The experimental protocol conforms to the ethical standards of the Declaration of Helsinki and was approved by the ethics committee of the Affiliated Hospital of Guangdong Medical University (Zhanjiang, Guangdong, China). Bone marrow samples were obtained from the femoral cavity during the operation of five female patients aged 35 to 45 years with traumatic femoral head fractures. Each sample was treated individually and used for subsequent experiments. All participants provided written informed consent. The bone marrow samples were collected from each participant and hBMSCs were isolated using Ficoll density gradient centrifugation. Subsequently, the bone marrow samples were transferred to a 15-ml centrifuge tube and diluted with PBS at a ratio of 1:1. An equal volume of Histopaque^®-1077 (Sigma-Aldrich; Merck KGaA) was then added to the diluted bone marrow sample and centrifuged at 400 × g and room temperature for 20 min without accelerating or braking. Cells in the middle layer were extracted and further resuspended in Modified Eagle's Medium α (α-MEM) supplemented with 10% (v/v) fetal bovine serum (FBS), 10 U/ml penicillin G and 10 mg/ml streptomycin (all from Gibco; Thermo Fisher Scientific, Inc.). The obtained cells were incubated at 37°C in an incubator containing air with 5% CO₂ and saturated humidity. The medium was changed every three days; when the cells reached ~90% confluence, they were digested with 0.25% trypsin-EDTA (Gibco; Thermo Fisher Scientific, Inc.) and passaged at a 1:3 ratio. The cells from passages five to six were used in all subsequent experiments.

The cells from passage three were used to identify phenotypes. When the cells had reached 80% confluence, they were digested with 0.25% trypsin-EDTA (Gibco; Thermo Fisher Scientific, Inc.), centrifuged at 800 × g for 3 min at room temperature and resuspended in PBS. The cell surface antigens CD90, CD73, CD105 and CD45 were detected via flow cytometry using the Human MSC Analysis Kit (cat. no. 562245; BD Biosciences). Data were analyzed using FlowJo 10.0.7 software (FlowJo LLC).

The hBMSCs were seeded in 6-well plates at a density of 1×10⁵ cells per well. The hBMSCs were cultured with adipogenic induction medium (α-MEM containing 10% FBS, 10 mg/ml insulin, 1 mM dexamethasone, 50 mM 3-isbutyl-1-methylxanthine and 50 µM indomethacin; all from Sigma-Aldrich; Merck KGaA) and chondrogenic induction medium (α-MEM containing 1% FBS, 1% insulin-transferrin-sodium selenite, 20 nM dexamethasone, 10 ng/ml TGF-β and 40 µg/ml proline; all from Sigma-Aldrich; Merck KGaA) and stained with Oil Red O or Alcian Blue staining solution (Cyagen Biosciences, Inc.). The images were acquired under a microscope and stored.

The RH strain of T. gondii (kindly provided by Dr Young-Ha Lee, Department of Infection Biology, Chungnam National University School of Medicine, Republic of Korea) was allowed to proliferate in ARPE-19 cells (CRL-2302; American Type Culture Collection). T. gondii was collected and rinsed in PBS, centrifuged at 1,000 × g for 5 min at room temperature, and this was repeated twice. Purified tachyzoites were collected in 1 ml Hank's balanced salt solution (Gibco; Thermo Fisher Scientific, Inc.) and oscillated at 3.57 × g in a constant temperature shaker at 37°C for 4 h. After centrifugation at 14,000 × g thrice for 5 min each at 4°C, the supernatant was stored as TgESPs.

To visualize vesicle-like structures in TgESPs, the TgESPs were deposited on a nickel grid and stained with uranyl acetate and lead citrate, and images of the samples were acquired using a JEM-1400 transmission electron microscope (JEOL, Ltd.).

The effect of TgESPs on the proliferation of hBMSCs was evaluated using the Cell Counting Kit-8 (CCK-8; Zeta Life). In brief, hBMSCs were seeded in 96-well plates at a density of 3,000 cells per well and incubated with different concentrations of TgESPs (0, 1, 10, 20, 30, 40 and 50 µg/ml) for 24, 48 and 72 h. The wells were then filled with 10 µl CCK-8 solution and incubated in 5% CO₂ in a humidified environment for 2 h at 37°C. The absorbance of the cells at 450 nm was measured using a microplate reader (BioTek Instruments, Inc.) and a cell proliferation curve was generated.

The hBMSCs were seeded in 12-well plates at a density of 1×10⁵ cells per well. Once they reached >80% confluence, the medium was changed to the osteogenic induction medium (OIM), i.e. Dulbecco's modified Eagle's medium (DMEM) with low glucose (LG) content, supplemented with 100 nM dexamethasone (Sigma-Aldrich; Merck KGaA), 50 µM L-ascorbic acid-2-phosphate (Sigma-Aldrich; Merck KGaA), 10 mM β-glycerophosphate (Sigma-Aldrich; Merck KGaA) and 10% FBS (Gibco; Thermo Fisher Scientific, Inc.). The experimental group was incubated in OIM supplemented with 1 or 10 µg/ml TgESPs, whereas the control group was incubated in OIM without TgESPs, and the inhibitor group was treated with 25 µM Icg-001 (MCE) or 2 mM 2-deoxy-D-glucose (2-DG; Sigma-Aldrich; Merck KGaA) for three days to induce osteogenic differentiation. The osteogenic induction medium was changed every three days.

The cellular mineral deposition was evaluated using alizarin red staining. The cells were fixed with 4% paraformaldehyde for 15 min at room temperature. Subsequently, they were washed thrice with PBS, treated with Alizarin Red S staining solution (Oricell) for 20 min at room temperature and washed thrice with PBS. The images were acquired and saved using a camera and microscope.

Cells were seeded in 6-well plates at a density of 2×10⁵ cells per well. Total RNA was extracted from the hBMSCs using TRIzol reagent (Invitrogen; Thermo Fisher Scientific, Inc.) at 7 days after osteogenic induction and reverse-transcribed into cDNA using HiScript III RT SuperMix (Vazyme), ChamQ Universal SYBR qPCR Master Mix (Vazyme) was used for qPCR, and both cDNA synthesis and qPCR were performed according to the manufacturer's protocol. The qPCR thermocycling conditions were as follows: 95°C for 10 sec and 60°C for 30 sec, performed for 40 cycles. The mRNA expression levels of target genes were normalized to those of the internal control (β-actin) and quantified using the 2^−ΔΔCq method (24). The primers used for PCR are listed in Table SI.

Cells were lysed using RIPA buffer (Invitrogen; Thermo Fisher Scientific, Inc.) 14 days after the induction of osteogenic differentiation. Protein concentrations were determined using a BCA protein assay kit (Beyotime Institute of Biotechnology). For each lane, 20 µg of protein were loaded on a 10 or 12% SDS-PAGE gel, resolved and electrotransferred to a polyvinylidene difluoride membrane (Merck Millipore). The membrane was then placed in 5% skimmed milk (0.5 g skimmed milk powder dissolved in 10 ml Tris-buffered saline), blocked at room temperature for 1-2 h, and incubated with a primary antibody at 4°C overnight. After washing it thrice with Tris-buffered saline containing Tween 20 (TBS-T), the membrane was incubated with a horseradish peroxidase-conjugated secondary antibody for 1 h at room temperature. After washing the membrane thrice with TBST (10 min each), immunoreactive bands were visualized using a chemiluminescence detection reagent (Merck Millipore). Finally, protein intensity was quantified using the ImageJ software version 1.8.0 (National Institutes of Health). All the antibodies used for western blot analysis are listed in Table SII.

Cells were induced to undergo osteogenic differentiation in OIM with or without TgESPs for 7 days. In addition, RUNX family transcription factor 2 (Runx2) expression was measured using IF. In brief, the cells were fixed in 4% paraformaldehyde for 20 min, permeabilized in 0.5% Triton X-100 (Sigma-Aldrich; Merck KGaA) for 30 min and blocked in 1% bovine serum albumin (Beijing Solarbio) for 1 h. The cells were then washed thrice with PBS (5 min each) and incubated overnight at 4°C with a specific primary antibody against Runx2. The cells were subsequently incubated with AlexA-568-conjugated secondary antibody for 1 h at room temperature. The nuclei were stained with DAPI. Immunofluorescence was observed using a fluorescence microscope (Olympus Corporation). Details of all antibodies used for IF are provided in Table SII.

The hBMSCs were cultured in OIM for 7 days. Cells were harvested following trypsin digestion and re-seeded in Seahorse XF24 cell culture microplates (Seahorse Bioscience) at a density of 2×10⁴ cells per well. Cells were cultured overnight in OIM and the OCR and ECAR were measured using the Seahorse XF Cell Mito Stress Test kit (Seahorse Bioscience) and Seahorse XF Glycolysis Rate Assay kit (Seahorse Bioscience), respectively, according to the manufacturer's instructions. The Cell Mito Stress Test assesses ATP production and maximal mitochondrial respiration by successively adding specific inhibitors, such as oligomycin, carbonyl cyanide 4-(trifluoromethoxy) phenylhydrazone (FCCP, a potent mitochondrial oxidative phosphorylation uncoupling agent that promotes mitochondrial membrane depolarization. In this study, we employed FCCP to investigate the impact of TgESP on cellular energy metabolism.) and rotenone/antimycin A (Rot/AA). Afterwards, the glycolysis rate assay was carried out using Rot/AA and 2-DG to measure ECAR and intracellular glycolysis capacity. The OCR and ECAR were measured and analyzed using an XF24 Flux Analyser (Seahorse Bioscience).

For lactic acid measurement, a lactic acid detection kit (cat. no. BC2235; Beijing Solarbio) was used according to the manufacturer's protocol. The culture medium of the cells was collected, and mixed with working solution of the lactic acid detection kit. Subsequently, the mixture was incubated with the color reagent at 37°C in the dark for 20 min. Finally, the absorbance at 570 nm was measured using a microplate reader (BioTek Instruments, Inc.) and the lactic acid concentration was calculated based a standard curve.

The cell wound healing assay was performed as previously described (25). In brief, hBMSCs were seeded at a density of 2×10⁵ cells per well in 6-well plates and treated with 1 or 10 µg/ml TgESPs for 24 h. After this treatment, a vertical scratch was made in the cell monolayer using a 20-µl micropipette tip. The cells were then washed once with PBS and incubated with serum-free basal medium. The migration of hBMSCs was observed and recorded using Leica DMI3000 microscope (Leica Microsystems GmbH) at 0 and 24 h. The cell migration distance was quantified by measuring the cell distance between the two sides using Photoshop 2021 (Adobe Inc.).

The cell adhesion assay was conducted following a previously reported protocol (26). In brief, hBMSCs were seeded in 6-well plates and treated with 1 and 10 µg/ml TgESPs for 24 h. After this treatment, the cells were digested, resuspended and seeded at a density of 1×10⁴ cells per well in 96-well plates coated with polylysine (Beijing Solarbio). After allowing the cells to attach for 1 h, non-adherent cells were washed off with PBS. Subsequently, 10 µl MTS assay solution was added to each well and incubated at 37°C for 2 h. The absorbance was measured at 490 nm using a microplate reader (BioTek Instruments, Inc.). The relative cell adhesion rate was thus evaluated.

Statistical analyses were performed using GraphPad Prism 8.0 software (GraphPad Software; Dotmatics). Comparisons of data between two groups were performed using an unpaired two-tailed Student's t-test, while data were compared among three or more groups by one-way ANOVA followed by Tukey's post-hoc test. Values are expressed as the mean ± standard deviation and were based on results from at least three independent experiments. P<0.05 was considered to indicate a statistically significant difference.

hBMSCs were isolated and cultured from human bone marrow samples and passaged three to four times. After 10 days of primary culture, the attached hBMSCs appeared fibroblast-like and spindle-shaped and grew to 90% confluency (Fig. 1A). Alizarin Red S, Oil Red O and Alcian blue staining revealed that hBMSCs had the potential for osteogenic, adipogenic and chondrogenic differentiation, respectively (Fig. 1B). When the confluence of hBMSCs in passage three reached 90%, the surface markers assessed via flow cytometry showed that expression of the positive markers CD73 (99.9%), CD90 (99.1%) and CD105 (99.3%) was >95%, while that of the negative marker CD45 (0.2%) was <2% (Fig. 1C). These features suggest that the cells isolated from human bone marrow can be considered hBMSCs.

Ramírez-Flores et al (23) observed that T. gondii can secrete tubular structures composed of vesicles in vitro, which disperse into a single exosomes-like vesicle after centrifugation. In the present study, TgESPs extracted by centrifugation also contained numerous vesicles (~100-160 nm in diameter) with the morphological characteristics of exosomes, as observed under the transmission electron microscope (Fig. 1A). The isolate also contained some fine particulate matter, which may be other soluble proteins.

To address the effect of TgESPs on hBMSC proliferation, the CCK-8 assay was performed. The results indicated that 1, 30, 40 and 50 µg/ml TgESPs had no inhibitory effect on the proliferation of hBMSCs, while 10 and 20 µg/ml TgESPs had a positive impact on the growth of hBMSCs after 24 and 48 h of treatment compared to that in the control group (P<0.05) (Fig. 2B). Further investigation of the effect of TgESPs on hBMSC behavior revealed that low concentrations of TgESPs promoted cell adhesion, and both 1 and 10 µg/ml concentrations enhanced cell migration in hBMSCs (Fig. S1). These results indicate that TgESPs not only promote the proliferation of hBMSCs at low concentrations, but also enhance cell adhesion and migration.

To understand the effect of TgESPs on the osteogenic differentiation of hBMSCs, the effects of 0, 1, 10 and 20 µg/ml TgESPs on osteogenic differentiation of hBMSCs were examined using alizarin red staining. After 14 days of osteogenic differentiation induction, compared with that in the control group, TgESP treatment enhanced the osteogenic mineralization ability of hBMSCs. However, no obvious differences in mineralization were observed between 10 and 20 µg/ml (Fig. 2C); consequently, two concentrations, 1 and 10 µg/ml, were used in the subsequent experiments.

To elucidate the effect of TgESPs on osteogenic differentiation of hBMSCs in vitro, the expression levels of the osteogenesis-specific genes alkaline phosphatase (ALP), secreted phosphoprotein 1 (SPP1), RUNX2 and bone gamma-carboxyglutamate protein (BGLAP) were determined using RT-qPCR. The mRNA expression levels of ALP, SPP1, RUNX2 and BGLAP were significantly increased after seven days of TgESP treatment (P<0.05) (Fig. 2D). Next, the effect of TgESPs on the protein levels of Runx2, osteopontin (OPN) and osterix (OSX) was determined using western blotting and IF. The results showed that after ESP treatment, the protein expression of Runx2, OPN and OSX significantly increased in a concentration-dependent manner (P<0.05) (Fig. 2E-H). These findings indicate that TgESPs have a positive role in the osteogenic differentiation of hBMSCs.

The role of the Wnt/β-catenin signaling pathway in the osteogenic differentiation of BMSCs is crucial. To determine the mechanism by which TgESPs affect the osteogenic differentiation of hBMSCs, the expression of the Wnt/β-catenin signaling pathway was studied after TgESP treatment. First, the protein expression of LEF1, Dvl segment polarity protein 3 (DVL3), β-catenin and cyclin D1 was detected using western blotting. The results indicated that the protein expression of LEF1, DVL3, β-catenin and cyclin D1 was significantly increased after TgESP treatment (P<0.05) (Fig. 3A and B). This suggests that TgESPs may affect the Wnt/β-catenin signaling pathway during osteogenic differentiation of hBMSCs. After Icg-001 treatment, calcium deposition in hBMSCs was significantly reduced compared to that in the controls and addition of TgESPs restored the inhibitory effect of Icg-001 (Fig. 3C). In addition, the expression levels of osteogenic-specific genes and proteins, as well as β-catenin, were determined using RT-qPCR, western blotting and IF. The results suggested that the mRNA and protein expression of Runx2, OPN, OSX and β-catenin was significantly inhibited by Icg-001. The inhibition of these genes and proteins by Icg-001 was reversed due to the repressive effects of TgESP treatment (Fig. 3D-H).

Previous studies suggested that the cellular OCR and mitochondrial DNA content increase during osteogenic differentiation of hBMSCs (27,28), and that TgESPs can regulate host cell metabolism, particularly the TCA cycle (22). Therefore, it was hypothesized that TgESPs may also regulate the energy metabolism of hBMSCs during osteogenic differentiation. The results indicated that the OIM group with induced osteogenic differentiation had significantly higher OCR in hBMSCs than the LG-DMEM group without induced osteogenic differentiation. Upon addition of TgESPs, the basal, maximal, ATP production and respiration rates, as well as the spare respiratory capacity, were increased. However, no significant difference in OCR was observed among the induced osteogenic differentiation groups (Fig. 4A and B). This observation suggested that TgESPs had no significant effect on the mitochondrial OXPHOS of hBMSCs. Furthermore, the glycolysis rate of hBMSCs was measured and it was observed that the ECAR increased following hBMSC induction and that basal and compensatory glycolysis were enhanced after TgESP treatment (Fig. 4C and D). Subsequently, lactic acid production in cells was detected; the results indicated that hBMSCs produced significantly more lactic acid after 72 h of TgESP treatment (Fig. 4E). Next, to assess the biochemical basis of glycolysis, the gene and protein expression of several rate-limiting enzymes involved in critical steps of glycolysis was examined. The mRNA levels of PFKFB3, PFKFB4, enolase (ENO)3 and LDHA were significantly increased at three and seven days after osteogenic differentiation induction. In addition, TgESPs significantly increased the protein expression of PFKFB3 and LDHA in a concentration-dependent manner (Fig. 4F-H).

According to the above-mentioned results, TgESPs can enhance glycolysis during the osteogenic differentiation of hBMSCs. Therefore, in the present study, the effect of glycolysis on the osteogenic differentiation of hBMSCs was investigated in further detail. After treatment with 2-DG (an inhibitor of glycolysis), the ECAR and lactate production in hBMSCs were significantly inhibited; however, the ECAR and lactate secretion in hBMSCs were restored after simultaneous treatment with TgESPs and 2-DG (Fig. 5A and C). Quantification of the ECAR data showed that basic and compensatory glycolysis was able to reverse the inhibition effect of 2-DG after TgESP treatment (Fig. 5B). Alizarin red staining indicated that calcium deposition in the 2-DG group was much lower than that in the control group and TgESP group, but it increased again in the TgESPs+2-DG group as compared with that in the 2-DG group (Fig. 5D). Furthermore, RT-qPCR and western blotting results indicated that 2-DG inhibited the rate-limiting enzymes of glycolysis, while TgESPs reversed this inhibition (Fig. 5E, F and H). In addition, it was found that 2-DG inhibited the mRNA and protein expression of Runx2, OPN and OSX, while TgESPs reversed the inhibitory effect of 2-DG on osteogenic differentiation (Fig. 5E-I). These results suggest that TgESPs can affect glycolysis in hBMSCs and enhance their osteogenic differentiation ability.

The regulation of glycolysis in MSCs has been reported to be regulated by the Wnt/β-catenin signaling pathway (29). Therefore, it was hypothesized in the present study that TgESPs can affect glycolysis through the Wnt/β-catenin signaling pathway during the osteogenic differentiation of hBMSCs. Upon assessing the glycolytic rate, both basal and compensatory glycolysis were found to be significantly lower in the OIM+Icg-001 group than those in the OIM group (P<0.05). In addition, TgESP treatment was observed to reverse the inhibitory effect of Icg-001 on glycolysis (Fig. 6A and B). Furthermore, cellular lactate production showed the same trend as that described above (Fig. 6C). Next, the gene and protein expression of rate-limiting enzymes in the critical step of glycolysis were examined. The mRNA expression of PFKFB3, PFKFB4, ENO3 and LDHA was significantly reduced in the Icg-001 group (P<0.05), but addition of TgESPs increased the expression of these genes again (Fig. 6D). Furthermore, the inhibition of the PFKFB3 and LDHA proteins caused by Icg-001 was partially reversed by TgESPs (Fig. 6E and F). These results suggest that β-catenin is crucial in regulating downstream glycolysis and promoting the osteogenic differentiation of hBMSCs.