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. 2023 Jan 28;6(2):e1247. doi: 10.1002/jsp2.1247

DLX5 regulates the osteogenic differentiation of spinal ligaments cells derived from ossification of the posterior longitudinal ligament patients via NOTCH signaling

Tao Tang 1,2, Zhengya Zhu 1, Zhongyuan He 1,2, Fuan Wang 1,2, Hongkun Chen 1, Shengkai Liu 2, Mingbin Zhan 2, Jianmin Wang 1, Wei Tian 3, Dafu Chen 3, Xinbao Wu 3, Xizhe Liu 2,✉, Zhiyu Zhou 1,2,✉, Shaoyu Liu 1,2
PMCID: PMC10285757  PMID: 37361333

Abstract

Background

Ossification of the posterior longitudinal ligaments (OPLL) is common disorder characterized by heterotopic ossification of the spinal ligaments. Mechanical stimulation (MS) plays an important role in OPLL. DLX5 is an essential transcription factor required for osteoblast differentiation. However, the role of DLX5 during in OPLL is unclear. This study aims to investigate whether DLX5 is associated with OPLL progression under MS.

Methods

Stretch stimulation was applied to spinal ligaments cells derived from OPLL (OPLL cells) and non‐OPLL (non‐OPLL cells) patients. Expression of DLX5 and osteogenesis‐related genes were determined by quantitative real‐time polymerase chain reaction and Western blot. The osteogenic differentiation ability of the cells was measured using alkaline phosphatase (ALP) staining and alizarin red staining. The protein expression of DLX5 in the tissues and the nuclear translocation of NOTCH intracellular domain (NICD) was examined by immunofluorescence.

Results

Compared with non‐OPLL cells, OPLL cells expressed higher levels of DLX5 in vitro and vivo (p < 0.01). Upregulated expression of DLX5 and osteogenesis‐related genes (OSX, RUNX2, and OCN) were observed in OPLL cells induced with stretch stimulation and osteogenic medium, whereas there was no change in the non‐OPLL cells (p < 0.01). Cytoplasmic NICD protein translocated from the cytoplasm to the nucleus inducing DLX5 under stretch stimulation, which was reduced by the NOTCH signaling inhibitors (DAPT) (p < 0.01).

Conclusions

These data suggest that DLX5 play a critical role in MS‐induced progression of OPLL through NOTCH signaling, which provides a new insight into the pathogenesis of OPLL.

Keywords: cyclic stretch, DLX5, mechanical stimulation, NOTCH signaling, OPLL, osteogenic differentiation

The mechanism of NOTCH signaling and regulating DLX‐5 medited osteogenic differentiation

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1. INTRODUCTION

Ossification of the posterior longitudinal ligament (OPLL) is a common spinal disease with ectopic bone formation in the posterior longitudinal ligament (PLL). 1 It is often associated with severe compression of the spinal cord and nerve roots, resulting in symptoms of paralysis and other myelopathy. 2 Surgical intervention is required when symptoms are severe with evidence of compression to the nerve root or spinal cord. 3 At present, spinal cord decompression has become an established therapy option for patients with advanced OPLL. 4

OPLL is a multifactorial degenerative disease of the spine, caused by both environmental and genetic risk factors. 5 The environmental factors include diet, metabolism, age, and physical strain stimulation. 6 Anterior cervical discectomy with fix multiple cervical segments has become an effective method for preventing postoperative ossification progression in OPLL. 7 , 8 Several studies have shown that mechanical stimulation (MS) enhance osteogenic differentiation of OPLL spinal ligament cells(OPLL cells). 9 , 10 , 11 , 12 , 13 Excessive mechanical loading of rat caudal vertebrae producing tensile strain and resulting in cartilage formation in spinal ligaments. 14 However, the mechanism of MS in OPLL osteogenic differentiation has been not fully elucidated.

Distal‐Less Homeobox5 (DLX5) is required for the development and maturation of the olfactory system, cranial neural crest, and skeletal system. 15 , 16 , 17 Additionally, DLX5 is largely involved in osteoblast differentiation and MS‐induced osteogenesis. 18 DLX5 also acts as a mechanical sensitive gene closely related to MS. 19 , 20 , 21 , 22 Since, it has been reported that mechanical stress may activate DLX5 for the bone formation, we assume that DLX5 has the same effect in OPLL cells.

In this study, we explored the possible mechanisms by which DLX5 might regulated osteogenic differentiation in OPLL cells. Our study will provide a novel insight into molecular mechanisms of the pathogenesis of OPLL.

2. METHODS

2.1. Patients and ethics

This study was approved by the Ethics Committee of the Seventh Affiliated Hospital of Sun Yat‐sen University (Certificate No. 2020SYSUSH‐055), and informed consent was obtained from each patient. Eleven OPLL patients and 11 non‐OPLL patient controls were selected for this study. Patients were diagnosed and staged based on clinical history and physical exam and computed tomography studies. PLL specimens from all patients were collected during cervical spine surgical decompression.

2.2. Cell culture and osteogenic differentiation

PLL tissues were washed three times with phosphate‐buffered saline (PBS) and were carefully dissected to avoid surrounding tissue contamination. The tissues were cut and ground into small pieces and digested by enzymes following the protocol. The cell suspension was collected, neutralized and added to DMEM medium (Gibco, Life Technologies) containing 10% fetal bovine serum (FBS, Gibco, Life Technologies). Cells were cultured in a humidified incubator at 37°C and 5% carbon dioxide. Cells were detached using 0.25% trypsin EDTA (Gibco, Life Technologies) for passaging and third generation were used for subsequent experiments. OPLL cells and non‐OPLL cells were identified by vimentin staining (Figure 1A).

FIGURE 1.

FIGURE 1

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(A) Vimentin was visualized by immunostaining (IF) staining in spinal ligament cells. The scale bars = 20 μm. (B and C) Quantitative polymerase chain reaction and western blot analysis showing the mRNA and protein expression level of DLX5 in ossification of the posterior longitudinal ligament (OPLL) and non‐OPLL cells, data was presented as mean fold changes in OPLL cells as compared to control cells, n = 3, two‐tailed student's t‐test. (D and E) IF staining and quantification assay of DLX5 in ligament tissues. Scale bar = 500 μm, n = 3, two‐tailed student's t‐test. All data were presented as means ± SEMs. Immunofluorescence staining of DLX5 was performed in the posterior longitudinal ligament tissues of the OPLL and N‐OPLL groups (figure1E). Although the local immunofluorescence intensity was higher in the N‐OPLL group than in the OPLL group within the orange box, the overall immunofluorescence intensity was higher in the OPLL group than in the N‐OPLL group. In other words, Image J software was used to quantify the immunofluorescence intensity not in the orange box but in the whole image (figure1D). *p < 0.05; **p < 0.01; ***p < 0.001

For osteogenic induction culture, cells were cultured in an osteogenic medium for 7 or 14 days. Subsequently, alkaline phosphatase (ALP) staining and alizarin red staining were performed to examine the osteogenic differentiation.

2.3. Cyclic stretch

PLL cells were subjected to stretch according to previously described methods with minor modifications. 23 Briefly, cells were seeded on an elastic silicone membrane (Dow Corning) coated with 5 μg/cm2 collagen I (Thermo Fisher Scientific) at a density of 1 × 104 cells/cm2. After reaching about 70% confluence, the culture mediums were changed to 1% FBS DMEM for 24 h. Cells were then subjected to a cyclic stretch (10% elongation, 0.5 Hz) at 37°C in a humidified atmosphere of 5% CO2.

2.4. RNA extraction and real‐time quantitative polymerase chain reaction

RNA extraction was performed using the RNAeasyTM animal RNA isolation Kit (Beyotime) following the manufacturer's protocol. RNA was converted to cDNA by PrimeScript™ RT Master Mix (TaKaRa). Real‐time quantitative polymerase chain reaction (RT‐qPCR) was performed using qPCR Mix (Thermo Fisher Scientific) in a CXF‐96 Real‐Time System (Bio‐Rad). The reaction mixture contained the following components in a total volume of 10 μl: 5 μl Fast SYBR green master mix, 2 μl RNase‐free dH2O, 2 μl cDNA, and 0.5 μl of each primer (Table S1). Relative expression was calculated using the 2−ΔΔCt and gene expression levels were normalized to GAPDH.

2.5. Western blotting

Cells were lysed with RIPA (Thermo Fisher Scientific) buffer supplemented with protease inhibitor cocktail (Thermo Fisher Scientific) and PMSF (Boster). Protein extracts were resolved by electrophoresis on 4%–12% Bis‐Tris NuPAGE gel (Invitrogen) and transferred to PVDF membranes (Invitrogen). The membranes were blocked for 1 h with 5% nonfat milk, incubated with primary antibodies overnight at 4°C overnight and HRP‐conjugated goat anti‐rabbit antibodies for 1 h at room temperature (Abcam). The antigen‐antibody complexes were visualized using the ECL reagent (EpiZyme). The immunoreactive bands were quantified using ImageJ software and normalized to the corresponding GAPDH bands.

2.6. Immunofluorescence staining

PLL tissues were fixed in 4% paraformaldehyde for 24 h and washed three times with PBS. After decalcification and dehydration, the samples were embedded in OCT compound and 10 μm cryosections were made with a cryostat. Sections were permeabilized and blocked in TBST (Biosharp) containing 0.3% Triton X‐100 (Sigma) and 5% bovine serum albumin (BioFroxx). Sections were incubated with an anti‐DLX5 antibody (Abcam, 1:100) overnight at 4°C and goat anti‐rabbit secondary antibody (Abcam, 1:300) for 1 h at room temperature. Nuclei were counterstained with 4,6‐diamidino‐2‐phenylindole (DAPI, Abcam) for 5 min. Stains were visualized using a fluorescence microscope (Leica).

For immunostaining (IF) staining in cultured cells, after fixation, permeabilization and blocking, cells were incubated with primary antibodies overnight and followed by corresponding secondary antibodies for 1 h. The following IF protocol was the same as that described above.

2.7. Transfection and γ‐secretase inhibition

The ShRNA sequences targeting human DLX5 were cloned into hU6‐MCS‐CBh‐gcGFP‐IRES‐puromycin vector (Genechem). Their sequence were as follows respectively: shDLX5: 5′‐CTCAGGAATCGCCAACTTT‐3′; scrambled control: 5′‐TTCTCCGAACGTGTCAC‐GT‐3′. Lentiviruses were collected 48 h after transfection, mixed with polybrene (8 μg/ml, Sigma Aldrich) and used for infection. Transfection efficiency was detected by GFP expression under fluorescence microscopy and efficiency of DLX5 knockdown was determined by qPCR.

To inhibit NOTCH signaling in vitro, cells were pretreated with the 10 μM γ‐secretase inhibitors: N‐[N‐(3,5‐difluorophenacetyl)‐l‐alanyl]‐(S)‐phenylglycine‐t‐butyl ester (DAPT; Sigma Aldrich) at 30 min before MS.

2.8. Statistical analysis

SPSS version 20.0 (SPSS) was used for statistical analysis. All values are presented as means ± standard error of the means. The Shapiro–Wilk normality test was performed to evaluate the normality of the data distribution. Statistical significance (p ≤ 0.05) was analyzed by Student's t‐test (two groups) or one‐way analysis of variance (ANOVA) (more than two groups). Mann–Whitney U test was performed for non‐normally distribution data.

3. RESULTS

3.1. DLX5 is highly expressed both in vitro and vivo

IF staining of vimentin was used to determine the cell phenotype and the results showed that most of spinal ligament cells were fibroblasts (Figure 1A). To confirm the expression of DLX5 in ligament cells, mRNA and protein levels of DLX5 by RT‐PCR and western blot. DLX5 expression was significantly increased in OPLL cells compared to controls (Figure 1B,C). These results were consistent with the expression of DLX5 detected by IF staining (Figure 1D,E). Overall, our results demonstrated that DLX5 was highly expressed in OPLL cells and tissues.

3.2. High expression of DLX5 and high osteogenic potential in osteogenic induction

To confirm osteogenic capacity, cells from OPLL and non‐OPLL group were cultured in osteogenic medium. Over 7 or 14 days' culture period, osteogenic differentiation was significantly enhanced by ALP staining and alizarin red staining in OPLL group (Figure 2A–D). The increasing of DLX5 expression was observed at mRNA levels after osteogenic induction for 7 days (Figure 2E). These results were consistent with the expression of DLX5 detected by IF staining (Figure 2F,G). In conclusion, OPLL cells had a higher expression level of DLX5 and osteogenic potential compared to the non‐OPLL cells during induction of osteogenic differentiation.

FIGURE 2.

FIGURE 2

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High expression of DLX5 and high osteogenic potential in ossification of the posterior longitudinal ligament cells during induction of osteogenic differentiation. (A–D) The osteogenic potential was analyzed using alkaline phosphatase (ALP) staining and alizarin red staining after osteogenic induction for 7 or 14 days, n = 3, two‐tailed Student's t‐test, scale bars = 200 μm. DMEM, DMEM medium; OM, osteogenic medium. (E) Quantitative polymerase chain reaction analysis showing the expression of DLX5 after 7 days' osteogenic induction in different groups, data showing fold changes compared with the respective controls (DMEM group), n = 3, one‐way ANOVA analysis. (F and G) Representative images show immunostaining and the quantification of DLX5 positive cells after 7 days, n = 3, two‐tailed student's t‐test. Scale bar = 50 μm. All data were presented as means ± SEMs. *p < 0.05; **p < 0.01; ***p < 0.001

3.3. DLX5 expression is increased by stretch stimulation

To determine the effect of stretch stimulation on DLX5 expression, we detected its expression levels from ligament cells in culture in response to stretch stimulation. In OPLL cells, significantly increased DLX5 was observed following stretch from 3 to 9 h compared to static group (0 h), whereas non‐OPLL group exhibited no significant change compared to corresponding group (Figure 3A,B). The increased DLX5 expression in the OPLL cells was confirmed by IF staining using an antibody against DLX5 (Figure 3C,D). To examined if the effects of osteogenic differentiation could be induced by stretch stimulation, the expression of osteogenesis‐related genes was detected by qPCR and western blot. The 9 h group had the highest expression levels of OSX, RUXN2 and OCN compared with other groups (Figure 3E,F). Above all, these results indicated that cyclic stretch induced DLX5 expression and osteogenic differentiation in OPLL cells. We then used selected 9 h of stretch stimulation for further study.

FIGURE 3.

FIGURE 3

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Cyclic stretch induced DLX5 expression and osteogenic differentiation in ossification of the posterior longitudinal ligament (OPLL) cells. (A and B) Quantitative polymerase chain reaction (qPCR) and western blot analysis detecting the mRNA and protein expression level of DLX5 in OPLL and non‐OPLL cells after stretch for 7 days, data was presented as mean fold changes in stretch group as compared to control group, n = 3, one‐way ANOVA analysis. (C and D) Representative images show immunostaining and the quantification of DLX5 positive cells after stretch for 7 days, n = 3, two‐tailed student's t‐test. Scale bar = 50 μm. (E and F) qPCR and western blot analysis detecting the mRNA and protein expression level of osteogenic genes after stretch for 7 days both in OPLL and non‐OPLL cells. Data showing fold changes compared with the respective controls, n = 3, one‐way ANOVA analysis. All data were presented as means ± SEMs. *p < 0.05; **p < 0.01; ***p < 0.001

3.4. Knockdown of DLX5 inhibited osteogenic differentiation

To study the effect of DLX5 in osteogenic differentiation, DLX5 expression was knocked down in OPLL cells using lentiviral‐mediated shRNA methods (Figure 4A–C). DLX5 expression decreased following transfection group compared to scramble control (Figure 4D). The expression of osteogenic genes including OSX, RUNX2, and OCN were down‐regulated, and upregulation of these genes enhanced by stretch was robustly blunted by DLX5 knockdown (Figure 4E). Furthermore, the activity of ALP and alizarin red staining in DLX5 knockdown group was significantly decreased after different days of stretch stimulation (Figure 4F,G). These results suggest that stretch‐induced osteogenic differentiation in OPLL cells was inhibited after knockdown of DLX5 expression.

FIGURE 4.

FIGURE 4

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Knockdown of DLX5 inhibited the osteogenic differentiation. (A) Construction of lentivirus vector. (B–D) Transfection efficiency and knockdown efficiency were confirmed by GFP expression and quantitative polymerase chain reaction (qPCR). The scale bars = 50 μm. (E) qPCR analysis detecting the mRNA expression level of OSX, RUNX2, and OCN of different groups in ossification of the posterior longitudinal ligament cells after 7 days of stretch, data was presented as mean fold changes compared to control static group, n = 3, one‐way ANOVA analysis. (F and G) In the DLX5 knockdown experiment, the osteogenic potential was analyzed using ALP staining and alizarin red staining after stretch stimulation for different days, n = 3, two‐tailed Student's t‐test, scale bars = 200 μm. All data were presented as means ± SEMs. *p < 0.05; **p < 0.01; ***p < 0.001

3.5. DLX5 expression is regulated by NOTCH signaling

Previously, it has been reported that DLX5 expression was activated by NOTCH signaling during MSCs osteoblastic differentiation. To assess if NOTCH signaling is essential for the expression of DLX5, NOTCH signaling was inhibited using the gamma‐secretase inhibitor DAPT before stretch stimulation. In this study, DAPT treatment inhibited the expression of DLX5 in OPLL cells (Figure 5A,B). The DLX5 expression was consistent with the expression of DLX5 detected by IF staining (Figure 5C,D). Besides, no statistically significant difference in NOTCH‐related genes, including ligands JAG1, DLL1, and DLL3, receptors NOTCH1, NOTCH2, and NOTCH4, and transcriptional targets HES1, HES5, and HEY2, were noted between transfection group and scramble control, except for NOTCH3 (Figure 5E). The relationship between DLX5 and NOTCH3 needs to be studied further. These results indicated that DLX5 was regulated by NOTCH signaling, while the NOTCH signaling were hardly affected by knockdown of DLX5.

FIGURE 5.

FIGURE 5

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DLX5 expression is regulated by NOTCH signaling. (A and B) Quantitative polymerase chain reaction and western blot analysis of the DLX5 expression treated with DAPT or DMSO vehicle after 7 days of stretch, data was presented as mean fold changes compared to vehicle control, n = 3, two‐tailed Student's t‐test. (C and D) Representative images show immunostaining and the quantification of DLX5 positive cells after stretch for 7 days, scale bar = 50 μm, n = 3, two‐tailed Student's t‐test. Scale bars = 50 μm. (E) Relative expression of NOTCH ligands (JAG1, DLL1, and DLL3), receptors (NOTCH1, NOTCH2, and NOTCH4), and transcriptional targets (HES1, HES5, and HEY2) in shDLX5 group compared to scramble control after stretch for 7 days, n = 3, two‐tailed Student's t‐test. All data were presented as means ± SEMs. *p < 0.05; **p < 0.01; ***p < 0.001

3.6. NOTCH signaling regulates DLX5‐medited osteogenic differentiation

First, NOTCH signaling activation was examined by expression NOTCH intracellular domain (NICD), an active form of NOTCH and NOTCH transcription factors HES1. The increasing of NICD and HES1 expression was observed at protein levels after stretch stimulation (Figure 6A). In response to MS, NICD translocated from the cytoplasm to the nucleus in IF assays which inhibited by DAPT treatment (Figure 6B). We then investigated whether NOTCH regulates DLX5‐medited osteogenic differentiation during stretch stimulation. After knock down of DLX5, DAPT treatment significantly decreased the OSX, RUNX2, and OCN expression compared to scramble group (Figure 6C). The activity of ALP and alizarin red staining was significantly inhibited after 7 of 14 days of stimulation (Figure 6D,E). In conclusion, NOTCH signaling and DLX5 had synergistic effects on regulating osteogenic differentiation in OPLL cells.

FIGURE 6.

FIGURE 6

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DLX5‐medited osteogenic differentiation was regulated by NOTCH signaling. (A) Western blot analysis detecting the protein expression level of NOTCH intracellular domain (NICD) and HES1 after stretch stimulation for 7 days in ossification of the posterior longitudinal ligament cells, data was presented as mean fold changes in stretch group as compared to control group, n = 3, two‐tailed Student's t‐test. (B) Immunostaining staining shows nuclear localization of NICD after stimulation for 7 days, n = 3, two‐tailed Student's t‐test. Scale bars = 20 μm. (C) The quantitative polymerase chain reaction analysis showing the expression level of OSX, RUNX2, and OCN in different groups after stimulation for 7 days. Data showing respective expression fold changes compared to DMSO vehicle, n = 3, one‐way ANOVA analysis. (D and E) The osteogenic potential was analyzed using ALP staining and alizarin red staining with DAPT treatment or knock down of DLX5 after stretch stimulation for 7 or 14 days, n = 3, one‐way ANOVA analysis, scale bars = 200 μm. All data were presented as means ± SEMs. *p < 0.05; **p < 0.01; ***p < 0.001

4. DISCUSSION

OPLL is a common spinal disease with ectopic bone formation in the PLL and the pathogenesis of OPLL has not been defined. 3 Notably, accumulating studies have demonstrated that excessive cervical activity is one of the leading causes of the development of OPLL. 5 , 24 , 25 , 26 However, the molecular mechanism of this disease is poor understanding. In this study, OPLL cells and non‐OPLL cells were acquired from human PLL tissues. These results suggested that DLX5 enhance osteogenic differentiation through the NOTCH signaling pathway under MS, which indicated that DLX5 is essential for OPLL progression.

In embryogenesis, the DLX5 is expressed on cells in several tissues, including branchial arches, confined brain regions, expanded appendages, and bones. 27 Numerous studies have reported that MS upregulate expression of DLX5 in human BMSCs. 28 , 29 , 30 After external stimuli, activation of cAMP/PKA/CREB signaling upregulated C/EBPβ expression, which binds to the DLX5 promoter to suppress its transcription. 22 Besides, the DLX5 expression was upregulated during BMP‐2—mediated osteogenic differentiation. 19 , 21 BMP‐2 signaling responds to mechanical stress via YAP/TAZ nucleocytoplasmic translocation. 31 , 32 , 33 , 34 In addition, DLX5 was an important transcriptional factor involved in bone development and cellular differentiation. 35 The deletion of the STAT5A in mouse results in increase of trabecular bone and cortical bone mass in mice by regulating the expression of DLX5. 36 Studies have shown that DLX5 was essential for bone development, and its activated was increased by mechanical stress. However, the role of DLX5 in regulating OPLL cells osteogenic differentiation is still unclear. In this study, DLX5 expression is increased by stretch stimulation and DLX5 knockdown inhibited osteogenic gene expression in OPLL cells, which was consistent with prior findings from other researchers. 36 , 37 It has been suggested that DLX5 was involved in osteogenic gene transcription, driving OPLL cells toward osteogenic differentiation under MS.

NOTCH signaling is involved in a multiple cellular processes, including cell proliferation, differentiation, and apoptosis. 38 Physical contact with NOTCH ligands on adjacent cells activates NOTCH receptors, leading to the release of NICD. 39 It has been demonstrated that NOTCH1 acts as a mechanosensor in endothelial cells, which response to mechanical stress and regulates cellular junctions and triggers endothelial proliferation. 40 Besides, Cyclic stretch stimulation enhanced NOTCH3 expression in vascular smooth muscle cells. 41 In human ligamentum flavum cells, NOTCH promoted cell differentiation by interacting with RUNX2 and OSX. 42 Conversely, NOTCH signaling inhibits Wnt signaling and interacts with RUNX2 to suppress osteoblast differentiation in immature osteoblastic cells. 43 Our results showed that the protein expression of NICD and HES1 were significantly increased after stretch stimulation compared with control. These findings were consistent with those from previous studies. 28 , 44 , 45 It is worth mentioning that NICD nuclear translocation occurs rapidly after MS and mediates OPLL cells osteogenic differentiation. Stretch causes NICD cleavage, however the exact mechanism of nuclear translocation of the NICD induced by stretch stress is still understood. We hypothesized that MS may result in conformational changes in NOTCH receptors that enhance their binding affinity for proteases (ADAM and γ‐secretase) or increasing expression levels or activities of NOTCH receptors and ligands.

The effect of DAPT treatment on the osteogenic potential of OPLL cells was also investigated. The results shown that DAPT treatment significantly inhibited the DLX5‐mediated osteogenic differentiation, which is inconsistent with previous reports. 46 , 47 Multiple factors, including cell type, cell differentiation stage, and timing of NOTCH activation, may account for this discrepancy. It was supported that NOTCH signaling inhibits osteogenic differentiation of MSCs in the early stages but promotes this process in the late stages. 48 , 49 OPLL cells represent late stage differentiated cells, hence, activated NOTCH signaling upregulated osteogenic‐related genes expression and enhanced osteogenic differentiation. In addition, the expression of DLX5 was decreased when NOTCH signaling was inhibited by DAPT when MSCs was exposed to pulsed electromagnetic field. 29 This study demonstrated that DAPT treatment inhibits DLX5‐medited osteogenic differentiation of OPLL cells in response to mechanical stretch. Consequently, mechanical stretch‐induced NOTCH signaling activation promotes the translocation of NICD into the nucleus, which positively regulated osteogenic genes expression by enhancing DLX5 (Figure 7). Besides, no statistically significant difference in NOTCH‐related genes were detected between shDLX5 and scramble control, except for NOTCH3. The relationship between DLX5 and NOTCH3 needs further investigation.

FIGURE 7.

FIGURE 7

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The mechanism of NOTCH signaling regulating DLX5‐medited osteogenic differentiation. NOTCH activation is normally activated by cell–cell interactions. On ligand‐receptor binding, the NOTCH receptor heterodimers dissociate and undergo cleavage by ADAM and γ‐secretase complex, releasing a NOTCH intracellular structure domain (NICD) and mechanical stimulation promotes NICD cleavage. When NICD enters the nucleus, its binding to CSL may trigger an allosteric change. Mastermind (MAML) then recognizes the NICD/CSL interface and recruit coactivators (Co‐A) to activate the transcription of target genes. NOTCH signaling was blocked by treatment with the γ‐secretase inhibitor DAPT. DLX5‐medited osteogenic differentiation is regulated by NOTCH signaling.

There are some limitations of this study that should be acknowledged. First, due to the small sample size limitations, we were unable to classify the disease types into segmental, continuous, mixed, and localized OPLL in this investigation, and a larger sample scale is necessary to strengthen our conclusions. Second, it was challenging to acquire particular PLL tissue structure during surgical procedures, preventing us from performing additional experimental validation at tissue level. Finally, the results presented in this study were almost based on the OPLL cells, further extensive research with alternative ttw mouse will be required to powerfully prove our conclusions.

5. CONCLUSIONS

In this study, DLX5 was found to be highly expressed of OPLL patients both in vitro and in vivo. In OPLL cells, DLX5 plays an important role in osteogenic differentiation, and the molecular mechanism was partly due to NOTCH signaling respond to stretch and regulates DLX5‐medited osteogenic differentiation. This finding provide a new perspective to understanding of OPLL pathogenesis.

5.1. Antibody

The following primary antibodies were used: rabbit monoclonal anti‐DLX5 antibody (Abcam, Britain, 1:1000), rabbit monoclonal anti‐Cleaved Notch1 antibody (Cell Signaling Technology, 1:1000), rabbit monoclonal anti‐HES1 antibody (Abcam, 1:1000), rabbit monoclonal anti‐ RUNX2 antibody (Abcam, 1:1000), rabbit monoclonal anti‐OSX antibody (Abcam, 1:1000), rabbit monoclonal anti‐GAPDH antibody (Cell Signaling Technology, 1:1000), and goat anti‐rabbit HRP (Abcam, 1:1000).

AUTHOR CONTRIBUTIONS

Tao Tang, Zhengya Zhu, and Zhongyuan He: conceptualization, methodology, investigation, writing original draft. Fuan Wang and Jianmin Wang: methodology and data analysis. Hongkun Chen, Shengkai Liu, Mingbin Zhan: methodology and figures preparation. Dafu Chen: project administration and funding acquisition. Wei Tian and Xinbao Wu: conceptualization, supervision, and project administration. Xizhe Liu: conceptualization, supervision, writing, editing, and provided funding support. Zhiyu Zhou: conceptualization, supervision, project administration, and funding acquisition. Shaoyu Liu: conceptualization, resources, supervision, project administration, and funding acquisition. All authors have read and agreed to the published version of the manuscript.

CONFLICT OF INTEREST

The authors declare no conflict of interest.

Supporting information

TABLE S1. Primers used for RT‐qPCR

Click here for additional data file. (31.1KB, docx)

ACKNOWLEDGMENTS

This work was supported by the National Natural Science Foundation of China (U22A20162, 31900583, 32071351, 81772400, 82102604, 81960395), the Natural Science Foundation of Guangzhou City (201807010031), Foundation of Shenzhen Committee for Science and Technology Innovation (JCYJ20190809142211354, GJHZ20180929160004704), Sanming Project of Medicine in Shenzhen (SZSM201911002), the Beijing Municipal Health Commission (Grant No. BMHC‐2021‐6, BMHC‐2019‐9, BMHC‐2018‐4, PXM2020_026275_000002), AOCMF Translational approaches for bone constructs (AOCMF‐21‐04S), Sun Yat‐sen University Clinical Research 5010 Program (2019009), and Academic Affairs Office of Sun Yat‐sen University (202211583, 202211589). Special thanks are extended to Cheng Ruijuan in Active Technology Limited and Zhang Guangxu in Accuramed Technology (Shanghai) Limited (https://www.accuramed.com/) for technical support. We would like to thank Drs. Manman Gao to funding support and Guoliang Chen to providing language help and in this manuscript.

Tang, T. , Zhu, Z. , He, Z. , Wang, F. , Chen, H. , Liu, S. , Zhan, M. , Wang, J. , Tian, W. , Chen, D. , Wu, X. , Liu, X. , Zhou, Z. , & Liu, S. (2023). DLX5 regulates the osteogenic differentiation of spinal ligaments cells derived from ossification of the posterior longitudinal ligament patients via NOTCH signaling. JOR Spine, 6(2), e1247. 10.1002/jsp2.1247

Tao Tang, Zhengya Zhu, and Zhongyuan He contributed equally to this work and should be considered as co‐first authors.

Contributor Information

Xizhe Liu, Email: liuxizhe@mail.sysu.edu.cn.

Zhiyu Zhou, Email: zhouzhy23@mail.sysu.edu.cn.

DATA AVAILABILITY STATEMENT

All data generated or analyzed during this study are available from the corresponding author on reasonable request.

REFERENCES

  • 1. Inamasu J, Guiot BH, Sachs DC. Ossification of the posterior longitudinal ligament: an update on its biology, epidemiology, and natural history. Neurosurgery. 2006;58(6):1027‐1039. discussion 1027–1039. [DOI] [PubMed] [Google Scholar]
  • 2. Epstein NE, Grande DA, Breitbart AS. In vitro characteristics of cultured posterior longitudinal ligament tissue. Spine (Phila Pa 1976). 2002;27(1):56‐58. [DOI] [PubMed] [Google Scholar]
  • 3. Bernstein DN, Prong M, Kurucan E, et al. National trends and complications in the surgical management of ossification of the posterior longitudinal ligament (OPLL). Spine (Phila Pa 1976). 2019;44(22):1550‐1557. [DOI] [PubMed] [Google Scholar]
  • 4. Boody BS, Lendner M, Vaccaro AR. Ossification of the posterior longitudinal ligament in the cervical spine: a review. Int Orthop. 2019;43(4):797‐805. [DOI] [PubMed] [Google Scholar]
  • 5. Xu C, Zhang Z, Liu N, et al. Small extracellular vesicle‐mediated miR‐320 e transmission promotes osteogenesis in OPLL by targeting TAK1. Nat Commun. 2022;13(1):2467. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 6. Nam DC, Lee HJ, Lee CJ, Hwang SC. Molecular pathophysiology of ossification of the posterior longitudinal ligament (OPLL). Biomol Ther (Seoul). 2019;27(4):342‐348. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 7. Khuyagbaatar B, Kim K, Park WM, Kim YH. Influence of sagittal and axial types of ossification of posterior longitudinal ligament on mechanical stress in cervical spinal cord: a finite element analysis. Clin Biomech (Bristol, Avon). 2015;30(10):1133‐1139. [DOI] [PubMed] [Google Scholar]
  • 8. Chen J, Wang X, Wang C, Yuan W. Rotational stress: role in development of ossification of posterior longitudinal ligament and ligamentum flavum. Med Hypotheses. 2011;76(1):73‐76. [DOI] [PubMed] [Google Scholar]
  • 9. Shi L, Miao J, Chen D, Shi J, Chen Y. Endoplasmic reticulum stress regulates mechanical stress‐induced ossification of posterior longitudinal ligament. Eur Spine J. 2019;28(10):2249‐2256. [DOI] [PubMed] [Google Scholar]
  • 10. Yang H, Shi L, Shi G, et al. Connexin 43 affects osteogenic differentiation of the posterior longitudinal ligament cells via regulation of ERK activity by stabilizing Runx2 in ossification. Cell Physiol Biochem. 2016;38(1):237‐247. [DOI] [PubMed] [Google Scholar]
  • 11. Sawada T, Kishiya M, Kanemaru K, et al. Possible role of extracellular nucleotides in ectopic ossification of human spinal ligaments. J Pharmacol Sci. 2008;106(1):152‐161. [DOI] [PubMed] [Google Scholar]
  • 12. Iwasawa T, Iwasaki K, Sawada T, et al. Pathophysiological role of endothelin in ectopic ossification of human spinal ligaments induced by mechanical stress. Calcif Tissue Int. 2006;79(6):422‐430. [DOI] [PubMed] [Google Scholar]
  • 13. Yan L, Gao R, Liu Y, He B, Lv S, Hao D. The pathogenesis of ossification of the posterior longitudinal ligament. Aging Dis. 2017;8(5):570‐582. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 14. Tsukamoto N, Maeda T, Miura H, et al. Repetitive tensile stress to rat caudal vertebrae inducing cartilage formation in the spinal ligaments: a possible role of mechanical stress in the development of ossification of the spinal ligaments. J Neurosurg Spine. 2006;5(3):234‐242. [DOI] [PubMed] [Google Scholar]
  • 15. Sugii H, Grimaldi A, Li J, et al. The Dlx5‐FGF10 signaling cascade controls cranial neural crest and myoblast interaction during oropharyngeal patterning and development. Development. 2017;144(21):4037‐4045. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 16. Garaffo G, Conte D, Provero P, et al. The Dlx5 and Foxg1 transcription factors, linked via miRNA‐9 and ‐200, are required for the development of the olfactory and GnRH system. Mol Cell Neurosci. 2015;68:103‐119. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 17. Yang H, Cao Y, Zhang J, et al. DLX5 and HOXC8 enhance the chondrogenic differentiation potential of stem cells from apical papilla via LINC01013. Stem Cell Res Ther. 2020;11(1):271. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 18. Papadogiannis F, Batsali A, Klontzas ME, et al. Osteogenic differentiation of bone marrow mesenchymal stem cells on chitosan/gelatin scaffolds: gene expression profile and mechanical analysis. Biomed Mater. 2020;15(6):064101. [DOI] [PubMed] [Google Scholar]
  • 19. Lee MH, Kim YJ, Kim HJ, et al. BMP‐2‐induced Runx2 expression is mediated by Dlx5, and TGF‐beta 1 opposes the BMP‐2‐induced osteoblast differentiation by suppression of Dlx5 expression. J Biol Chem. 2003;278(36):34387‐34394. [DOI] [PubMed] [Google Scholar]
  • 20. Tanno M, Furukawa KI, Ueyama K, Harata S, Motomura S. Uniaxial cyclic stretch induces osteogenic differentiation and synthesis of bone morphogenetic proteins of spinal ligament cells derived from patients with ossification of the posterior longitudinal ligaments. Bone. 2003;33(4):475‐484. [DOI] [PubMed] [Google Scholar]
  • 21. Holleville N, Matéos S, Bontoux M, Bollerot K, Monsoro‐Burq AH. Dlx5 drives Runx2 expression and osteogenic differentiation in developing cranial suture mesenchyme. Dev Biol. 2007;304(2):860‐874. [DOI] [PubMed] [Google Scholar]
  • 22. Lee HL, Qadir AS, Park HJ, et al. cAMP/protein kinase A signaling inhibits Dlx5 expression via activation of CREB and subsequent C/EBPβ induction in 3T3‐L1 preadipocytes. Int J Mol Sci. 2018;19(10):3161. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 23. Li JM, Zhang Y, Ren Y, et al. Uniaxial cyclic stretch promotes osteogenic differentiation and synthesis of BMP2 in the C3H10T1/2 cells with BMP2 gene variant of rs2273073 (T/G). PLoS One. 2014;9(9):e106598. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 24. Tokuhashi Y, Ajiro Y, Umezawa N. A patient with two re‐surgeries for delayed myelopathy due to progression of ossification of the posterior longitudinal ligaments after cervical laminoplasty. Spine (Phila Pa 1976). 2009;34(2):E101‐E105. [DOI] [PubMed] [Google Scholar]
  • 25. Chen D, Chen Y, Li T, Shi L, Pan M, Chen D. Role of Cx43‐mediated NFsmall ka, CyrillicB signaling pathway in ossification of posterior longitudinal ligament: an in vivo and in vitro study. Spine (Phila Pa 1976). 2017;42(23):E1334‐E1341. [DOI] [PubMed] [Google Scholar]
  • 26. Chen S, Zhu HF, Wang GL, Xie Z, Wang JY, Chen J. Combined use of leptin and mechanical stress has osteogenic effects on ossification of the posterior longitudinal ligament. Eur Spine J. 2018;27(8):1757‐1766. [DOI] [PubMed] [Google Scholar]
  • 27. Tan Y, Testa JR. DLX genes: roles in development and cancer. Cancers (Basel). 2021;13(12):3005. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 28. Ziouti F, Ebert R, Rummler M, et al. NOTCH signaling is activated through mechanical strain in human bone marrow‐derived mesenchymal stromal cells. Stem Cells Int. 2019;2019:5150634. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 29. Bagheri L, Pellati A, Rizzo P, et al. Notch pathway is active during osteogenic differentiation of human bone marrow mesenchymal stem cells induced by pulsed electromagnetic fields. J Tissue Eng Regen Med. 2018;12(2):304‐315. [DOI] [PubMed] [Google Scholar]
  • 30. Ongaro A, Pellati A, Bagheri L, et al. Characterization of notch signaling during osteogenic differentiation in human osteosarcoma cell line MG63. J Cell Physiol. 2016;231(12):2652‐2663. [DOI] [PubMed] [Google Scholar]
  • 31. Zouani OF, Kalisky J, Ibarboure E, Durrieu MC. Effect of BMP‐2 from matrices of different stiffnesses for the modulation of stem cell fate. Biomaterials. 2013;34(9):2157‐2166. [DOI] [PubMed] [Google Scholar]
  • 32. Shekaran A, García JR, Clark AY, et al. Bone regeneration using an alpha 2 beta 1 integrin‐specific hydrogel as a BMP‐2 delivery vehicle. Biomaterials. 2014;35(21):5453‐5461. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33. Tan S, Fang JY, Yang Z, Nimni ME, Han B. The synergetic effect of hydrogel stiffness and growth factor on osteogenic differentiation. Biomaterials. 2014;35(20):5294‐5306. [DOI] [PubMed] [Google Scholar]
  • 34. Wei Q, Holle A, Li J, et al. BMP‐2 signaling and mechanotransduction synergize to drive osteogenic differentiation via YAP/TAZ. Adv Sci (Weinh). 2020;7(15):1902931. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 35. Acampora D, Merlo GR, Paleari L, et al. Craniofacial, vestibular and bone defects in mice lacking the Distal‐less‐related gene Dlx5. Development. 1999;126(17):3795‐3809. [DOI] [PubMed] [Google Scholar]
  • 36. Lee KM, Park KH, Hwang JS, et al. Inhibition of STAT5A promotes osteogenesis by DLX5 regulation. Cell Death Dis. 2018;9(11):1136. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 37. Kawane T, Komori H, Liu W, et al. Dlx5 and mef2 regulate a novel runx2 enhancer for osteoblast‐specific expression. J Bone Miner Res. 2014;29(9):1960‐1969. [DOI] [PubMed] [Google Scholar]
  • 38. McGirt LY, Jia P, Baerenwald DA, et al. Whole‐genome sequencing reveals oncogenic mutations in mycosis fungoides. Blood. 2015;126(4):508‐519. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 39. Kopan R, Ilagan MX. The canonical Notch signaling pathway: unfolding the activation mechanism. Cell. 2009;137(2):216‐233. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 40. Mack JJ, Mosqueiro TS, Archer BJ, et al. NOTCH1 is a mechanosensor in adult arteries. Nat Commun. 2017;8(1):1620. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 41. Zhu JH, Chen CL, Flavahan S, Harr J, Su B, Flavahan NA. Cyclic stretch stimulates vascular smooth muscle cell alignment by redox‐dependent activation of Notch3. Am J Physiol Heart Circ Physiol. 2011;300(5):H1770‐H1780. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 42. Qu X, Chen Z, Fan D, et al. Notch signaling pathways in human thoracic ossification of the ligamentum flavum. J Orthop Res. 2016;34(8):1481‐1491. [DOI] [PubMed] [Google Scholar]
  • 43. Zanotti S, Canalis E. Notch signaling and the skeleton. Endocr Rev. 2016;37(3):223‐253. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 44. Karakaya C, van Turnhout MC, Visser VL, et al. Notch signaling regulates strain‐mediated phenotypic switching of vascular smooth muscle cells. Front Cell Dev Biol. 2022;10:910503. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 45. Masumura T, Yamamoto K, Shimizu N, Obi S, Ando J. Shear stress increases expression of the arterial endothelial marker ephrinB2 in murine ES cells via the VEGF‐Notch signaling pathways. Arterioscler Thromb Vasc Biol. 2009;29(12):2125‐2131. [DOI] [PubMed] [Google Scholar]
  • 46. Hilton MJ, Tu X, Wu X, et al. Notch signaling maintains bone marrow mesenchymal progenitors by suppressing osteoblast differentiation. Nat Med. 2008;14(3):306‐314. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 47. Tu X, Chen J, Lim J, et al. Physiological notch signaling maintains bone homeostasis via RBPjk and hey upstream of NFATc1. PLoS Genet. 2012;8(3):e1002577. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 48. Lawal RA, Zhou X, Batey K, et al. The notch ligand Jagged1 regulates the osteoblastic lineage by maintaining the osteoprogenitor pool. J Bone Miner Res. 2017;32(6):1320‐1331. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 49. Goel PN, Moharrer Y, Hebb JH, et al. Suppression of notch signaling in osteoclasts improves bone regeneration and healing. J Orthop Res. 2019;37(10):2089‐2103. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

TABLE S1. Primers used for RT‐qPCR

Click here for additional data file. (31.1KB, docx)

Data Availability Statement

All data generated or analyzed during this study are available from the corresponding author on reasonable request.

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