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American Journal of Translational Research logoLink to American Journal of Translational Research
. 2019 Jun 15;11(6):3398–3411.

High-throughput sequencing analysis of the expression profile of microRNAs and target genes in mechanical force-induced osteoblastic/cementoblastic differentiation of human periodontal ligament cells

Yun Wu 1,2, Yanjing Ou 1,2, Chufang Liao 1,2, Shanshan Liang 1,2, Yining Wang 1,2
PMCID: PMC6614645  PMID: 31312353

Abstract

Mechanical tension force directs the lineage commitment of periodontal ligament cells (PDLCs) to osteogenesis; however, the underlying mechanisms, especially those at the post-transcriptional level, remain unclear. In the present study, we developed an in vitro force-loading model for PDLCs. Then, high-throughput sequencing was used to identify the expression profile of microRNAs (miRNAs) for stretched PDLCs. The candidate target genes of differentially expressed miRNAs were predicted by bioinformatics analysis. A total of 47 miRNAs were found to be differentially expressed in stretched and non-stretched PDLCs; of these, 31 were upregulated and 16 were downregulated. Further, 9 osteogenesis-related miRNAs (miR-221-3p, miR-138-5p, miR-132-3p, miR-218-5p, miR-133a-3p, miR-145-3p, miR-143-5p, miR-486-3p, and miR-21-3p) were validated by quantitative reverse transcription-polymerase chain reaction (RT-qPCR). Gene Ontology (GO) and Kyoto Encyclopedia of Gene and Genome (KEGG) pathway analysis were then carried out to reveal the potential functions of predicted target genes. Among the top 20 enriched pathways, the Hippo signaling pathway was selected for further functional analysis. Several important components of the Hippo signaling pathway, including YAP1, WWTR1, TEAD2, CTGF, DVL2, GDF5, GLI2, LIMD1, WTIP, LATS1, and TEAD1, were predicted to be target genes of differentially expressed miRNAs and were determined to be upregulated in stretched PDLCs. Among them, YAP1, WWTR1, TEAD2, CTGF, DVL2, and GDF5 were positive regulators of osteogenesis. These findings may provide a reliable reference for future studies to elucidate the biological mechanisms of orthodontic tooth movement (OTM).

Keywords: Periodontal ligament cells, osteogenesis, high-throughput sequencing, microRNA, orthodontic tooth movement

Introduction

Orthodontic tooth movement (OTM) relies on coordinated tissue resorption and formation in the surrounding bone and periodontal ligament. The periodontal ligament (PDL) is a dense fibrous connective tissue structure consisting of collagenous fiber bundles, cells, neural and vascular components, and tissue fluids [1]. Its primary function is to support the teeth in their sockets while allowing the teeth to withstand considerable chewing forces. The PDL acts as an important medium, transducing the mechanical force from the teeth to the alveolar bone, and then transferring it into biochemical signals [2,3]. Bone formation and resorption happens at the tension side and compressive side, respectively [4]. To date, the underlying mechanisms by which the PDL perceives mechanical loading and initiates the process of bone remodeling have not yet been fully elucidated.

PDL cells (PDLCs), a heterogeneous cell population within the PDL, are considered to be the first cellular recipients of mechanical stimuli. Previous studies have demonstrated that PDLCs possess a portion of mesenchymal stem cells, which can further differentiate to an osteoblastic and cementoblastic phenotype and involve in the alveolar bone and root cementum remodeling process during the orthodontic tooth treatment [5-7]. Besides, it was shown that multiple regulators, including cytokines, growth factors, cytoskeleton proteins, and signaling pathways (e.g., ERK1/2, p38 mitogen-activated protein kinases (MAPK) signaling pathway, ephrin B2/EphB4 signaling) are involved in the modulation of the osteogenic activities of PDLCs under mechanical stimuli [8-10]. Recently, another regulatory mechanism involved in osteoblastic/cementoblastic differentiation of PDLCs after mechanical force loading have been reported to be post-transcriptional modulation of gene expression by microRNAs (miRNAs).

miRNAs are short non-coding RNAs composed of approximately 22 nucleotides; they regulate gene expression by selectively binding to the 3’ untranslated regions (3’-UTR) of target mRNAs, resulting in mRNA degradation or protein translation repression [11,12]. In recent years, increasing evidence has indicated that miRNAs are involved in the cellular response to various mechanical stimuli in diverse organs and cells. Among them, miR-33-5p was identified as a novel mechano-sensitive miRNA that positively regulates osteoblast differentiation by repressing the expression of high mobility group AT-hook 2 (Hmga2) [13]. Additionally, miR-503-5p is reported to function as a mechano-responsive miRNA which can inhibit osteogenic differentiation of bone marrow-derived mesenchymal stem cells (BMSCs) subjected to mechanical stretch and bone formation in OTM tension sides [14]. Under tensile stress, miR-154-5p was found to negatively regulate osteogenic differentiation of adipose tissue-derived stem cells (ADSCs) through the Wnt/planar cell polarity (PCP) signaling pathway by directly targeting Wnt11 [15]. In addition, miRNAs also participate in osteogenesis of PDLCs under mechanical loading. Previous studies have shown that miR-132 regulates the osteogenic differentiation of PDLCs after fluid shear stress treatment through the mTOR signaling pathway [16]. Of note, miR-21 has been reported to play a role in osteogenic differentiation of PDLSCs exposed to stretch in vitro, and can modulate OTM and alveolar bone remolding in the presence of both normal and inflammatory microenvironments in vivo [17,18]. Further, miR-195-5p was found to be downregulated and negatively correlated with osteogenic differentiation of PDLCs under cyclic tension straining for 72 hours [19]. Although several miRNAs have been identified to play a role in stretch-induced osteogenic differentiation of PDLCs, the functional roles and regulatory mechanisms of miRNAs during osteogenesis and bone formation under force-loading have not been fully explored.

High-throughput sequencing techniques have been successfully used to identify miRNAs in a variety of organisms. Compared with microarray assay, next-generation sequencing allows deep sequencing in a short period of time, and has been able to identify new, previously unreported miRNAs [20]. In the present study, to further clarify the regulatory effect of miRNAs in osteoblastic/cementoblastic differentiation of stretched PDLCs, we examined differentially expressed miRNAs in stretched PDLCs and non-stretched PDLCs using high-throughput sequencing. Relative target genes of differentially expressed miRNAs were also predicted using bioinformatics analysis. Then, these putative target genes were analyzed by Gene Ontology (GO) and Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway analyses. Additionally, the interaction between osteogenesis-related transcription factors and changed miRNAs was predicted using bioinformatics tools in order to investigate osteoblastic/cementoblastic mechanisms of PDLCs under force loading.

Materials and methods

Cell culture

Healthy third molar teeth were collected in a clinical setting from individuals aged 14-25 years. Informed consent was obtained from all participants and the study protocol was approved by the Ethics Committee, School of Stomatology, Wuhan University, Wuhan, China. Human PDLCs were obtained from human tissue scraped from the roots of the extracted teeth. In brief, extracted teeth were rinsed three times with phosphate-buffered saline (PBS) containing 10% penicillin and streptomycin. Tissue was obtained from the middle one-third of the roots, and was placed into a 25 cm2 tissue culture flask containing α-modified essential medium (α-MEM; HyClone, Logan, UT, USA) supplemented with 20% fetal bovine serum (FBS; HyClone, Logan, UT, USA), 100 U/mL penicillin G, and 100 μg/mL streptomycin. The explants were incubated at 37°C with 5% CO2. When sub-confluence was reached, PDLCs were passaged; cells at passages 3-6 were used for the following experiments.

Flow cytometry analysis

To identify the phenotype of PDLCs, cell surface markers were evaluated by flow cytometry. Briefly, a total of 1 × 106 cells were collected and incubated with FITC-labeled anti-CD146 antibodies (Biolegend, San Diego, California), anti-CD90 antibodies (Biolegend, San Diego, California), or anti-CD45 antibodies (Biolegend, San Diego, California) for 30 min at 4°C. Then, cells were washed three times with PBS and analyzed by flow cytometry analysis. The results were analyzed with FlowJo software (Tree Star, Ashland, OR, USA).

Osteoblast and adipocyte differentiation

PDLCs were seeded in six-well plates at a density of 1 × 105 per well. After reaching 70-80% confluence, cells were incubated with osteogenic medium containing 50 μg/ml ascorbic acid, 10 mmol/L β-glycerophosphate, and 10 nmol/L dexamethasone. The medium was changed every three days. After osteogenic induction for 7 days and 21 days, extracellular matrix calcification was investigated with alkaline phosphatase (ALP) staining with a BCIP/NBT kit (Beyotime, China) and Alizarin red staining, respectively.

For adipocyte differentiation, PDLCs were cultured in adipogenic medium, according to the manufacturer’s instructions (Cyagen, Suzhou, China). Two weeks later, Oil Red O staining was used to detect lipid formation.

Application of tension force system

External mechanical stimulation was achieved with a Flexcell FX-5000TM tension system (Flexcell International Corp., Burlington, NC, USA). PDLCs were plated onto six-well Bioflex plates coated with type I collagen (Flexcell International Corp., Burlington, NC, USA) at a density of approximately 1 × 105 cells/well. When the culture reached 70%-80% confluence, they were stretched with 10% equibiaxial strain at 0.1 Hz for 24 h in a Flexcell FX-5000TM tension system (Flexcell International Corp., Burlington, NC, USA). The control cells were cultured without stretching.

Western blot analysis

For Western blot analysis, cell lysates were prepared with radio-immunoprecipitation assay (RIPA) lysis buffer supplemented with phenylmethylsulfonyl fluoride (PMSF) (Roche, Germany) and phosphatase inhibitor (Roche, Germany). The samples were separated on a 10% sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE), and transferred onto 0.22 μm polyvinylidene difluoride (PVDF) membranes. The membranes were blocked with 5% non-fat milk for 1 h at room temperature. Subsequently, these membranes were incubated with primary antibodies, including rabbit anti-runt-related transcription 2 (RUNX2) (1:500; Abclonal, Boston, MA), rabbit anti-secreted phosphoprotein 1 (SPP1) (1:1000; Abclonal, Boston, MA), rabbit anti-cementoblastoma-derived protein 1 (CEMP1) (1:500; Abcam, Cambridge, UK), and mouse anti-glyceraldehyde 3-phosphate dehydrogenase (GAPDH) (1:5000; Abcam, Cambridge, UK) monoclonal antibody, overnight at 4°C. Membranes were then washed three times with TBST and incubated with horseradish peroxidase (HRP)-conjugated secondary antibody for 1 h at room temperature. Immuno-reactive bands were detected using an enhanced chemiluminescence (ECL) kit, and were quantitatively analyzed with Image J software. GAPDH was used as the internal control.

DNA library construction and high-throughput sequencing

Total RNA was isolated from the two groups of cells (normal PDLCs and PDLCs stretched for 24 h), according to the manufacturer’s instructions. RNA molecules sized 18-30 nt were enriched by PAGE. Then, the 3’ and 5’ adapters were ligated to the RNAs. The ligation products were constructed by reverse transcription polymerase chain reaction (RT-PCR), and PCR products sized 140-160 bp were enriched to generate a cDNA library and sequence using Illumina HiSeqTM2500 (Illumina, Inc., San Diego, CA, USA).

Alignment and identification of miRNAs

Reads obtained from the sequencing machines were further filtered to achieve clean tags. All the clean tags were aligned with small RNAs using the GenBank 209 and Rfam11.0 databases to identify and remove rRNA, scRNA, snoRNA, snRNA, and tRNA. Meanwhile, the clean tags were aligned with the reference genome, and those mapped to exons, introns, and repeat sequences were also removed. Then, the clean tags were searched against miRBase21 to identify known miRNAs. All unannotated tags were aligned with the reference genome. Novel miRNA candidates were identified according to their genome positions and hairpin structures as predicted by Mireap v0.2 software.

Analysis of differentially expressed miRNAs

miRNAs with a fold change ≥ 2 and a P-value < 0.05 were considered to be significantly differentially expressed.

Prediction of miRNA target genes and functional enrichment analysis

Based on the differential expression profile of miRNAs, the candidate target genes were predicted using three software programs (RNAhybrid v. 2.1.2 + svm-light v. 6.01, Miranda v. 3.3a, and TargetScan v. 7.0). The intersection of the results was chosen as the predicted miRNA target genes. GO terms and KEGG pathway annotation were used to further identify the biological functions of the target genes. Genes with a false discovery rate (FDR) < 0.05 were considered to be significantly enriched target candidate genes.

Quantitative RT-PCR (RT-qPCR)

To measure the mRNA level of miRNAs and other genes, total RNA was isolated from PDLCs and rat periodontal ligament tissue using a mirVanaTM miRNA isolation kit (Invitrogen, Carlsbad, CA, USA). Then, 1 μg of total RNA was reverse-transcribed using a Mir-X miRNA First-Strand Synthesis kit and PrimerScript RT Reagent kit with gDNA Eraser (Takara, Tokyo, Japan). SYBR Premix Ex Taq II (Takara, Tokyo, Japan) was used for RT-qPCR analysis, according to the manufacturer’s instructions. The relative expression of mRNA or miRNA was calculated using the 2-ΔΔCt method and normalized to the expression of GAPDH or U6, respectively. All PCRs were performed in triplicate. The miRNA primer sets were purchased from GenePharma (Suzhou, China). And the primer pairs of the relative genes are shown in Table 1.

Table 1.

Primer sequences for RT-qPCR

Gene Forward primer (5’-3’) Reverse primer (5’-3’)
ALP CGAGATACAAGCACTCCCACTTC CTGTTCAGCTCGTACTGCATGTC
BSP CAGGGCAGTAGTGACTCATCC TAGCCCAGTGTTGTAGCAGA
COL1A1 CGATGGATTCCAGTTCGAG TAGGTGATGTTCTGGGAGGC
SPP1 GTGGCCACATGGCTAAACCCT GACTTACTTGGAAGGGTCTGTGG
OCN GGTGCAGCCTTTGTGTCCAA CCTGAAAGCCGATGTGGTCA
RUNX2 AACCCTTAATTTGCACTGGGTCA CAAATTCCAGCAATGTTTGTGCTAC
CAP CTGCGCGCTGCACATGG GCGATGTCGTAGAAGGTGAGCC
CEMP1 GGGCACATCAAGCACTGACAG CCCTTAGGAAGTGGCTGTCCAG
SATB2 GCCCTGGTCTTCTTTCTCCC CGGAAGAGTTGGTTGGCTCT
MSX2 GCCTCGGTCAAGTCGGAAAA CGGCGCGCACTCACTT
TEAD1 CCATTCCAGGGTTTGAGCCT GCTTGGTTGTGCCAATGGAG
YAP1 TCCCAGATGAACGTCACAGC TCATGGCAAAACGAGGGTCA
LATS1 TGAAGAGCGAAGGGAATCTCG TTTAAAAGGTTTTACCCGCATCA
WTIP CCACCTACTGTGTCGTCGTT TCAGAGCTCAGTGACGTGC
GLI2 GCAGGTGTATCCCACGGAAA CTGGCATCCTCCCAGCATAG
GDF5 GACTCCCCAAACTCCTCACTT CACCCAACACAGTGCAGATGA
DVL2 AGTGAGCATGGCGCTGG TCGCTGGTCATGAGGGTAGA
CTGF GTTTGGCCCAGACCCAACTA GGCTCTGCTTCTCTAGCCTG
LIMD1 GAGAGATGGATGCTCACCCG GTAGAGGTTCCCCATGGCCT
WWTR1 TGGACCAAGTACATGAACCACC GACTGGTGATTGGACACGGT
TEAD2 CCCCATCTACTGACCTCCCA GGTTCCACGAAGGCTGAGAA
GAPDH AACAGCGACACCCACTCCTC CATACCAGGAAATGAGCTTGACAA
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OTM

Twelve eight-week-old male Wistar rats weighing 180 to 220 g were used in this study. All procedures involving animals were performed in accordance with the ethical standards of our institution, and were approved by the Institutional Animal Care and Use Committee of Wuhan University. In brief, we used a 0.2 mm nickel-titanium coil spring (TOMY International Inc., Tokyo, Japan) to connect the maxillary left first molar to the incisors. A tension gauge was utilized to calculate the force magnitude in the stretch direction. The non-stretched right upper first molars with no appliance acted as the corresponding control. For each condition in our experiment, six animals were used at each time point. After 3 days, the PDL tissue from each rat’s distal stretched side and the control side was collected for RT-qPCR analysis.

Measurement of OTM

Rats were sacrificed 14 days after orthodontic force application. The maxillary alveolar bone together with three molars on the right and left sides were isolated and submitted to X-ray photography (Bruker Corp., Billerica, MA, USA). Mesial molar movement was determined by the distance between the cemento-enamel junctions of the orthodontic loaded first molars and the second molars. The measurement of OTM was performed using the ruler tool in Adobe Photoshop software (Adobe Inc., San Jose, CA, USA).

Hematoxylin and eosin (H&E) staining

The maxillary alveolar bone and the three molars from each rat were fixed with 4% paraformaldehyde for 24 h at room temperature. They were then decalcified for 3 months using 10% EDTA (pH = 7.4) and then dehydrated and embedded in paraffin. All specimens were cut into 5 μm sections and were then deparaffinized and rehydrated. H&E staining was performed for histological and histomorphometric analysis.

Statistical analysis

Statistical analysis was undertaken by SPSS 16.0 software (IBM, Armonk, NY, USA) or GraphPad Prism 5.0 software. All quantitative data were presented as mean ± standard deviation (SD) of at least three independent experiments. Statistical difference was assessed by student’s t test or one-way analysis of variance (ANOVA) in either two or multiple groups. A P-value < 0.05 was considered statistically significant.

Results

Morphology and characterization of PDLCs

PDLCs derived from periodontal ligament tissue reached confluence in two weeks, exhibiting spindle, fibroblast-like morphology (Figure 1A). Flow cytometry analysis indicated that cultured PDLCs were positive for mesenchymal stem cell markers (CD146, CD90) and negative for CD45 (Figure 1B). ALP and alizarin red staining revealed calcium deposits and mineralized nodule formation in PDLCs after osteogenic induction for 7 days and 21 days, respectively (Figure 1C, 1D). The relative expression of all osteogenic genes was increased during the process of PDLC osteogenesis (Figure 1F). Moreover, staining with Oil Red O showed the formation of lipid clusters in cultured cells (Figure 1E). These results indicate that PDLCs were of mesenchymal origin and had multi-differentiation potential.

Figure 1.

Figure 1

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Morphology and characterization of human periodontal ligament cells (PDLCs). (A) Cultured single cells grew to confluence in two weeks, exhibiting spindle, fibroblast-like morphology. (B) Flow cytometry analysis indicated that cultured PDLCs were positive for mesenchymal stem cell markers CD146 (42.64%) and CD90 (99.08%), and negative for CD45 (0.51%). (C) ALP and (D) Alizarin red staining indicated the presence of calcium deposits and mineralized nodules in PDLCs after osteogenic induction for 7 days and 21 days, respectively. (E) Lipid accumulation was detected by Oil Red O staining of PDLCs cultured for 14 days with adipogenic medium. (F) Relative expression levels of the osteogenic/cementoblastic markers ALP, RUNX2, COL1A1, OCN and CEMP1 were upregulated during the process of osteogenesis of PDLCs (P < 0.05). *P < 0.05 vs. control.

Mechanical stretch-induced osteogenic/cementoblastic differentiation of PDLCs

PDLCs at a density of 1 × 105 were seeded onto a six-well plate with a flexible silicon membrane. After mechanical loading for 24 h, the stretched cells showed a higher expression level of osteogenic/cementoblastic genes and proteins compared with the cells in the control group (Figure 2A, 2B). This indicates that the osteoblastic/cementoblastic differentiation of PDLCs was increased when mechanical force was imposed. See the original Western blot images in Figures S1 and S2.

Figure 2.

Figure 2

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Effect of mechanical stretch on osteogenic/cementoblastic differentiation of cultured human periodontal ligament cells (PDLCs). A. The osteogenic/cementoblastic markers RUNX2, SATB2, MSX2, BSP, SPP1, OCN, CAP, and CEMP1 were detected by RT-qPCR analysis and were found to be upregulated in the tension group (P < 0.05). B. Western blot analysis revealed that stretched PDLCs had higher expression levels of osteogenic/cementoblastic proteins. *P < 0.05 vs. control.

Differentially expressed miRNAs

To analyze the variance in miRNAs during mechanical force loading, we compared the known miRNA expressions between stretched and non-stretched PDLCs to identify the differentially expressed miRNAs. A heat map and cluster analysis was applied to visualize the miRNAs differentially expressed between the two groups. The analyses showed that 47 known miRNAs were differentially expressed, of which 31 were upregulated and 16 were downregulated in the stretched PDLCs compared with the normal PDLCs (Figure 3A). The overall distribution of the differentially expressed miRNAs is displayed in a scatter plot (Figure 3B).

Figure 3.

Figure 3

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Hierarchical clustering analysis, scatter plot, and in vitro validation of differentially expressed miRNAs in stretched and non-stretched periodontal ligament cells (PDLCs). (A) Cluster analysis of 47 differentially expressed miRNAs. Red color represents a relative high expression level. Green color shows a relative low expression level. In the stretched group, 31 miRNAs were significantly upregulated and 16 were downregulated compared with the control group. (B) miRNA scatter plot. Red dots represent significantly upregulated miRNAs in the stretched PDLCs compared with the control cells. Green dots show significantly downregulated miRNAs. Blue dots represent equally-expressed miRNAs. (C) RT-qPCR was applied to verify the expression levels of differentially expressed miRNAs. Consistent with (D) the RNA-Seq results, miR-218-5p, miR-138-5p, miR-221-3p, and miR-132-3p were remarkably upregulated, while miR-133a-3p, miR-145-3p, miR-143-5p, miR-486-3p, and miR-210-3p were significantly downregulated in the stretched group (P < 0.05). *P < 0.05 vs. control.

Validation of differentially expressed miRNAs in vitro by RT-qPCR

To verify the results obtained from our RNA-Seq experiment, nine osteogenesis-related miRNAs were chosen for validation by RT-qPCR. The function and the validated osteogenesis-related targets of those nine miRNAs were reviewed according to the published literature by searching PubMed (Table S1). Consistent with the RNA-Seq results, miR-218-5p, miR-138-5p, miR-221-3p, and miR-132-3p were remarkably upregulated, while miR-133a-3p, miR-145-3p, miR-143-5p, miR-486-3p, and miR-210-3p were significantly downregulated in the stretched group (Figure 3C, 3D). These results indicated that mechanical force could induce changes in the expression profile of miRNAs in PDLCs.

Validation of miRNA expression in rats

To further confirm the expression levels of several conserved miRNAs after mechanical force loading, we established a rat model for OTM (Figure 4A). X-ray photographs and quantitative analysis (Figure 4B, 4C) indicated that OTM was evident after 14 days of force application at the maxillary first molar. HE staining revealed that there was more bone tissue formation at the tension site compared with the control site (Figure 4D). Next, we isolated PDL tissue from both the stretched site and the control site after 3 days of orthodontic force loading. RT-qPCR was performed to detect the expression levels of six conserved miRNAs, including miR-221-3p, miR-138-5p, miR-132-3p, miR-210-3p, miR-133a-3p, and miR-133a-5p. The results were consistent with our RNA-Seq and in vitro experiment (Figure 4E).

Figure 4.

Figure 4

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Rat orthodontic tooth movement (OTM) model and validation of differentially expressed miRNAs in vivo. A. A rat model for OTM. B. X-ray photography of orthodontic force induced tooth movement in rats after 14 days. Black arrow: force loading direction; scale bar: 500 μm. C. Tooth movement measurement between cemento-enamel junctions of orthodontic loaded first molars and second molars for the tension force-stimulated site (TS) and the control site (CS). The results showed significantly more tooth movement in the tension site (P < 0.05). D. HE staining revealed more bone tissue volume in the stretched group (right) compared with the control group (left). Scale bar: 100 μm; r: root; p: periodontium; b: alveolar bone. E. Six conserved miRNAs, including miR-221-3p, miR-138-5p, miR-132-3p, miR-210-3p, miR-133a-3p, and miR-133a-5p, were detected by RT-qPCR analysis. The results were consistent with our RNA-Seq and in vitro experiment (P < 0.05). *P < 0.05 vs. control.

Identification of differentially expressed miRNAs controlled by osteoblast-specific transcription factors

RUNX2, osterix (OSX), SATB2, MSX2, and DLX5 are well-known transcription factors that play a vital role in osteogenic differentiation. In the present study, the expressions of RUNX2, SATB2, and MSX2 were upregulated in PDLCs after mechanical loading (Figure 2A). In order to verify the regulatory effect of miRNAs on these transcription factors, we used three software programs to search for miRNA targeted osteogenesis-related transcription factors. As shown in Table 2, several miRNAs were predicted to target the above-mentioned transcription factors.

Table 2.

Osteogenesis-related transcription factor targeted by differentially expressed miRNAs

Transcription factor Function miRNA
RUNX2 Osteogenesis miR-218-5p, miR-625-3p, miR-150-5p, miR-2682-5p, miR-7-1-3p, miR-221-3p, miR-133a-5p
Osterix Osteogenesis miR-486-3p, miR-145-3p, miR-143-5p, miR-133a-3p
SATB2 Osteogenesis miR-218-5p, miR-1-3p, miR-31-5p, miR-376a-3p
MSX2 Osteogenesis miR-7-1-3p, miR-708-5p, miR-942-3p
DLX5 Osteogenesis miR-376a-3p, miR-942-3p
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Functional analysis of mechanical stretch-responsive miRNAs

GO is an international standardized gene function classification system that comprehensively describes the properties of genes. To explore the potential functions of differentially expressed miRNAs in stretched PDLCs, we performed GO enrichment analysis to identify the functions significantly associated with the predicted target genes of the changed miRNAs. The most enriched GO terms in the biological process, cellular component, and molecular function domains are shown in Figure 5A.

Figure 5.

Figure 5

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Functional analysis of mechanical stretch-responsive miRNAs. A. GO enrichment analysis indicated that target genes of differentially expressed miRNAs had a broad range of functions, including biological process, cellular components, and molecular function. B. Top 20 enriched pathways are shown in the scatter plot. The horizontal axis represents rich factor. The vertical axis shows the specific 20 enriched pathways.

KEGG analysis can identify the main pathways in which the target gene candidates of the changed miRNAs could be involved. The top 20 enriched pathways of the target genes are displayed in Figure 5B. Among them, the MAPK signaling pathway, cAMP signaling pathway, and Hippo signaling pathway were found to be involved in the regulation of osteogenesis.

Hippo signaling pathway is differentially expressed in stretched and non-stretched PDLCs

The Hippo signaling pathway is involved in mechano-transduction and osteogenesis. According to our RNA-Seq results, the Hippo signaling pathway was obviously enriched in biological function analysis of target genes associated with differentially expressed miRNAs. Furthermore, several key components of the Hippo signaling pathway, such as YAP1, WWTR1, TEAD2, CTGF, DVL2, GDF5, GLI2, LIMD1, WTIP, LATS1, and TEAD1, were predicted to be target genes of the differentially expressed miRNAs (Figure 6A); the upregulation of these components in PDLCs submitted to mechanical force loading was validated by RT-qPCR (Figure 6B). Among them, YAP1, WWTR1, TEAD2, CTGF, DVL2, and GDF5 were found to be positive regulators of osteogenesis.

Figure 6.

Figure 6

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Differentially expressed miRNAs targeted core components of the Hippo signaling pathway. A. Regulatory networks of known differentially expressed miRNAs and the components of the Hippo signaling pathway. The red and green solid circles represent upregulated and downregulated miRNAs, respectively. The light grey box represents the components of the Hippo signaling pathway. B. RT-qPCR confirmed that the expression levels of several key components of the Hippo signaling pathway, including YAP1, WWTR1, TEAD2, CTGF, DVL2, GDF5, GLI2, LIMD1, WTIP, LATS1 and TEAD1, were upregulated in PDLCs submitted to mechanical force loading (P < 0.05). *P < 0.05 vs. control.

Discussion

OTM can be induced by the constant application of orthodontic force. PDL and alveolar bone cells can perceive and transfer the mechanical force into molecular events. As a result, bone formation and resorption occur at the tension and compressive sites, respectively. To date, the underlying mechanisms in this process have not been fully elucidated. In our study, we first developed an in vitro force-loading model for PDLCs to mimic the process of OTM. Then, we applied high-throughput sequencing to determine the expression profile of miRNAs in stretched and non-stretched PDLCs. We also performed prediction and functional analysis of target genes. Our findings suggest that multiple miRNAs may be involved in mechanical force-induced osteoblastic/cementoblastic differentiation of PDLCs.

A number of in vitro studies have demonstrated that PDLCs are mechanosensitive. Cyclic stretch can induce several biological changes in PDLCs, such as cell realignment, cytoskeleton organization, apoptosis, and angiogenesis [21-24]. In addition, previous studies have also reported that cyclic tension can promote the osteogenic differentiation of PDLCs. An in vitro study showed that cyclic tension resulted in increased osteogenic gene expression of PDLCs [25]. Another study reported that mechanical loading may control osteoblastic/cementoblastic differentiation of PDLCs mediated by angiotensin II signaling [10]. In this study, we observed higher expression of osteoblastic/cementoblastic genes and proteins in PDLCs after applying mechanical tension force loading, which was consistent with previous studies.

miRNAs are a type of small and non-coding RNA that regulate gene expression at the post-transcriptional level. Several previous studies have reported the expression profile of miRNAs in PDLCs after force stimulation. One study detected the expression levels of miRNAs in PDLCs treated with fluid shear stress using microRNA arrays [16]. Another study explored miRNA expression differences between normal PDLCs and PDLCs that underwent 72 hours of 12% cyclic tension force stimulation. The above study screened 3100 miRNAs, of which 17 miRNAs were upregulated and 15 miRNAs were downregulated [26]. In our study, we cultured PDLCs under 10% equibiaxial strain at 0.1 Hz for 24 hours, and applied high-throughput sequencing to identify the expression profile of miRNAs. According to our results, the expression levels of 47 miRNAs were found to be altered after stretching, of which with 31 were upregulated and 16 were downregulated. The differences in results obtained by other scholars compared to our results may be attributed primarily to the analytical methods. Compared with microarray analysis, high-throughput sequencing can identify new miRNAs that have previously been unreported, including those evident at low expression levels. Additionally, the effect of mechanical stimulation on cells may be related to the device, magnitude, duration, and frequency. Next, we selected nine osteogenesis-related miRNAs to be further validated by RT-qPCR. The results of our RT-qPCR analysis were consistent with the RNA-Seq experiment. The relative function and validated targets of those nine miRNAs in the process of osteogenesis were also reviewed in this study (see Table S1). However, further studies must be performed to determine whether these osteo-miRNAs are also involved in the process of osteogenic/cementoblastic differentiation of stretched PDLCs.

Osteogenesis is a well-organized process involving various factors. In recent years, transcription factors have been found to combine with miRNAs to regulate osteogenic differentiation of MSCs. RUNX2, an early and primary osteogenic transcription factor, is responsible for the activation of osteoblast marker genes, including collagen type I alpha 1 chain (COL1A1), SPP1, and bone sialoprotein (BSP) [27]. Additionally, miR-133a was reported to be a key negative regulator of osteogenic differentiation of vascular smooth muscle cells, and this effect appeared to be mediated by targeting RUNX2 [28]. As a downstream gene of RUNX2, OSX is specifically expressed during osteoblast differentiation and is critical for bone formation [29]. It has been shown that miR-143 and miR-145 can repress osteogenic differentiation through the regulation of OSX expression, forming a feedback loop with Kruppel-like factor 4 (KLF4) and OSX in odontoblasts [30]. DLX5 is a bone-inducing transcription factor that is expressed in the later stages of osteoblast differentiation [31]. A previous study showed that miR-203 and miR-320b can negatively regulate BMP2-induced osteoblast differentiation by suppressing DLX5, which in turn inhibits RUNX2 and OSX [32]. MSX2 has been found to act as an upstream factor of RUNX2 [33]. miRNA array analysis has revealed that miR-124a and miR-181a are important regulatory factors for osteoblastic differentiation of mouse-induced pluripotent stem cells; both can directly target DLX5 and MSX2 [34]. SATB2 is an important determinant of osteoblastic differentiation and can enhance the activity of RUNX2 and activate transcription factor 4 (ATF4) [35]. To date, several miRNAs have been reported to regulate osteoblastic differentiation by directly binding to the 3’-UTR of SATB2 mRNA [36,37]. In summary, the above evidence suggests that regulation of transcription factors controlled by miRNAs is a crucial mechanism underlying osteogenic differentiation. In the present study, RUNX2, OSX, DLX5, MSX2, and SATB2 were predicted to be the targets for differentially expressed miRNAs in PDLCs after force stimulation. Furthermore, the expression levels of RUNX2, MSX2, and SATB2 were found to be upregulated in tension force loading PDLCs. Whether the differentially expressed miRNAs regulate the mechanical stretch-induced osteogenesis of PDLCs by targeting these osteogenic specific transcription factors requires further study.

Increasing evidence indicates that the Hippo signaling pathway is regulated by mechanical stress. Several core components that are reported to be positive regulators of osteogenesis are involved in the Hippo signaling pathway. Yes-associated protein (YAP) and transcriptional coactivator with PDZ-binding motif (TAZ) are key downstream effectors of the Hippo signaling pathway, and have potential to function in mechano-transduction and osteogenic differentiation [38]. Cyclic stretch enhances the osteogenesis of PDLCs via YAP-activation [39], and TAZ is indicated as a key mediator, promoting ADSC commitment to the osteoblast lineage [40]. TEA domain transcription factor 2 (TEAD2) is located in the nucleus, where it can form a complex with YAP and co-activate the expression of multiple genes involved in cell fate determination, proliferation, and survival [41]. Connective tissue growth factor (CTGF) is a target gene of YAP; its expression is known to be induced by mechanical stimulation and is involved in osteogenic differentiation of mechanical stretched-associated ADSCs [42]. Growth differentiation factor 5 (GDF5) and Dishevelled 2 (DVL2) are another two components involved in the Hippo signaling pathway; they have been shown to be involved in the osteogenic differentiation of ligamentum flavum (LF)-derived stem cells (LFSCs) and synovial fibroblasts, respectively [43,44]. Recently, a number of studies have demonstrated an association between miRNA and the Hippo signaling pathway during osteogenesis of human MSCs (hMSCs). One study showed that miR-135b-5p positively regulates osteogenic differentiation of hMSCs by controlling the expression levels of LATS1 and MOB1B, key negative regulators of the Hippo signaling pathway, resulting in subsequent activation of the Hippo signaling pathway [45]. In our study, YAP1, WWTR1, TEAD2, CTGF, DVL2, GDF5, GLI2, LIMD1, WTIP, LATS1, and TEAD1 were determined to be the target genes of differentially expressed miRNAs, and also were confirmed to be upregulated in stretched PDLCs. Therefore, we speculate that the changed miRNAs can regulate the mechanical stretch-induced osteogenic/cementoblastic differentiation of PDLCs via components of the Hippo signaling pathway.

In conclusion, we first identified and characterized the expression profile of miRNAs in PDLCs after mechanical force loading using high-throughput sequencing. Furthermore, we explored a regulatory network between differentially expressed miRNAs and transcription factors to identify the potential mechanisms underlying osteogenic/cementoblastic differentiation of stretched PDLCs. This study provides a basis for further insight into the molecular mechanisms involved in bone remolding during OTM.

Acknowledgements

This work was financially supported by the National Natural Science Foundation of China (No. 81500888 and 81571011), the Youth Medical Experts’ Project of Wuhan, Research Fund for the Doctoral Program of Higher Education (20120141130009) and Wuhan Applied Basic Research Project (No. 201606010101004). We are very thankful for the technical assistance and bioinformatics analysis provided by Gene Denovo.

Disclosure of conflict of interest

None.

Supporting Information

ajtr0011-3398-f7.pdf (391.3KB, pdf)

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