The multi-potential bone marrow stromal cell line KusaO was obtained from Dr Julian W Quinn (The University of Melbourne, Melbourne, Australia) and was used for the evaluation of osteogenesis in vitro (13). Osteogenic induction was performed as previously described (14). Specifically, cells were cultured at 37°C and 5% CO₂ in osteogenic medium, which consisted of low glucose (LG)-DMEM (Gibco; Thermo Fisher Scientific, Inc.) supplemented with 10% foetal bovine serum (FBS; Gibco; Thermo Fisher Scientific, Inc.), 100 nM dexamethasone (MilliporeSigma), 50 μM ascorbic acid (MilliporeSigma) and 20 mM β-glycerophosphate (MilliporeSigma). Cells were also cultured in complete medium, which comprised LG-DMEM containing 10% FBS, as a control.

ENO (CAS no. 11029-12-2; MedChemExpress) at different concentrations (7.5, 15, 30 and 60 μM) was added to the osteogenic medium to evaluate the effect of ENO on the osteogenic differentiation of KusaO cells in vitro. The cells were treated for 24, 48 or 72 h to assess cell proliferation, 3 days for reverse transcription-quantitative PCR (RT-qPCR) and alkaline phosphatase (ALP) staining, 7 days for western blotting, and 14 days for alizarin red S (ARS) staining. The medium was replaced every 3 days during the treatment period. The changes in osteoblastic genes were detected using RT-qPCR, and ALP staining was used to assess the osteogenic differentiation of KusaO cells. Western blotting was used to evaluate the protein expression of osteoblastic markers. ARS staining was used to assess the mineralization of KusaO cells.

Cell proliferation was assessed using the CellTiter 96^® AQueous One Solution Cell Proliferation Assay (MTS) (Promega Corporation) according to the manufacturer's protocol. The cells were seeded at a density of 5×10³ cells/well in 96-well plates. After a 24-h incubation, the cells were treated with ENO at various concentrations (7.5, 15, 30 and 60 μM) for 24, 48 and 72 h at 37°C and 5% CO₂. Subsequently, 10 μl MTS was added to each well and incubated at 37°C with 5% CO₂ for 4 h. The absorbance was measured at 490 nm using a Bio-Rad 680 microplate reader (Bio-Rad Laboratories, Inc.) and cell proliferation was expressed as a percentage of the control culture value, which was considered 100% viable.

Cells were cultured in osteogenic medium with or without various concentrations of ENO (7.5, 15, 30 and 60 μM) and incubated for 3 days at 37°C and 5% CO₂. Subsequently, the Alkaline Phosphatase Detection Kit (MilliporeSigma) was used, according to the manufacturer's protocol. Briefly, cells were fixed in 95% ethanol for 10 min at room temperature and were subsequently incubated with ALP regent at 37°C for another 10 min. After rinsing with PBS; ight microscopy (Nikon Corporation) was used to capture images of the cells with purple ALP staining. To semi-quantify positive ALP staining, Fiji ImageJ2 software was used (15).

Cells were cultured in osteogenic medium containing different concentrations of ENO (0, 7.5, 15, 30 and 60 μM) for 14 days at 37°C and 5% CO₂. After fixing the cells with 95% ethanol for 10 min at room temperature, they were stained with 0.5% ARS (cat. no. A5533; MilliporeSigma) solution (pH 4.2) for 10 min at room temperature to visualize mineral deposits, which appeared as red-stained areas. Images were captured using a UMAX Scanner (PowerLook 2100XL-USB; VueScan) and a light microscope (Nikon Corporation).

Cells were cultured in osteogenic medium containing 30 μM ENO, and MMP9 protein (cat. no. 909-MM-010; R&D Systems, Inc.) was added at different concentrations (100 and 200 ng/ml) to observe whether the effects of ENO on osteogenesis could be reversed by MMP9. The cells were treated for 3, 7 or 14 days at 37°C and 5% CO₂, and the cells were then collected for further RT-qPCR after 3 days, western blotting after 7 days and ARS staining after 14 days to examine changes in osteoblastic genes and proteins, as well as the mineralization of cells.

Cells were plated in 24-well plates and were cultured in osteogenic medium containing different concentrations of ENO (0, 15, 30 and 60 μM) for 3 days at 37°C and 5% CO₂, with 6 replicates for each concentration (n=6). RT-qPCR was performed to detect the changes in the expression levels of osteoblastic genes after ENO treatment, and the experiment was performed as described in our previous study (14). Briefly, total RNA was extracted using TRIzol^® (Invitrogen; Thermo Fisher Scientific, Inc.), and cDNA was synthesized with PrimeScript RT Master Mix (cat. no. RR036B; Takara Biotechnology, Ltd.), according to the manufacturers' protocols. The cDNA then underwent qPCR with Power UP SYBR Green Master Mix (cat. no. A25742, Thermo Fisher Scientific, Inc.) on an Analytik Jena qTOWER (Analytik Jena GmbH). The amplification conditions for qPCR were as follow: 50°C for 2 min and 95°C for 5 min, followed by 40 cycles at 95°C for 15 sec and 60°C for 1 min, with a melt curve stage of 95°C for 15 sec, 60°C for 1 min and 95°C for 15 sec. The relative mRNA expression levels of all genes were normalized to the housekeeping gene Gapdh and were calculated using the 2^−ΔΔCq method (16). The primers used were listed in Table I and Mmp9 primers were purchased from Sino Biological, Inc. (cat. no. MP200552).

For western blotting, proteins were extracted from cells with or without treatment using RIPA lysis and extraction buffer (Thermo Fisher Scientific, Inc.) with Halt Protease Inhibitor Cocktail (100X; Thermo Fisher Scientific, Inc.). The concentration of extracted proteins was determined using the BCA Protein Assay (Thermo Fisher Scientific, Inc.), and 20 μg proteins were separated by SDS-PAGE on 10% gels made using TGX FastCast acrylamide kits (Bio-Rad Laboratories, Inc.). The proteins were run alongside a molecular weight marker (Pierce; Thermo Fisher Scientific, Inc.) for 1.5 h at 100 V. The proteins were then transferred from the gel to PVDF membranes using the Trans-Blot Turbo Transfer System (Bio-Rad Laboratories, Inc.). The membranes were incubated in 5% non-fat milk in TBS-1% Tween for 1 h at room temperature and then incubated overnight at 4°C with primary antibodies against Runx2 (1:1,000; cat. no. 12556; Cell Signaling Technology, Inc.), osteopontin (OPN; 1:1,000; cat. no. ab63856; Abcam) MMP9 (1:500; cat. no. 10375-2-AP; Wuhan Sanying Biotechnology) and β-actin (1:10,000; cat. no. ab32572; Abcam). The next day, the membranes were incubated with HRP-conjugated anti-rabbit and anti-mouse secondary antibodies (1:3,000; cat. nos. 7074 and 7076; Cell Signaling Technology, Inc.) for 1 h at room temperature. For signal development, BeyoECL Plus reagent (Beyotime Institute of Biotechnology) was used according to the manufacturer's recommendations. Images were acquired in the dark using the ChemiDoc XRS Imaging System (Bio-Rad Laboratories, Inc.) and protein expression levels were analysed using Image Lab 5.2.1 software (Bio-Rad Laboratories, Inc.).

Total RNA was extracted from KusaO cells with or without 30 μM ENO treatment under 3-day osteogenic induction using TRIzol. Purity was assessed using a NanoDrop (OD260/280 and OD260/230 ratios; NanoDrop; Thermo Fisher Scientific, Inc.) and integrity was evaluated using an Agilent 4200 TapeStation (Agilent Technologies, Inc.). Libraries were prepared by Chengqi Biotechnology, which involved mRNA isolation, fragmentation and cDNA synthesis. Library quality was assessed using an Agilent 4200 TapeStation. Sequencing was performed on a NovaSeq 6000 platform (Illumina, Inc.) using NovaSeq6000 S4 Reagent kit v1.5 (cat. no. 20028312; Illumina, Inc.), generating 150 bp strand-specific paired-end reads. The loading concentration of the final library was 100 pM.

Bioinformatics analysis was carried out by Chengqi Biotechnology. Briefly, raw sequencing reads underwent quality control using FastQC (v0.11.9, https://www.bioinformatics.babraham.ac.uk/projects/fastqc/) and trimming with Trimmomatic (v0.36, http://www.usadellab.org/cms/?page=trimmomatic). Clean reads were aligned to HISAT2 (2.2.0, https://daehwankimlab.github.io/hisat2/). FeatureCounts (v2.0.4, https://subread.sourceforge.net/featureCounts.html) was used to count the read numbers mapped to each gene. Differential expression analysis was conducted with edgeR (v3.40.2, https://bioconductor.org/packages/release/bioc/html/edgeR.html), identifying differentially expressed genes (DEGs) based on a |log2 fold change| >2 and adjusted P<0.05. Functional enrichment analysis of DEGs was subsequently performed. ClusterProfiler R (v4.6.2, https://bioconductor.org/packages/release/bioc/html/clusterProfiler.html) was used to test the statistical enrichment of DEGs in Kyoto Encyclopedia of Genes and Genomes (KEGG) pathways. Gene Set Enrichment Analysis (GSEA) can be used to determine whether a predefined set of genes exhibits statistically significant differences between control and ENO groups. The local version of the GSEA tool was used in the present study (http://www.broadinstitute.org/gsea/index.jsp), and KEGG datasets were used for GSEA.

All animal experiments were approved by the Institutional Animal Care and Use Committee of Ruiye Bio-tech Guangzhou Co., Ltd. (approval no. RYEth-20210716213), and were carried out in accordance with The Code of Ethics of the World Medical Association (17). The animal experiments were performed at the animal laboratory of Ruiye Bio-tech Guangzhou Co., Ltd. by the authors. Animals were maintained in a standard room at 22±2°C with 40-60% humidity, under a 12-h light/dark cycle, with ad libitum access to food and water according to the institutional animal guidelines. A total of 12 female Sprague-Dawley rats (weight, 200-250 g) underwent open femoral fracture surgery as previously reported (18). Rats were anesthetized with an intraperitoneal injection of 0.6% pentobarbital sodium (40 mg/kg). Subsequently, the femurs were exposed and cut down the middle to construct femur fracture models. The fractures were then internally fixed using intramedullary 1.2-mm Kirschner wires. After surgery, the rats were housed in individual cages with soft bedding, and easy access to food and water. To minimize the pain of surgery, buprenorphine (0.05 mg/kg) was administered subcutaneously every 8-12 h for 48-72 h post-surgery (17). (The rats were randomly divided into three groups (n=4/group): Saline group, 30 mg/kg ENO group and 60 mg/kg ENO group. After 3 days of recovery from the operation, ENO was injected locally at the fracture sites every 3 days for 6 weeks, with different concentrations used for each group. The Saline group of rats received saline injections as a control.

A total of 6 weeks after the operation, all rats were sacrificed by an overdose of pentobarbital sodium (0.6%) anaesthesia at a dose of 300 mg/kg by intraperitoneal injection. Death was confirmed by the cessation of heartbeat and respiratory movements, along with the absence of reflexes. Fractured femurs were collected for stereomicroscopy (SMZ25; Nikon Corporation) and were fixed for micro-CT analysis. Femurs were fixed in 10% neutral-buffered formalin solution for 3 days at room temperature. To assess the conditions of bone regeneration, the femurs were scanned using the Skyscan 1176 μCT scanner (Bruker Corporation). Reconstructive images were generated using NRecon v1.6 software (Bruker Corporation) and were then analysed by CTAn v1.9 (Bruker Corporation). Two-and three-dimensional images were generated by Data-viewer and CTvox software (Bruker Corporation), respectively. For bone fracture healing analysis, the defect size of the cortical bone was selected. The bone mass of each group was analysed using CTAn v1.9. The parameters measured included bone volume/tissue volume (BV/TV) and bone mass density (BMD).

After micro-CT scanning, all femurs from the rats were decalcified in 10% EDTA at 37°C for 10 days followed by 1 month in 10% EDTA. Subsequently, the femurs were processed for paraffin embedding as previously described (19). For further staining, 5-μm sections were utilized. Haematoxylin and eosin (H&E) and Safranin O/Fast green were applied to evaluate the healing of femur fractures as described previously (19), which included bone regeneration and the expression of proteoglycan at the callus bone areas. Images were captured using a stereomicroscope (SMZ25; Nikon Corporation) and a light microscope (Nikon Corporation).

Immunohistochemistry was applied using a Mouse and Rabbit specific HRP/DAB detection IHC kit (cat. no. SAP-9100; OriGene Technologies, Inc.) as described previously (19). Sections were treated according to the manufacturer's protocol, including incubation with primary antibodies overnight at 4°C. The primary antibodies included OPN (1:200; cat. no. ab63856; Abcam), Collagen I (1:200; cat. no. ab6308; Abcam), Collagen II (1:200; cat. no. 32160702; MilliporeSigma), osteocalcin (OCN; 1:200; cat. no. ab133612; Abcam) and MMP9 (1:200; cat. no. 10375-2-AP; Wuhan Sanying Biotechnology). Images were captured under a light microscope (Nikon Corporation) and were analysed using Fiji ImageJ2 software.

Data are presented as the mean ± SD. All data analyses were performed using GraphPad Prism 10.0 (Dotmatics) by one-way ANOVA followed by Dunnett's multiple comparisons test. At least three independent experiments were performed. P≤0.05 was considered to indicate a statistically significant difference.

To investigate the potential impact of ENO on osteoblast differentiation of mesenchymal stem cells (MSCs), KusaO cells, which have multi-differentiation abilities in vitro were employed and cultured in either complete medium or osteogenic medium supplemented with different concentrations of ENO (0, 7.5, 15, 30 and 60 μM). To determine whether ENO affected the proliferation of KusaO cells, an MTS assay was used, and the results showed that ENO had no significant effect after 24, 48 or 72 h of treatment (Fig. 1A). ALP is recognized as an important early marker of osteoblast differentiation and was measured to assess the effects of ENO on early osteogenesis. The results showed the levels of ALP were significantly increased in a dose-dependent manner after KusaO cells were treated with different concentrations of ENO (Fig. 1B and C). Additionally, the mineralization of KusaO cells was observed through ARS staining, and the results indicated that ENO significantly increased the mineral deposits of KusaO cells in a dose-dependent manner (Fig. 1D and E). Consistently, RT-qPCR results demonstrated that osteoblastic genes, including Runx2, Alpl, Spp1, Bglap, Bsp and Sp7, were markedly upregulated by 30 μM ENO (Fig. 1F). Western blotting further confirmed that only 30 μM ENO promoted the expression levels of the osteoblastic proteins, Runx2 and OPN, which was consistent with the gene expression findings (Fig. 1G and H). In summary, ENO promoted the osteogenic differentiation of KusaO cells in vitro, with the best effects observed in response to 30 μM ENO.

Based on the in vitro effects of ENO on promoting the osteoblastic differentiation of KusaO cells, the present study further explored its effects on bone regeneration in a rat model of fracture healing in vivo. After 7 days of post-surgery recovery, 30 and 60 mg/kg ENO were injected into the fracture sites. Subsequently, stereomicroscopy, micro-CT and histological staining were applied to detect new bone formation after ENO treatment (Fig. 2A). The stereomicroscope detected bone callus formation at the fracture sites, with little callus formation observed in both ENO groups, whereas more callus formation was found in the Saline group (Fig. 2B). Similarly, the micro-CT showed that the size of bone calluses in the ENO treatment groups was much less than that in the Saline group (Fig. 2D). In addition, non-union was detected in the Saline group, whereas integration at the fracture site was markedly better after ENO treatment (Fig. 2C). Furthermore, bone parameters, including BMD and BV/TV, were significantly increased following ENO treatment, with the best results observed in the 30 mg/kg ENO group (Fig. 2D), suggesting that ENO induced more new bone formation compared with the Saline group. Consistently, H&E staining showed more bone formation in the ENO groups compared with that in the Saline group, and woven bone formation was detected in the ENO groups, indicating faster and earlier bone remodelling after ENO treatment (Fig. 2E). Fracture healing is considered the process of endochondral formation, which was measured by Safranin O/Fast Green staining (Fig. 2F); The present results indicated a notable amount of proteoglycan (red), and oval or round chondrocyte-like cells at the bone callus sites in the Saline group, whereas little could be seen in the ENO groups, suggesting that the process of endochondral formation was accelerated by ENO. Notably, 30 mg/kg ENO exhibited the best effects on promoting cartilage resorption and new bone formation.

Immunohistochemical staining was used to evaluate the expression of osteoblast-related proteins at the callus sites of the femur (Fig. 3). The results showed that ENO not only increased the expression of OCN (Fig. 3A and E), which is a marker of late osteoblastic differentiation, and OPN (Fig. 3B and F), but also decreased collagen II expression (Fig. 3D and H) at the fracture sites. In addition, 60 mg/kg ENO significantly promoted collagen I expression (Fig. 3C and G). Chondrocytes could be seen in the Collagen II-positive areas in the Saline group (black arrows) (Fig. 3D). These findings are consistent with the results of histological staining, which showed reduced cartilage and increased new bone formation after ENO treatment.

To determine the underlying mechanisms governing the effects of ENO on MSC osteogenesis, RNA-sequencing was performed. Total RNA was isolated from cells with or without ENO treatment and RNA-sequencing analysis was then performed. The results showed that 771 DEGs were identified, with 260 genes showing significant downregulation and 511 genes showing upregulation after ENO treatment compared with in the control group (Fig. 4A and B). KEGG analysis revealed that the cells treated with ENO were enriched in the 'Wnt signaling pathway' and 'cytokine-cytokine receptor interaction' pathways, which are classic signalling pathways associated with osteogenesis (20,21) (Fig. 4C). The 'estrogen signalling pathway', which is highly related to osteoporosis, was also involved in the role of ENO. These results highlight the role of ENO in osteogenesis. In addition, 'protein digestion and absorption' was involved in the effect of ENO on KusaO cells induced by osteogenic factors (Fig. 4C), indicating that ENO was involved in regulating matrix degradation. To further identify the gene sets that are related with osteogenesis with or without ENO treatment, GSEA was performed. ENO-induced genes were enriched in 'bone mineralization' (Fig. 4D and E), 'bone development' (Fig. 4F and G) and 'ossification' (Fig. 4H and I), indicating that ENO exerted effects that are highly associated with osteogenesis and bone formation. Collectively, these RNA-sequencing data showed that ENO may be highly associated with osteogenic differentiation and bone formation.

Since the KEGG pathway analysis showed that 'protein digestion and absorption' was involved in the role of ENO in osteogenesis, proteinase-related genes were selected from the DEGs. Among these genes, Mmp9 was downregulated and Prss35 was upregulated by ENO treatment (Fig. 5A). Mmp9 is known to be associated with endochondral ossification (22), while there is little evidence to support its involvement in osteogenic differentiation. However, there are few reports on Prss35 in bone homeostasis. As such, the present study further examined the mRNA and protein expression levels of MMP9 after ENO treatment in vitro and in vivo. The results revealed that, in vitro, both the mRNA and protein expression levels of MMP9 were decreased after ENO treatment in a dose-dependent manner (Fig. 5B-D). Similarly, in vivo, the expression of MMP9 at callus sites was significantly decreased by ENO treatment (Fig. 5E). In the Saline group, MMP9 was strongly expressed at the callus sites, whereas much less MMP9-positive staining could be detected in the ENO groups.

Based on the findings of RNA-sequencing analysis and confirmation that MMP9 was decreased after ENO treatment, the present study further explored whether MMP9 protein treatment could reverse the role of ENO in the osteogenic differentiation of KusaO cells. The results showed that ENO increased the mineral deposits of KusaO cells, which was consistent with the aforementioned results; however, recombinant MMP9 protein treatment attenuated their mineralization in the presence of ENO (Fig. 6A and 6B). Moreover, the mRNA expression levels of several osteoblastic genes, including Alpl, Runx2, Spp1, Sp7 and Bsp, were downregulated by MMP9 treatment, while there was no significant difference in the mRNA expression levels of Bglap between the groups with or without MMP9 treatment (Fig. 6C). Consistent results were detected regarding the expression levels of osteoblast-related proteins, including Runx2 and OPN, which were significantly decreased by 200 ng/ml MMP9 treatment (Fig. 6D and E).