cell Bank of the Chinese Academy of Sciences (Shanghai, China).(37-39) The hFOB 1.19 cell line was maintained in Dulbecco’s Modi?ed Eagle’s Medium (DMEM, Hyclone) with 15% FBS (Gibco) and 1% penicillin and streptomycin (Hyclone) and 0.3 mg/mL G418 (Sigma). The cells were maintained at 34.5°C culture conditions of 5% CO2 and 95% humidity and were not used beyond passage 10. For the experiments, confluent cells were removed using 0.25% trypsin containing 10 mM EDTA (Hyclone), resuspended in antibiotic-free growth medium and plated onto six-well plates at a density of 2.0 × 105 cells per well (if not mentioned). To induce osteoblastic mineralization, hFOB 1.19 cells were seeded in 6-well plates with osteogenic medium containing 100 nM dexamethasone, 50 µM of ascorbic acid and 10 mM ß-glycerophosphate (Sigma). hBMSCs were isolated and expanded as reported previously with some modifications.(40) Human bone marrow aspirates were obtained from healthy male donors (A-C) (A, 40 years old; B, 34 years old; C, 25 years old) during routine orthopedic surgical procedures. Institutional Review Board approval was obtained. hBMSCs was maintained in a -Minimum Essential Medium (a -MEM, Hyclone) with 10% FBS (Gibco) and 1% penicillin and streptomycin (Hyclone). The cells were maintained at 37°C culture conditions of 5% CO 2 and 95% humidity and were not used beyond passage 5. For transfection of miRNA or siRNA oligos, the medium was transfected by LipofectamineTM2000 (Invitrogen, USA) was used according to the Manufacturer's Instruction. mimic -103a or inhibitor-103a (Guangzhou Ribobio Co. , LTD) was transfected at the concentration of 200 nM, and Runx2 siRNA was transfected at the concentration of 50 nM. agomir -103a or antagomir-103a (Guangzhou RiboBio Co. , LTD) was transfected at the concentration of 200 µM.

Cyclic mechanical stretch application hFOB 1.19 cells were seeded on six-well BioFlexTM culture plates (The culture plates w ith flexible silicone rubber membranes coated with collagen type I, Flexce ll

International Corporation, Hillsborough, USA). The cells were cultured for 48~72 h to reach 90% con?uency, at which time the growth medium was replaced. The cyclic mechanical stretch at 0.5 Hz sinusoidal curve at 8% elongation was applied (8% elongation, 80000 µe, Sin, 0.5 Hz, CMS) using an FX-5000TTM Flexercell Tension PlusTM unit (Flexcell International Corporation, Hillsborough, USA). The cells were incubated in a humidi?ed atmosphere at 34.5°C and 5% CO2 while stretching. Cells were harvested immediately when CMS stimulation finished.

miRNA extraction The total RNA from the collected cells was extracted using TRIzol reagent (Invitrogen, USA) according to the manufacturer's instructions. The total RNA was collected for real-time PCR (qRT-PCR) analysis.

Alkaline phosphatase staining ALP presence of the cell layers was assessed as follows. The cultured cells were rinsed with PBS three times and ?xed with 4% paraformaldehyde for 10 min at 48°C. The ? xed cells were soaked in 0.1% naphthol AS-MX phosphate (Sigma Aldrich, St. Louis,

MO)

and

0.1%

fas t

red

violet

LB

salt

(Sigma)

in

56

mM

2-amino-2-methyll,3-propanediol (pH 9.9, Sigma) for 10 min at room temperature, washed with PBS, and were then observed under an digital camera.

Assay for alkaline phosphatase activity hFOB1.19 cells were rinsed two times with ice-cold PBS, scrapped from the dishes and suspended in ddH 2O. This was followed by three cycles of freezing and thawing. ALP activity was determined at 405 nm using p-nitrophenyl phosphate (pNPP) (Sigma Aldrich) as the substrate. A 50 ml of sa mple was mixed with 50 ml of pNPP (1 mg/ml) in 1 M diethanolamine buffer containing 0.5 mM MgCl2 (pH 9.8) and incubated at 37°C for 15 mins on a bench shaker. The reaction was stopped by the addition of 200 ml of 2 M NaOH per 200 µl of reaction mixture. Total protein content was determined by the BCA method with protein assay kit (PIERCE, Rockford, IL). ALP activity was calculated as nmol p-nitrophenol per minute per mg protein, and

presented as fold changes over the non-loading group at the respective time points. All experiments were conducted in triplicate.

Cytoskeleton staining The F-actin cytoskeleton in hFOB1.19 cells was checked by staining. hFOB1.19 cells were plated at the density of 2 × 103 cells/cm2 in 2 ml of medium on six-well flexible silicone rubber BioFlexTM plates. After cells attaching to the plates, mechanical tension was applied. After being subjected to mechanical stretching for 3 days, cells were fixed with 4% paraformaldehyde for 30 min, washed with PBS containing 0.05% Tween-20 twice, then permeabilized with 0.1% Triton X-100 for 5 min, and blocked with 1% BSA for 30 min. Phalloidin-FITC at 1:500 (green) (catalog number: P5282, Sigma ) was used to incubate with cells for 1 h at room temperature. Nuclei were counterstained with Hochest 33258 (blue) (catalog number: H1398, Life). Cells were visualized with a confocal microscope (LEICA TCSSP5, Wetzlar, Germany).

Alizarin red staining Cells were fixed in 70% ice-cold ethanol for 1 h and rinsed with double -distilled H2O (ddH2O). Cells were stained with 40 mM Alizarin red S (Sigma), pH 4.0, for 15 min with gentle agitation. Cells were rinsed five times with ddH2O and then rinsed for 15 min with 1× PBS while gently agitating.

Cell proliferation assay For the cell proliferation assay, hFOB1. 19 cells were seeded on six -well ?exible silicone rubber BioFlexTM plates at a density of 1.0 × 104 cells per well for approximately 12 h at 34.5°C and 5% CO2 before proceeding with the assay. Add 1/10th volume of AlamarBlue reagent (Invitrogen) was added directly into the culture medium at time point 0, 24, 48 and 72 h respectively followed by 8% CMS treatment at 34.5°C and 5% CO2. After 4-hour -incubation in the dark, 100ml medium sample was transfered into 96-well plate (Corning, Corning, NY) and measured the

absorbance at 570 and 590 nm using a Multiscan UV visible spectrophotometer (Safire2; TECAN, Mannedorf, Switzerland). Non-seeded BioFlexTM plate with the same medium was used as blanks. Cells at baseline prior to CMS loading served as control. Record results using fluorescence or absorbance according to the manufacturer's instructions.

Runx2 3' UTR cloning and luciferase Assay Runx2 mRNA 3'UTR containing the miR-103a -binding sequences for the human Runx2 gene (gene ID 860) were amplified by PCR from human genomic DNA. The primer sequences used in this study were as follows : RUNX2-3UTR-F : 5' CCGCTCGAGAATTCCTCAGCAGTGGC

3';

RUNX2-3UTR-R

GAATGCGGCCGCTAACAAAACCAAAAAAGCCATTTTATTG

3' .

: The

5' PCR

product was then subcloned into the XhoI, NotI site downstream of the stop codon in the pmiR -RB-REPORT TM

empty Vector (Guangzhou RiboBio Co. , LTD).

Binding-region mutations were achieved using a QuikChange Site-Directed Mutagenesis Kit (Stratagene) following the manufacturer’s instructions. Transient transfection of hFOB1.19 cells was carried out in six -well plates with Lipofectamine 2000 (Invitrogen) following the manufacturer’s instruction. The cells were co-transfected with 200 ng of the luciferase constructs and 50 ng of the pRL-TK (Promega, USA) Renilla luciferase plasmid, and luciferase assays were performed with the dual-luciferase reporter assay system (Promega) according to the manufacturer’s instructions. Luminescent signals were quantified by luminometer (Glomax, Promega), and each value from the Renilla luciferase construct was normalized by Firefly luciferase.

RNA isolation and real-time PCR Total RNA from bone tissues or cells was extracted with TRIzol Reagent (Invitrogen) according to the manufacturer's instructions. First-strand cDNA was synthes ized from 1 µg of total RNA by incubating for 1 h at 42°C with Superscript III reverse transcriptase (Invitrogen, Mulgrave, Australia) following oligo (dT) priming.

After reverse transcription reaction, qRT-PCR was performed by LightCycler480 system (Roche, Mannheim, Germany) using SYBR Premix Ex TaqTM (Takara, Dalian, China) according to the manufacturer’s instructions. All amplifications were normalized by GAPDH. Data were analyzed using the comparison Ct (2-? ? Ct) method and expressed as fold change compared to respective control. Each sample was analyzed in triplicate. The primer sequences used in this study were as follows: GAPDH:

forward,

5`-CCTCTGACTTCAACAGCGAC-3`;

5`-TCCTCTTGTGCTCTTGCTGG-3`;

Col1a1:

5`-CAGCCGCTTCACCTACAGC-3`; 5`-TTTTGTATTCAATCACTGTCTTGCC-3`;

reverse, forward, reverse,

Runx2:

forward,

5`-GCCTTCAAGGTGGTAGCCC-3`; reverse, 5`-CGTTACCCGCCATGACAGTA -3`;

OCN:

forward,

5`-GAAGCCCAGCGGTGCA-3`;

5`-CACTACCTCGCTGCCCTCC-3`;

ALP:

5`-GAGTCGGACGTGTACCGGA-3`; 5`-TGCCACTCCCACATTTGTCAC-3`;

forward, reverse,

OPG:

5`-CCTCTCATCAGCTGTTGTGTG-3`; 5`-TATCTCAAGGTAGCGCCCTTC-3`;

reverse,

forward, reverse,

RANKL:

forward,

5`-CACTATTAATGCCACCGAC-3`; reverse, 5`-GGGTATGAGAACTTGGGATT-3`; PANK3:

forward,

5`-TTTTGGCCGAAGAGGGAACTT-3`;

5`-TAGCACCGTCTGCAATGTTGA-3`; 5`-TTGACTCAGTCGGATTCAATGG-3`;

PANK2:

reverse, forward, reverse,

5`-CAGAAGCAGAGGATACGGATTTT -3`;

Western blot analysis For Western blot analysis, cells were lysed in lysis buffer (50 mM Tris-HCl, pH 7. 4, 150 mM NaCl, 0.1% SDS, 1% Nonidet P-40, 1 mM PMSF and protease inhibitor cocktail (10 mg/mL leupeptin, 10 mg/mL pepstatin A, and 10 mg/mL aprotinin) on ice for 30 min. Protein fractions were collected by centrifugation at 15,000g at 4 °C for 10 min and then subjected to 10% SDS-PAGE and transferred to polyvinylidene difluoride (PVDF) membranes. The membranes were blocked with 5% BSA and

incubated with specific antibodies overnight at 4°C. A horseradish peroxidase–labeled secondary antibody was added and visualized using the enhanced chemiluminescence detection system (Millipore, Billerica, MA) as recommended by the manufacturer. Immunoreactive bands were quantitatively analyzed in triplicate by normalizing the band intensities to GAPDH on scanned films with Alpha Image software. We used primary antibodies recognizing human Runx2 Rabbit mAb (1:1000, Cell Signaling Technology, Inc , No. 8486), active-beta-catenin Rabbit mAb, phospho-p44/42 MAPK Erk1/2 Rabbit mAb, p44/42 MAPK Erk1/2 Rabbit mAb, GAPDH Rabbit mAb (1:1000, all purchased from Cell Signaling Technology, Inc) to examine the concentrations of proteins in the lysates, respectively.

Bone histomorphometric analyses We measured the structure of distal femurs with a SCANCO Medical µCT 40 scanner to produce the images and analyzed them with SCANCO evaluation software for segmentation, three-dimensional morphometric analysis, density and distance parameters (SCANCO Medical AG, Switzerland). Three-dimensional structural parameters analyzed included: TV (total tissue volume; contains both trabecular and cortical bone), BV/TV (trabecular bone volume per tissue volume), Tb.Th (trabecular thick-ness), Tb.Sp (trabecular separation), SMI (structure model index), Conn.Dn (connectivity density). For assessment of bone formation, we injected green fluorescent calcein (Sigma; 5 mg per kg body weight) into the mice on days 7 and 2 before euthanasia. Trabecular sections were subjected to tartrate-resistant acid phosphatase (TRAP) staining to analyze static parameters or left unstained for collection of fluorochrome -based data. Bone static histomorphometric analyses for Ob.S/BS, Oc.S/BS, Ob.N/B.Pm and Oc.N/B.Pm, as well as bone dynamic histomorphometric analyses for MAR and BFR/BS, were performed using professional image analysis software (Image J, NIH, USA) under fluorescence microscopy (Leica image analysis system, Q500MC). The bone histomorphometric parameters were calculated and expressed according to the standardized nomenclature for bone histomorphometry.

Therapeutic inhibition of miR-103a in hindlimb-unloading mice Six-month-old

C57BL/6J

mice

received

daily

tail-vein

injections

of

Antagomir-103a (HU + Antagomir -103a group, 80 mg/kg body weight in 0.2 ml per injection), PBS (HU + PBS group, 0.2ml) for three consecutive days or no treatment (HU group) before hindlimb-unloading suspension. The mice were then subjected to hindlimb unloading through tail suspension for 28d and were euthanized. Tissues were harvested, measurements of miR -103a levels in tissues were performed. To maintain the effect of Antagomir-103a in vivo , the HU+Antagomir-103a group mice received another three consecutive injection of antagomir-103a on day 1-3 at the third week after the first injection.

Statistical analyses All numerical data are expressed as the mean ± s.d. Statistical differences among groups were analyzed by one -way analysis of variance with a post-hoc test (after normalization to baseline in the hindlimb-unloading study) to determine group differences in the study parameters. All statistical analyses were performed with SPSS software, version 13.0. Statistical differences between two groups were determined by the Student’s t test. P < 0.05 was considered statistically significant.

Results

Reduced mechanical loading leads to osteoporosis in a hindlimb unloading mouse model To investigate the mechanical loading preferentially regulating bone development in vivo, we adopted the hindlimb unloading (HU) mouse model which had been widely used to simulate weightlessness and to study various aspects of musculoskeletal loading (Fig. 1a) . Compared with blank control group (weight-bearing ad libitum, group-mean-fed, 6 months old age -matched adult mice, WB) mice, HU mice were tail-hanged for 28 days. The mice were weighed twice per week throughout the experiment. Body weight did not differ between groups (WB, HU) at baseline or at the

end of the 28-day unloading period (28.6±2.7 g vs. 30.1±2.5 g, respectively; P < 0.05). There was distinct thinning of bone tissue and increased vulnerability to fractures in HU mice (Fig. 1b). Similarly, micro computed tomography (microCT) showed that bone volume of distal femurs in HU mice was significantly lower than that in WB mice (Fig. 1c,d). Bone histomorphometric analysis revealed that the bone formation-related parameters (Ob.S/BS, MAR, BFR and N.Ob/B.Pm) were significantly lower in HU mice (Fig. 1e,f), whereas the bone resorption-related parameters (Oc.S/BS and N.Oc/B.Pm) were higher in HU mice (Fig. 1g,h).

Post-transcriptional regulation of Runx2 during CMS-induced osteoblast differentiation To investigate how mechanical loading regulated osteoblast differentiation in vitro, we constructed the CMS-induced osteoblast differentiation model. To obtain a relatively precise simulate cell-loading model, finite element analysis (FEA) was performed to clarify strain distribution in proximal femurs in our study (Fig. 2a). Tissue-level strains in intact human bone are usually less than 1000 µe. (41) Our FEA data also showed the strain loaded on proximal femur bone -level was about 815 ± 57 µe. However, the strain loaded on cells inside the bone is different from the tissue-level strain calculated by FEA.(42) According to the strain amplification mechanism, when mechanical loading was transduced from tissue-level to cellular -level it would be amplified about 100 times. (43-44) Thus, we adopted 8% cyclic mechanical stretch (8% elongation, 80000 µe, Sin, 0.5 Hz, CMS) on human preosteoblast cell line hFOB1.19. After loading for 3 days, quantitative Real-Time PCR (qRT-PCR) analysis showed that the expression of osteoblast marker genes Alkaline phosphatase (ALP), osteocalcin (Ocn) and collagen type I, alpha 1 (Col1a1) was increased in CMS group compare d with non-loading control cells (NC group) (Fig. 2b). Consistent with the above changes, 8% CMS treatment also enhanced ALP enzyme activity and ALP staining (Fig. 2c,d). Besides, we also found CMS could cause distinct rearrangement of cytoskeleton orienta tion compared with static group (Fig. 2e). However, no significant proliferation differences were found between the two groups in our study (Fig. 2f).

Regulation of osteoblastic growth and differentiation occurs through osteogenic signaling pathways among which ERK1/2 MAPK and Wnt/ß -catenin are activated in a duration- and type-of applied-mechanical loading-dependent manner.(45-47) We also found that 8% CMS significantly activated the ERK1/2 MAPK and Wnt/ß -catenin signaling pathways in our study (Fig. 2g). As the 8% CMS can induce osteoblast differentiation, we then used this model to investigate variation of Runx2, the 'master' regulator of osteoblast differentiation. (33-34) We found that Runx2 protein level was significantly increased in 8% CMS stimulation group compared with NC cells, whereas Runx2 mRNA level was only slightly enhanced (Fig. 2h,i). This observation suggested that Runx2 expression under CMS may be mainly regulated at post-transcriptional level, such as miRNA suppression of protein translation or ubiquitin -proteasome degradation.(48)

miR-103a directly targets Runx2 during CMS-induced osteoblast differentiation Runx2 mRNA has a considerably long 3'-untranslated region (3'UTR, 3777 nt), based on RNA blot or the sequence of cDNA clones. We hypothesized that Runx2 may be regulated by miRNAs through their binding to the 3'UTR of Runx2 mRNA, decreasing Runx2 protein level and preventing osteoblast differentiation. Thus, we performed bioinformatic analysis using three miRNA target prediction softwares (i.e, TargetScan, miRDB and miRanda) and miRBase to screen for Runx2-targeting miRNAs (Fig. 3a). miRWalk database was further applied to exclude the miRNAs that had been validated targeting to Runx2 in previous reports. 12 miRNAs including miR-7, miR-22, miR -23b, miR-103a, miR-107, miR -143, miR-154, miR -221, miR-320d, miR -374b, miR -375 and miR -384 were finally acquired to potentially target the human Runx2 3'UTR through multiple binding sites (Fig. 3a, Supplementary. 1a,b). To determine if these miRNAs are regulated during CMS-induced osteoblast differentiation, we treated hFOB1.19 cells under 8% CMS for 3 days and extracted miRNA for qRT-PCR analysis. qRT-PCR showed that the expression of 7 miRNAs including miR -103a, miR-23b and miR -374b was substantially reduced during

osteoblast differentiation (Fig. 3b). In contrast, the expression of miR -107, miR-143 and miR-154 was enhanced (Fig. 3b). To further verify if these chosen miRNAs can directly target Runx2, we constructed a wild -type (WT) Runx2 3'UTR luciferase reporter and then co-transfected with miRNAs mimics and U6 control (the oligo with nonspecific nucleotide sequence) in hFOB1.19 cells. The results of luciferase activity demonstrated that except for miR-107, miR-143 and miR -154, all the other miRNAs inhibited the luciferase reporter activity (Fig. 3c). Interestingly, among the candidate miRNAs, miR-103a mimic inhibited the luciferase reporter activity most significantly. Consistent with the luciferase reporter assays, Western blot analysis also showed that miR-103a mimic decreased Runx2 protein expression most significantly (Fig. 3d). Among the 12 candidate miRNAs, miR-103a and miR-107 are paralogs, which differ only at a single nucleotide near the 3' end of the miRNAs. The miR -103a/miR-107 were previously noted as being upregulated in obese mice and subsequently found to play the key role in insulin sensitivity. (49) Interestingly, in our study, we note that miR-107 presents the diverse effect in luciferase assay and its perturbation has no significant effect on Runx2 protein level (Fig. 3c,d), indicating that the homologous miRNAs may sometimes play the completely opposite roles in different kinds of cells and biological processes. To investigate whether the expression of miR -103a could be regulated by mecha nical force in vivo, we detected miR -103a levels in femurs from HU and WB mice. qRT-PCR showed that miR-103a levels in HU mice but not in WB mice were significantly increased compared with baseline mice (Fig. 3e). Considering that miR -103a demonstrated the most significant suppression on Runx2 both in luciferase assay and Western blot, we then focused on miR -103a for further study. To investigate if gain - and loss-of-function of miR-103a exert post-transcriptional regulation on Runx2 under CMS-loading condition, we used mimic -103a and inhibitor-103a to over-express and knockdown miR -103a in hFOB1.19 cells. Intracellular miR -103a levels were significantly up-regulated by mimic-103a treatment and markedly down-regulated by inhibitor -103a (Fig. 3f). After loading for 3 days under 8% CMS, overexpression of miR-103a decreased Runx2 protein level and knockdown of miR-103a increased Runx2 protein level, whereas only slightly Runx2

mRNA level changes were found (Fig. 3g,h). To further identify the miR -103a target region in the Runx2 mRNA, we constructed a Runx2 3'UTR luciferase reporter which contains mutant sequences of the miR-103a binding sites (MUT Runx2 3'UTR reporter) and then co-transfected with miR-103a oligos in hFOB1.19 cells (Fig. 3i). The luciferase reporter assay demonstrated that mimc-103a decreased, whereas inhibitor-103a increased the WT Runx2 3'UTR luciferase reporter activity but not the MUT Runx2 3'UTR reporter (Fig. 3j). Taken together, all our results demonstrate that miR-103a directly targets Runx2 through interaction with its 3'UTR during CMS-induced osteoblast differentiation.

miR-103a is a mechano -sensitive miRNA during CMS-induced osteoblast differentiation There are two loci align with pre-miR-103a in the human genome. Homo sapiens miR-103a-1 stem-loop located in human chromosome 5 and miR-103a-2 stem -loop located in human chromosome 20. For all known vertebrate species, each miR-103a/107 pa ralog exists within an intron in a gene which also encodes the pantothenate kinase enzyme, PANK. Mature miR -103a-1 and miR-103a-2 are produced from intron 5 of the two separate but related host genes PANK3 and PANK2; respectively (Fig. 4a). The PANK3 and PANK2 enzymes are important for co-enzyme A synthesis and thereby for lipid metabolism. (50-51) To clarify whether the mechanical loading-modulated miR-103a perturbation occurs via PANK3 and/or PANK2, we performed qRT-PCR with primers specific for PANK3 and PANK2. Interestingly, 8% CMS selectively decreased PANK3 but not PANK2 mRNA level with an expression pro?le similar to miR-103a (Fig. 4b). To clarify if miR-103a was specifically sensitive to mechanical stimuli in osteoblast differentiation, hFOB1.19 cells were cultured under osteogenic medium for 3 days and extracted miRNA for qRT-PCR analysis. The results showed that with osteogenic medium treatment, both miR -103a and its host gene PANK3 expressions weren't significantly altered compared with blank control cells (Fig. 4c). All these results indicate that: firstly, the observed CMS-related perturbation

of miR -103a is mainly due to selective production of miR -103-1 from the Pank3 gene locus; secondly, the expressions of miRNAs which located in the introns of their host genes are generally coherent with their corresponding host genes; lastly, miR-103a and its host gene PANK3 are specifically mechano-sensitive. To further clarify the long-term expression profile of miR-103a during CMS-induced osteoblast differentiation, we cultured hFOB1.19 cells under 8% CMS up to 21 days and detected the expression of miR -103a every three days. We found that miR-103a level dramatically downregulated at the early stage of osteoblast differentiation (day 0~9), and then gradually decreased up to day 21 (Fig. 4d). In contrast, we found that ALP and Ocn levels significantly upregulated during hFOB1.19 differentiation (Fig. 4e), whereas Runx2 protein level was strongly increased from the onset of osteoblast differentiation (day 3) and maintained at high level until day 21 (mineralization stage), at which point its level dramatically decreased (Fig. 4f). The modulation in expression and the discrepancy between Runx2 mRNA and protein accumulation at the late stage of osteoblast differentiation may be linked to, at least in part, miR -103a expression modulating Runx2 mRNA. Collectively, our results demonstrate that both miR-103a and its host gene PANK3 are sensitive to mechanical stimuli and negatively regulated during CMS-induced osteoblast differentiation, suggesting that miR-103a could be the mechano-sensitive miRNA.

miR-103a inhibits osteoblast activity and extracellular matrix mineralization during CMS-induced osteoblast differentiation To investigate the role of miR -103a in regulating osteoblast activity, we treated hFOB1.19 cells with miR-103a mimic/inhibitor and their NC-control oligos , respectively. After treated under 8% CMS for 3 days, ALP and Ocn levels were significantly downregulated by mimic-103a and upregulated by inhibitor-103a compared with NC-control oligos (Fig. 5a). Consistently, overexpression of miR -103a weakened ALP activity and ALP staining, whereas knockdown of miR -103a enhanced ALP activity and ALP staining (Fig. 5b,c). The similar role of miR-103a was found in

human bone marrow stromal cells (hBMSCs), suggesting that miR-103a is also important in the differentiation of osteoblasts from uncommitted mesenchymal progenitors (Supplementary Fig. 2a-c). To further investigate the function of miR -103a during extracellular matrix mineralization, we over-expressed and knockdown miR-103a with miR-103a agomir and antagomir (agomir/antagomir, a novel class of chemically engineered specific, efficient and longlasting miRNA agonist/inhibitor)(52) and NC-control (the oligo with nonspecific nucleotide sequence) in hFOB1.19 cells, respectively. To maintain the stable miR -103a expression, the miR -103a agomir and antagomir were supplemented every three days. The intracellular miR -103a levels were timely monitored by qRT-PCR (Fig. 5d). After cultured under 8% CMS for 21 days, Alizarin red staining showed significantly less mineral deposition in agomir-103a-treated cells and more mineralized extracellular matrix formation in antagomir-103a-treated cells than in each corresponding NC-control and mock-treated group (Fig. 5e). To

determine

if

miR-103a-regulated

osteoblast

differentiation

is

Runx2-dependent, we constructed Runx2 siRNA and examined its effect on miR-103a-regulated osteoblast differentiation (Fig. 5f) . We found that transfection of Runx2 siRNA significantly reduced ALP, Ocn mRNA levels and ALP activity (Fig. 5g). However, when we co-transfected miR -103a mimic and inhibitor with Runx2 siRNA, the functions of miR-103a oligos were completely blocked; ALP, Ocn mRNA levels as well as ALP activities persistently remained low, suggesting that the function of miR-103a in CMS-induced osteoblast differentiation is Runx2-dependent (Fig. 5g). To clarify if the expression of miR -103a in CMS-induced osteoblast differentiation was regulated by ERK1/2 MAPK and Wnt/ß-catenin signaling pathways, we separately blocked the two pathway with U0126 and IWR-1 (U0126, the inhibitor of ERK1/2 pathway; IWR-1, the inhibitor of Wnt/ß-catenin pathway) in hFOB1.19 cells and then detected the expression of miR-103a , respectively (Fig. 5h). qRT-PCR showed that there was no significantly perturbation of miR-103a level under the treatment of U0126 and IWR-1. Taken together, our results suggest that miR -103a plays a negative role in

CMS-induced osteoblast differentiation and subsequent mineralization through suppression of Runx2.

miR-103a counteracts the decrease of bone formation in the hindlimb unloading mouse model As we had found miR -103a could play an essential role during CMS-induced osteoblast differentiation in vitro, we further verified if it can play the same role in vivo. As miR-103a is evolutionarily conserved across many vertebrate species including mouse (Supplementary Fig. 3a), thus, we could explore the relationship between miR-103a expression and bone formation in mouse model. First, we examined the expression profile of miR-103a in different tissues in C57BL/6J mice. The results clearly showed that the expression of miR-103a in bone was much higher than in other tissues, indicating that miR -103a may play a major role in bone remodeling (Fig. 6 a). To further test whether therapeutic inhibition of miR -103a could counteract the mechanical unloading-induced bone loss, the HU groups were separately pretreated with Antagomir-103a (HU+Antagomir-103a, Antagomir-103a, 80 mg/kg) and PBS (HU+PBS, PBS, 0.2ml) for 3 consecutive injections via caudal vein before unloading (Fig. 6b). To maintain the effect of miR-103a on mice, the HU+Antagomir -103a mice received another injection of antagomir-103a on day 1-3 at the third week after the first injection (Fig. 6b). All mice were euthanized for analysis after 28-day-hindlimp unloading. Several pharmacological properties for antagomir-103a were evaluated further, including bioavailability, silencing activity, and duration of action. We firstly investigated the bioavailability and silencing activity of antagomir -103a in different tissues including bone, heart and muscle tissues, because miR-103a is expressed abundantly in these tissues. In mice treated with antagomir-103a, miR-103a was ef?ciently silenced in the above-mentioned tissue samples (Supplementary Fig. 3b-d). We also tested the duration of silencing that could be achieved after the injection of antagomir-103a. Levels of miR-103a in bone tissue were significantly downregulated for as long as 11 days after the last injection, indicating that silencing of miR -103a using antagomir is long lasting (Fig. 6c). These data show that antagomir-103a can

efficiently silence miR-103a in vivo. Consistent with the ability to silence miR-103a, we found that compared with HU and HU+PBS mice, HU+Antagomir -103a mice had higher amounts of Runx2 protein in femurs (Fig. 6d), indicating that antagomir-103a can partly counteract the down-regulated Runx2 expression caused by hindlimb unloading. We also detected other 11 screening miRNAs in our in vivo models. No significantly changes were observed among each group (Supplementary Fig. 4). More importantly, microCT showed mechanical unloading-caused bone loss was partly rescued by Antagomir -103a (Fig. 6e,f). Bone histomorphometric analysis revealed that the bone formation-related parameters (Ob.S/BS, MAR, BFR and N.Ob/B.Pm) were higher in HU+Antagomir -103a mice compared with HU and HU+PBS mice (Fig. 6g,h), whereas the bone resorption-related parameters (Oc.S/BS and N.Oc/B.Pm) were not significantly different in all groups (Fig. 6i,j). All these results indicated that: First, the expression of miR -103a and Runx2 in bone tissue could be modulated by mechanical loading; Second, mechanical unloading- induced decreased Runx2 protein level and bone loss is through, at least in part, up-regulating miR -103a level in vivo ; Third, therapeutic inhibition of miR -103a could partly counteract the bone loss in HU mice.

Discussion The present study demonstrated that miRNAs could be regulated by mechanical loading and play essential roles in osteoblast differentiation and bone formation. Specifically, we identified miR-103a as a novel mechano-sensitive miRNA which negatively regulates osteoblast differentia tion and bone formation under both physiological and pathological mechanical loading condition by repressing Runx2 expression at the post-transcriptional level. Furthermore, our experiments in vivo suggest that miR-103a could be a potential therapeutic regimen for pathological mechanical loading-caused disorders of skeletal development. To our knowledge, the identification of miR-103a as a mechano-sensitive miRNA in progenitor osteoblasts represents the first evidence linking mechanical stretch and miRNAs to bone formation.

Mechanical stretch has been previously reported as an important regulator in diverse biological and pathological processes. (53-58) The present study demonstrated that osteoblasts are subjected to the external applied CMS, leading to rapid adaptations in their cellular functions and structures. Furthermore, we found that CMS could significantly activate transcription factor, namely Runx2, the principal transcriptional regulator

of

osteoblast

differentiation,

and

contribute

to

the

control

of

osteoblastogenesis both in vitro and in vivo . As an essential bone-speci?c transcription factor in osteogenesis, Runx2 can bind to osteoblast-specific cis-element 2 (OSE2) found in the promoter region of all major osteoblast-related genes. (59) Besides, Runx2 has a considerably long 3'UTR (~4 kb), presumably containing multiple regulatory elements. Because of its large assortment of binding partners and co-modulators, Runx2 is likely subjected to be activated by several mechanisms in osteoblasts. (60) Phosphorylation of Smad by BMPs and TGF-ß activates Runx2 as a crucial event in osteoblast maturation. By contrast, PTH and the fibroblast growth factors (FGFs) directly phosphorylate Runx2, and TNF leads to ubiquitination of Runx2 through the ligases Smurf-1 and Smurf-2.(61) In our study, intriguingly, we found that Runx2 protein level was significantly induced by CMS, whereas its mRNA level was only slightly changed. All the above prompted us to investigate whether there exsits miRNA regulation, since miRNAs have emerged as a new regulatory mechanism of gene expression and a key modulator of variety of biological and pathological processes in recent years.(22 -27) Several studies indicated that mechanical force including shear stress and cyclic stretch, could modulate the expression of a panel of miRNAs whic h are involved in the cellular response to mechanical force in different cell lines. (53-57) More recently, multiple miRNAs have been demonstrated as important regulators of bone remodeling related gene expression at the post-transcriptional level and further contribute to human skeletal disorders, such as osteopenia or osteoporosis. (62-63) However, the function of specific mechano-sensitive miRNAs in CMS-induced osteoblasts differentiation have not been well characterized and are, therefore, poorly understood. In our study, we found that several Runx2-targeting miRNAs could be regulated by

cyclic mechanical stimulation, indicating these miRNAs might also play roles in CMS-induced osteoblast differentiation. Of note, the greatest fold increased changes are found in miR -107 and miR-143, yet both of which are failed to downregulate Runx2 protein level in our study. These two miRNAs had been reported to play a role in insulin-sensitivity regulation

(49,64)

. Considering the reciprocal relationship between

osteogenesis and adipogenesis, it's possible that miR-107 and miR-143 might also function through other targets involved in adipocyte metabolism that contributing to the osteoblast response to in vitro mechanical stimulation. However, there are no reports about the function of these two miRNAs in CMS-induced osteoblast differentiation, and further research might be needed to address this issue. miR -103a (homologous to miR -107) is recruited to repress the expression of Runx2 through sequence specific base pairing with multiple conserved binding sites in Runx2 3'UTR and further regulates osteoblast differentiation. miR-103a/miR -107 have now been predicted or experimentally confirmed in human. They were originally noted to be up-regulated in obese mice and subsequently found to have a key role in insulin sensitivity, suggesting that these miRNAs represent potential targets for the treatment of type 2 diabetes. (49) miR-103a has also been linked with chronic pain and intestinal cell proliferation. (65) Like many other miRNAs, miR -103a highly expressed in many tumor cells, indicating that miR -103a also participates in carcinogenesis and tumor progression. (66) More recently, several studies disclosed that miR -103a could promote adipocyte

differentiation

and

further

modulate

homeostasis

of

glucolipid

metabolism.(50-51) However, there is no report regarding the role of miR-103a in the regulation of osteoblast function. It’s the first time to link the expression of the miRNAs during cyclic stretch with a bone homeostasis phenotype known to be induced by mechanical stretch. A large number of miRNAs have been identified to be located within the intronic regions of protein-encoding genes (host genes), so-called 'intronic miRNAs'. Due to their specific location in genome, most of them are co-expressed with the host genes and play the similar roles. In our study, we found that 8% CMS selectively decreased PANK3 but not PANK2 (the host genes of miR-103a) mRNA level with an expression

profile similar to miR -103a, whereas both PANK2 and PANK3 expressions weren't significantly altered during osteoblast differentiation induced by osteogenic medium. PANK3, the host gene of miR-103a, was originally reported to play important roles in co-enzyme A synthesis and lipid metabolism(50-51) and found being downregulated in CMS-induced osteoblast differentiation in our study. As miR -103a is located within the intronic region of PANK3 and own the same promoter, so mechanical loading might regulate miR-103a expression via activate the promoter of PANK3. Nevertheless, the exactly role of PANK3 in osteogenesis had not been reported before and further studies are needed to explore whether PANK3 can also play any functions in osteoblast differentiation as well as its potential effects under CMS conditions. However, our data support the model that miR-103a might co-expressed with its host gene PANK3 to fine-tune Runx2 gene expression. Upon stimulation of the mechanosensors in osteoblast biology, key intracellular enzymes are induced, and MAPKs, Cox-2, NO, TNF-a and Wnt-ß-catenin are activated

in

a

duration-

and

type-of-applied-force-dependent

manner. (67)

Mechanostimulation of osteoblasts also induces secretion of growth factors, including IGF, VEGF, PDGF, bFGF, TGF-ß and the BMP, which are considered to be principal local regulators of osteogenesis. (60) The activation of intracellular signaling pathways and growth factors might converge to regulate specific miRNAs' perturbation. The activation of the ERK1/2 MAPK and Wnt–ß-catenin signaling pathways has an anabolic

osteoblastic

role. (45-47)

Inactivation of

ß-catenin

blunts

osteoblast

differentiation from mesenchymal progenitors, indicating that ß-catenin has an essential role in osteoblast differentiation in vivo.(67) In our study, we find that CMS can significantly activate both ERK1/2 and ß-catenin pathways and further activate their down-stream genes, including Runx2. However, we found that inhibition of these two signaling pathways don't affect the expression of miR-103a in our study, indicating that the expression of miR -103a is not under the control of the two signaling pathways in CMS-induced osteoblast differentiation. Mechanostimulation can also act as a therapeutic regimen for bone disease.(68-75) Dynamic strain applied on osteoblastic cells activates the Wnt-ß-catenin pathway and

thus induces osteoblastogenesis. (47) Bone fracture healing has also benefited from mechanostimulation of osteoblasts. Osteoblasts and osteocytes respond differently in varied mechanical environments and, as a result, rigid fixation of fractures results in direct bone formation and osteonal bridging of the fracture gap, whereas flexible fixation can result in indirect healing characterized by periosteal callus formation and enchondral bone formation. (73) In our study, our experimental evidence from in vivo studies also defined a new mechanism whereby miR -103a mediates its regulation role in bone formation. miR-103a was highly expressed in bone tissues, which indicates that it may play a key role in bone remodeling. Our data demonstrate that the change of miR-103a level in physiological mechanical condition results in an alteration in the amount of Runx2 protein . Runx2 activity further favors bone formation by promoting its downstream osteoblast-specific marker genes expression. While in the hindlimb-unloading mouse model, therapeutic inhibition of miR-103a in vivo may promote bone formation by exerting an anabolic effect under pathological mechanical conditions. However, as we know that miRNAs can regulate the mRNA levels of their targets, and pharmacological sile ncing of miRNAs using antagomirs might therefore lead to the regulation of lots of mRNAs.(52) According to bioinformatics analysis, t here are hundreds of predicted targets for miR -103a including parathyroid hormone (PTH) and s pecia l AT-rich sequence binding protein 2 (SATB2), which themselves also have well-known effects on bone biology. We cannot exclude the possibility that these miR-103a-targeting genes might also engaged in the mechanical stress-regulated bone formation in vivo. Likewise, increased bone formation after knockdown of miR -103a by its antagomir in HU mouse model cannot be attributed to elevated Runx2 protein level alone. Nevertheless, our results clearly showed that Runx2 is one of the main targets of miR -103a in mechanical stress-regulated bone formation both in vitro and in vivo. In summary, our study provides a new finding that a specific miRNA-miR-103a as a mechano-sensitive miRNA in osteoblasts which can be targeted in these cells to inhibit bone formation under both physiological and pathological mechanical conditions in vitro and in vivo . miR -103a functions through inhibiting its direct target

Runx2, the master regulator during osteogenesis, at the post-transcription level. These findings not only provide ne w insights into mechano-transduction signaling pathways, but also raise intriguing possibilities of using miRNAs for modulating bone tissue engineering in regenerative medicine. We anticipate that our study can provide a foundation for future investigations on the role of miRNAs in regulating the osteoblast cells response to mechanical loading, serve as an effective model system for studying gene regulation by miRNAs in the development of gene therapy for treating human bone remodeling disorders related to mechanical loading such as osteopenia and osteoporosis.

Conflict of Interest The authors have declared that no conflict of interest exists. Acknowledgements This work was supported by the National Natural Science Foundation of China (Grant

nos.

811 71705

&

81190133).

Chinese

Academy

of

Sciences

(No.XDA01030404), Science and Technology Commission of Shanghai Municipality (No. 12411951100 & 12410708600). Author contributions Bin Zuo: Conception and design, manuscript writing, collection of data, data analysis and interpretation; Junfeng Zhu and Jiao Li: Collection of data, data analysis and interpretation; ChuanDong Wang, XiaoYing Zhao, Guiquan Cai, Zheng Li: Provision of study material; Jianping Peng, Peng Wang, Chao Shen: Data analysis and manuscript revision; Jiake Xu, Xiaoling Zhang and Xiaodong Chen: Conception and design, manuscript revision, final approval of manuscript. References 1. Harada S, Rodan GA. Control of osteoblast function and regulation of bone mass. Nature. 2003;423:349-55. 2. Zaidi M. Skeletal remodeling in health and disease. Nat Med. 2007;13:791-801. 3. Hsieh YF, Turner CH. Effects of loading frequency on mechanically induced bone formation. J Bone Miner Res. 2001;16:918-24.

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formation-related parameters (Ob.S/BS, MAR, BFR and N.Ob/B.Pm) in BS, WB and HU mice. Ob.S/BS, osteoblast surface per bone surface; MAR, mineral apposition rate; N.Ob/B.Pm, number of osteoblasts per bone parameter. (g) Tartrate-resistant acid phosphata se (TRAP) and hematoxylin staining images of distal femurs from BS, WB and HU mice. Scale bar, 100 µm. (h) Histomorphometric analysis of bone resorption-related parameters (Oc.S/B.S, N.Oc/B.Pm) in BS, WB and HU mice. Oc.S/B.S, osteoclast surface per bone surface; N.Oc/B.Pm, number of osteoclasts per bone parameter. All data are shown as the mean ± s.d. *P < 0.05, **P < 0.01, ***P < 0.001. NS, not significant. All P values based on Student's t test. For each group mice n = 6. Figure 2 Cyclic mechanical loading could regulate osteoblast differentiation in vitro. (a) A three-dimensional finite element (FE) model of the proximal human femur. 29,841 elements were generated in the FE model. The strain magnitude distribution on the proximal femur bone tissue -level was about 815 ± 57µe (from 375 µe to 1,583 µe). (b) Quantitative Real-time PCR analysis (qRT-PCR) of Ocn, ALP and Col1a1 mRNA levels in hFOB1.19 cells after treated with 8% CMS for 3 d compared with static control cells. CMS, cyclic mechanical stretch (8% elongation, Sin, 0.5 Hz). (c) ALP activities in hFOB1.19 cells after treated with 8% CMS for 3 d compared with static control cells. (d) Representative images of ALP staining of hFOB1.19 cells after treated with 8% CMS for 3 d compared with static control cells. Scale bars, 10 mm. (e) Immunostaining of F-actin filaments showed rearrangement of cytoskeleton orientation in 8% CMS treated hFOB1.19 cells compared with static control cells. (f) Cell viability was examined by Alamar Blue after treated with 8% CMS for 72 h compared with static control cells. (g) Western blot analysis of Wnt/ß -catenin and Erk1/2 MAPK signaling pathway components in 8% CMS treated hFOB1.19 cells. (h) Western blot analysis of Runx2 protein in hFOB1.19 cells after treated with 8% CMS for 3 d compared with static control. (i) qRT-PCR analysis of Runx2 mRNA levels in hFOB1.19 cells after treated with 8% CMS for 3 d compared with static control. All the staining data were confirmed by three repeated tests. All data are shown as the mean ± s.d. *P < 0.05, **P < 0.01, ***P < 0.001. NS, not significant. All P values based on Student's t test. Figure 3 miR-103a targets Runx2 to functionally inhibit osteoblast activity in vitro. (a) Bioinformatic analysis by using miRNA target prediction softwares: TargetScan, miRDB, miRanda and miRWalk to screen for Runx2-targeting miRNAs. (b) qRT-PCR analysis of miRNAs expression in hFOB1.19 cells treated with 8% CMS for 3 d. miRNAs expression was expressed as a ratio to that in static controls. (c) The effect of NC-control and 12 selected miRNA mimics on luciferase activity of WT Runx2 3'UTR reporter in hFOB1.19 cells. (d) The effect of NC-control and 12 selected miRNA mimics on the amount of Runx2 protein in hFOB1.19 cells. (e) qRT-PCR analysis of miR-103a levels in femurs (normalized to those in BS mice) from WB and HU mice. (f) qRT-PCR analysis of miR-103a levels in hFOB1.19 cells after treated with mimic-103a, inhibitor-103a or their negative controls (mimic -NC, inhibitor-NC). (g)

The effect of mimic -103a, inhibitor-103a or their negative controls on the amount of Runx2 protein in hFOB1.19 cells. (h) The effect of mimic -103a, inhibitor -103a or their negative controls on Runx2 mRNA levels in hFOB1.19 cells. (i) A schematic diagram illustrating the design of luciferase reporters with the WT Runx2 3'UTR (WT 3'UTR) or the site-directed mutant Runx2 3'UTR (MUT 3'UTR). hRluc, human Renilla luciferase. (j) The effect of miR -103a mimic, inhibitor or their negative controls on the luciferase activity of WT Runx2 3'UTR or MUT Runx2 3'UTR reporter in hFOB1.19 cells. All data are shown as the mean ± s.d. from three independent experiments. *P < 0.05, **P < 0.01, ***P < 0.001. NS, not significant. All P values based on Student's t test. Figure 4 miR-103a levels are negatively regulated by cyclic mechanical loading in vitro. (a) Schematic diagram of genomic localization of miR -103a and miR -107. PANK genes are shown with exons as rectangles and introns as crooked lines. Note: the exon/intron lengths on these gene diagrams are not to scale. (b) qRT-PCR analysis of miR-103a, PANK2 and PANK3 levels in hFOB1.19 cells treated with cyclic mechanical stretch (8% elongation, Sin, 0.5 Hz) for 3 d compared with static control. (c) qRT-PCR analysis of miR-103a and PANK3 levels in hFOB1.19 cells after treate d with osteogenic medium for 3 d compared with blank control. (d) qRT-PCR analysis of the time course changes of miR-103a levels during 8% CMS-induced osteoblast maturation and mineralization in hFOB1.19 cells for 21 dcompared with static control. (e) qRT-PCR analysis of the time course changes of Ocn and ALP mRNA levels in hFOB1.19 cells under 8% CMS treatment for 21 dcompared with static control. (f) Western blot analysis of Runx2 protein amounts during 8% CMS-induced osteoblast maturation and mineraliz ation in hFOB1.19 cells for 21 d. All data are shown as the mean ± s.d. from three independent experiments. *P < 0.05, **P < 0.01, ***P < 0.001. NS, not significant. All P values based on Student's t test. Figure 5 miR-103a inhibits osteoblast activity and matrix mineralization in vitro. (a) The effect of mimic-103a, inhibitor-103a or their negative controls on Ocn and ALP mRNA levels in hFOB1.19 cells under 8% CMS for 3 d. (b) The effect of mimic-103a, inhibitor-103a or their negative controls on ALP activities in hFOB1.19 cells under 8% CMS for 3 d. (c) Representative images of ALP staining of hFOB1.19 cells after treated with mimic-103a, inhibitor -103a or their negative controls under 8% CMS for 3 d. Scale bars, 10 mm. (d) qRT-PCR analysis of the time course changes of miR-103a level in hFOB1.19 cells after treatment with Agomir-103a or Antagomir-103a under 8% CMS treatment for 21 d compared with mock control. (e) Staining of calcium deposition by Alizarin red in hFOB1.19 cells after treatment with 200 µM agomir-103a, antagomir-103a or the ir corresponding negative controls under 8% CMS for 21 d. Scale bars, 10 mm. All the staining data were confirmed by three repeated tests. (f) The knockdown efficiency of Runx2-specific siRNA (siRNA-Runx2) was confirmed by comparison to a scrambled control siRNA (siRNA-NC). (g) qRT-PCR analysis of Ocn, ALP mRNA levels and ALP activities in hFOB1.19 cells after co-tranfected siRNA-Runx2 w ith mimic-103a, inhibitor-103a or their corresponding negative controls under 8% CMS for 3 d. (h) Real-time PCR analysis of

miR-103a levels after treated with Wnt/ß-catenin signaling pathway inhibitor IWR-1 and Erk1/2 MAPK signaling pathway inhibitor U0126 in hFOB1.19 cells under 8% CMS for 3 d. All the staining data were confirmed by three repeated tests. All data are shown as the mean ± s.d. *P < 0.05, **P < 0.01, ***P < 0.001. NS, not significant. All P values based on Student's t test. Figure 6 Therapeutic inhibition of miR-103a partly counteracts mechanical unloading -caused decreasing bone formation in vivo. (a) qRT-PCR analysis of miR-103a levels in bone and other tissues from C57BL/6J mice. (b) A schematic diagram illustrating the experimental design (for each group mice, n=6). HU+PBS, hind-limb unloading mice treated with PBS; HU+Antagomir -103a, hind-limb unloading mice treated with antagomir -103a. (c) qRT-PCR analysis of miR-103a levels in femurs (normalized to those in WB mice) from HU, HU+PBS mice and HU+Antagomir-103a mice. (d) Western blot analysis of Runx2 protein amounts in femurs collected from WB, HU, HU+PBS, HU+Antagomir -103a mice (for each group mice, n=3). (e) Representative microCT reconstructive images of distal femurs in WB and HU, HU+PBS mice and HU+Antagomir -103a mice. Scale bar, 1 mm. (f) Three-dimensional microstructural parameters of the distal femurs from WB, HU, HU+PBS mice and HU+Antagomir -103a mice. (g) Representative images showing new bone formation assessed by double calcein labeling in each group. Scale bars, 10 µm. (h) Histomorphometric analysis of bone formation-related parameters (Ob.S/BS, MAR, BFR and N.Ob/B.Pm) in WB, HU, HU+PBS mice and HU+Antagomir-103a mice. ( i) TRAP and hematoxylin staining images of distal femurs in WB, HU, HU+PBS and HU+Antagomir -103a mice. Scale bar, 100 µm. (j) Histomorphometric analysis of bone resorption-related parameters (Oc.S/B.S, N.Oc/B.Pm) in WB, HU, HU+PBS and HU+Antagomir -103a mice. All data are shown as the mean ± s.d. *P < 0.05, **P < 0.01, ***P < 0.001. NS, not significant. All P values based on Student's t test.

Supplemental information MiR-103a: a novel mechano-sensitive microRNA inhibits bone formation through targeting Runx2 Bin Zuo,1*JunFeng Zhu,1*Jiao Li,2GuiQuan Cai,1Zheng Li,1Jianping Peng,1Peng Wang,1Chao

Shen,1ChuanDong

Wang,2 XiaoYing

Zhao,2Yan

Huang,2Jiake

Xu3,XiaoLin g Zhang,2‡XiaoDong Chen1‡

Supplementary Figure 1 The binding sites of predicted Runx2-targeting miRNAs with Runx2 3'UTR by bioinformatics analysis. (a) The binding sites of some predicted Runx2-targeting miRNAs with Runx2 3'UTR by bioinformatics analysis using miRNA target prediction softwares (Target Scan, miRDB, miRanda ). (b) The binding sites of 3 validated Runx2-targeting miRNAs with Runx2 3'UTR by bioinformatics analysis using miRNA target prediction softwares

miRWalk. Supplementary Figure 2 miR-103a inhibits osteoblast activity and matrix mineralization in hBMSCs. (a) The effect of mimic-103a, inhibitor -103a or their negative controls on Ocn and ALP mRNA levels in hBMSCs under 8% CMS for 3 d. ( b) The effect of mimic -103a, inhibitor-103a or their negative controls on the amount of Runx2 protein in hBMSCs. (c) Representative images of ALP staining of hBMSCs after treated with mimic -103a, inhibitor-103a or their negative controls under 8% CMS for 3 d. Scale bars, 10 mm. All the staining data were confirmed by three repeated tests. All data are shown as the mean ± s.d. from three independent experiments. *P < 0.05, **P < 0.01, ***P < 0.001. NS, not significant. All P values based on Student's t test. Supplementary Figure 3 Antagomir-103a target microRNA expression in multiple tissues. (a) Schematic diagram of the secondary structure of precursor of hsa-miR-103a and sequence comparison of mature miR -103a among mammals or vertebrates. Hsa for human, mmu for mouse, ptr for chimpanzee, mml for rhesus, laf for elephant, mdo for opossum, dno for armadillo, gga for chicken. (b) qRT-PCR analysis of miR-103a levels in bone tissues 48 h after injection of antagomir -103a. (c) qRT-PCR analysis of miR-103a levels in heart tissues 48 h after injection of antagomir-103a. (d) qRT-PCR analysis of miR -103a levels in muscle tissues 48 h after injection of antagomir-103a. All data are shown as the mean ± s.d. from three independent experiments. *P < 0.05, **P < 0.01, ***P < 0.001. NS, not significant. All P values based on Student's t test. For each group mice n = 3. Supplementary Figure 4 Levels of eleven predicted miRNAs in bone specimens from four groups mice. (a)-(k) qRT-PCR analysis of 11 screening miRNAs levels in femur s from HU, HU+PBS mice and HU+Antagomir -103a mice. The relative miRNA levels were normalized to the mean value of the WB group mice. All data are shown as the mean ± s.d. from three independent experiments. *P < 0.05, **P < 0.01, ***P < 0.001. NS, not significant. All P values based on Student's t test. For each group mice n = 4.

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