Within the tooth, there exists a soft tissue known as “pulp,” surrounded by dentin, a hard tissue composed of blood vessels and nerves [1]. In clinical dentistry, if the condition of the dental nerve is good or the nerve damage is weak, even if there is a cavity, a material called “mineral trioxide aggregate” (MTA) is used instead of nerve treatment [2]. MTA is recognized for its outstanding biocompatibility and high airtightness, making it a preferred option in caries treatment, with or without partial nerve removal [3]. However, the application of MTA becomes challenging when the actual tooth nerve is already damaged [4]. Although interest in dental pulp regeneration treatments has grown to overcome the limitations of nerve treatment, activating dental pulp stem cells (DPSCs) inside the tooth and commercializing them for dental pulp regeneration still pose significant challenges [5,6]. DPSCs, a type of adult stem cell existing in the dental pulp tissue, exhibit multipotency with an immunophenotype similar to bone marrow mesenchymal stem cells (BMSCs), osteoblasts, odontoblasts, muscle cells, and nerve cells [7,8]. DPSCs demonstrate a higher proliferation capacity than BMSCs. Furthermore, compared to other tissues, these stem cells are easily collected from patients, so they are considered ideal for tissue engineering approaches, such as tooth and nerve regeneration [9,10].

Chios gum mastic (CGM) is a resin extracted from the stems and leaves of Pistacia lentiscus var. Chia, a plant that grows only on the Greek island of Chios. It has been used for centuries in the Mediterranean and Middle East as a treatment for gastric and duodenal ulcers and as a food additive [11]. Over the past decade, CGM, recognized as a natural biocompatible material, has gained attention for its diverse pharmacological properties, including wound healing, antibacterial, anti-inflammatory, antioxidant, anti-ulcer, anti-diabetic, and anti-cancer activities [12,13]. Recent focus has shifted to the regenerative effects of CGM on tissues. Studies have revealed its role in promoting bone regeneration through the activation of protein kinase C, particularly in cells extracted from wisdom teeth demonstrating bone deterioration ability [14]. Ongoing research has delved into the anticancer properties of CGM, particularly in oral cancer and osteosarcoma. Furthermore, a previous study has highlighted that the combined use of CGM and low-level laser therapy promotes the differentiation of osteoblasts through the p38 mitogen-activated protein kinase (MAPK) pathway [15].

The growing interest in and demand for aesthetic and enduring dental procedures are driven by the recent super-aging of the domestic population and a desire for enhanced quality of life. Although research on odontogenic differentiation using DPSCs is being actively conducted at home and abroad, it remains in its early stages. Typically, for pulp regeneration, DPSCs are combined with scaffold materials and cytokines. However, bacterial infection poses a significant obstacle in this process, emphasizing the urgent need for antibacterial agents. CGM, a natural substance with excellent antibacterial, wound healing, and bone differentiation effects, emerges as a potential solution to overcome bacterial challenges within the oral cavity, fostering an environment conducive to dental pulp regeneration. Therefore, this study aims to validate the influence of CGM on the odontogenic differentiation of human DPSCs (hDPSCs) and to explore the potential of CGM, with its intrinsic antibacterial and differentiation-enhancing properties, as a promising new material in the realms of regenerative medicine and stem cell research applications.

Cell culture

hDPSCs (Catalog #: PT-5025) were purchased from Lonza Bioscience. hDPSCs were cultured and maintained in a Dulbecco’s Modified Eagle’s Medium (DMEM; Gibco) with 10% fetal bovine serum (FBS; Gibco) and 1% penicillin-streptomycin (Gibco) under 5% CO2 at 37°C. The cells used in this experiment were sub-cultured once every 2–3 days and did not exceed 20 passages.

CGM treatment

The CGM used in this study was procured from Sigma-Aldrich. To prepare for the experimental procedures, we created a stock solution of CGM at a concentration of 100 mM. This was achieved by dissolving CGM in ethanol, ensuring optimal solubility and stability for application in cell culture. This stock solution was then diluted to achieve final concentrations of CGM at 0, 1, 2.5, 5, 10, 25, and 50 μM. During the dilution process, for the 50 μM concentration of CGM, the final ethanol concentration was 0.05%. For the 1 μM concentration of CGM, the final ethanol concentration was 0.001%.

MTT assay

hDPSCs were seeded in a 96-well plate and incubated for 24 h to allow attachment to the well plate and stabilization. Subsequently, the cells were treated with CGM at concentrations of 0, 1, 2.5, 5, 10, 25, and 50 μM. This treatment was maintained for 24 to 96 h to assess the impact of CGM on cell viability. After the treatment period, the medium was removed from the well, and the cells were then treated with 100 μl of MTT solution (0.5 mg/ml) and incubated at 37°C for 4 h. The supernatant was then removed, and 100 ml of dimethyl sulfoxide (DMSO) was added to dissolve the formazan crystals formed on the cells. The absorbance of each sample was measured at 570 nm using a SpectraMax iD3 microreader (BioTek). Absorbance was measured in three independent experiments, and the results were expressed as mean ± standard deviation (SD).

Proliferation assay

The cell proliferation rate was determined using colony formation assay and transwell migration assay. hDPSCs were seeded in a six-well plate incubated for 24 h so that they could be attached to the well plate and stabilized. Each well was treated with 5 and 10 μM of CGM and allowed to stand for 7 days. Thereafter, the medium was removed from each well, washed once with phosphate-buffered saline (PBS), and fixed with methanol for 15 min. The cells were stained with 0.5% crystal violet dye for 30 min, washed three times with tap water, and sufficiently dried at room temperature. Then, the entire plate was photographed. Crystal violet-stained cells were dissolved in DMSO, and absorbance was measured at 570 nm using a SpectraMax iD3 microreader (BioTek). All experiments were performed three independent times. The membrane in the upper chamber of the transwell plate (Corning Costar) was coated with 30 μl of Matrigel and incubated for 4 h to solidify the gel. The cells were placed in Matrigel-coated transwell plates and treated with 5 and 10 μM of CGM for 48 h; the medium was serum free. The upper chamber was washed with PBS and fixed with 100% methanol for 10 min. The upper chamber was reacted with hematoxylin and eosin (H&E) staining reagents for 5 min each and then washed with PBS three times. After separating the membranes of the upper chamber, they were dehydrated by immersion in 70%, 80%, 90%, and 100% ethanol and mounted using malinol. The images were photographed and analyzed using Lionheart FX Automated.

Cell cycle analysis using flow cytometry

The cells were initially seeded in a 6-well plate and allowed to incubate for 24 h so that they could be attached to the well plate and stabilized. Following this, the cells underwent treatment with varying concentrations of CGM (0, 1, 2.5, 5, 10, 25, and 50 μM) for a duration of 48 h. The cells were suspended by trypsinization, washed once with PBS, and fixed in 70% ethanol for 24 h. The cells were harvested using a centrifuge and subsequently resuspended using a pipette in PBS containing 1% bovine serum albumin (BSA). Then, 10 μM of RNase was added to the cells and placed on ice for 30 min. Next, 10 μl of propidium iodide was added to each sample, and the analysis was conducted using the phycoerythrin (PE) wavelength in an fluorescence-activated cell sorting (FACS) analyzer. The G1/S/G2 phase was then systematically analyzed, quantified, and graphically represented for further examination.

Immunofluorescent staining

The hDPSCs were seeded into a Lap-Tek chamber plate, and the well plate was incubated for 24 h so that they could be attached to the well plate and stabilized. The next day, hDPSCs underwent treatment with CGM at concentrations of 0, 5, and 10 μM for 48 h. The cells were fixed with 4% paraformaldehyde (PFA) for 15 min and treated with a 0.2% Triton-X 100 (in PBS) solution for 10 min to permeate the cells with antibodies. As primary antibodies, Ki-67 and collagen type 1 were used. Primary antibodies (Cell Signaling Technology) were diluted at a 1:100 ratio in 1% BSA (in PBS) solution and applied to the cells for 24 h at 4°C. The cells were washed three times with PBS, and FITC-conjugated secondary antibody (1:100 dilution) was applied for 2 h at room temperature and then washed three times with PBS. Actin, a cytoskeleton, was stained with rhodamine phalloidin, a red fluorescent dye, and the nuclei were stained with DAPI. Each stained sample was observed, photographed, and analyzed using a Zeiss LSM 750 laser scanning confocal microscope.

Western blot assay

CGM-treated hDPSCs were harvested and lysed using RIPA buffer at 4°C for 2 h. The RIPA lysis buffer was prepared using the following ingredients: pH 7.6; 50 mM Tris–Cl, 300 mM NaCl, 0.5% Triton X-100, 2 μl/ml aprotinin, 2 mM PMSF, and 2 μl/ml Leupeptin. Proteins extracted from cells were quantified using a Bradford assay (Bio-Rad), and the amount of protein in each sample was set to 20 μg. Each sample was loaded by electrophoresis using a 10% SDS-PAGE gel, and then the protein loaded onto the gel was transferred to a PVDF membrane (Millipore). Membranes were blocked with 5% non-fat dry milk for 1 h, and primary antibodies (Cyclin A, Cyclin E, CDK2, GSK3β, p-GSK3β, Wnt3a, β-catenin, Axin1, Ki67, and β-actin; Cell Signaling Technology) were applied at 1:1,000 dilution overnight in the refrigerator. The membranes were washed 5 times for 10 min each with PBS, and a secondary antibody (anti-mouse-HRP conjugated and anti-rabbit-HRP conjugated secondary antibody; 1:5,000 dilution; Cell Signaling Technology) was applied for 1 h at room temperature. Membranes were then washed 5 times for 10 min each with PBS, and SuperSignal West Femto enhanced chemiluminescent substrate was applied to detect protein expression. Protein expression was documented using the ImageQuant LAS 500 chemiluminescence system (GE Healthcare).

RNA isolation and quantitative polymerase chain reaction (qPCR)

The hDPSCs were seeded in a 6-well plate and allowed to incubate for 24 h. Subsequently, the medium was replaced with a new differentiation medium containing CGM, and the cells were cultured for 48 h and 14 days. Total RNA (10 ng) was isolated from each sample using the RNeasy Mini Kit (Qiagen Inc.). The RNA from each sample was synthesized into cDNA using an M-MLV cDNA Synthesis Kit (Enzynomics) according to the manufacturer’s protocol and as previously described [16]. Genes were labeled using the SYBR Green Kit (Applied Biosystems), and gene expression was quantified by amplification at 40 cycles with the ABI 7500 Real-Time PCR Detection System (Applied Biosystems). The primers used in this experiment are shown in Table 1. Gene expression was calculated using the comparative Ct method, with the housekeeping gene GAPDH as a reference, to calculate the Ct values of the samples.

Table 1 . Sequences of primers.

Target gene		Primer sequence (5’ to 3’)
Wnt3a	Forward	TTTGGTGGGATGGTGTCTCG
	Reverse	ACCAGCATGTCTTCACCTCG
Axin	Forward	ACAGGATCCGTAAGCAGCAC
	Reverse	GGAATGTGAGGTAGGGGCAC
β-catenin	Forward	CTGAGGAGCAGCTTCAGTCC
	Reverse	GGCCATGTCCAACTCCATCA
ALP	Forward	CGGGCACCATGAAGGAAA
	Reverse	GGCCAGACCAAAGATAGAGTT
BSP	Forward	GGAACGGACATTCGGTCCTT
	Reverse	AGTCCGTCTAAGAAGCACGC
DMP-1	Forward	TGGTCCCAGCAGTGAGTCCA
	Reverse	TGTGTGCGAGCTGTCCTCCT
DSPP	Forward	GGGAATATTGAGGGCTGGAA
	Reverse	TCATTGTGACCTGCATCGCC
OP	Forward	CTCACACTCCTCGCCGTATT
	Reverse	GCTCCCAGCCATTGATACAG
RUNX2	Forward	CCAGATGGGACTGTGGTTAC
	Reverse	ACTTGGTGCAGAGTTCAGGG
GAPDH	Forward	GGTCACCAGGGCTGCTTTTA
	Reverse	CCCGTTCTCAGCCATGTAGT

Alizarin red staining and alkaline phosphatase (ALP) activity

hDPSCs were seeded in 24-well plates and allowed to incubate for 24 h. Subsequently, the medium was replaced with a new differentiation medium containing CGM. This odontogenic differentiation medium was composed of MEM alpha medium (HyClone) supplemented with β-glycerophosphate (10 mmol; Sigma-Aldrich) and l-ascorbic acid (0.2 mmol; Sigma-Aldrich), providing the necessary conditions for odontogenic differentiation. The cells were cultured in this medium for 7–14 days, with the media changed every 2 days. Both cells were fixed with ice-cold 70% ethanol and stained with alizarin red S to detect calcification. For quantification, the cells were stained with ethylpyridium chloride and transferred to a 96-well plate. Absorbance was measured at 540 nm using a SpectraMax iD3 microreader (BioTek). In the analysis of alizarin red S staining, absorbance was measured for both the control group and the CGM-treated groups. To quantify the relative calcification, we normalized the absorbance values of the CGM-treated groups against those of the control group. This was done by setting the absorbance value of the control group as the baseline (normalized to 1). The relative absorbance for the CGM-treated groups was then calculated using the following formula:

To measure ALP activity, we treated CGM under the same conditions as alizarin red staining and cultured the cells for 7 and 14 days. Following treatment, the cell lysates were incubated with 10 mM Tris–HCl (pH 7.6) buffer containing 0.1% Triton X on ice. Then, 50 μg of cellular fraction proteins were incubated with p-nitrophenol for 30 min at 37°C in the dark. The reaction was quenched by adding sodium hydroxide. The p-nitrophenol levels and ALP enzyme activity were measured by monitoring the SpectraMax iD3 microreader (BioTek) at a 410 nm excitation-emission wavelength. The ALP activity of each sample was normalized by the protein concentration.

Statistical analysis

All data were expressed as mean ± SD. The figures presented herein were derived from at least three independent experiments. Statistical analysis was performed using one-way analysis of variance (ANOVA) and Dunnett’s comparison. Differences with a probability (p-value) of less than 0.05 were considered statistically significant.

Low doses of CGM are not cytotoxic and activate DNA synthesis in the cell cycle progression in hDPSCs

An MTT assay was conducted to assess cytotoxicity based on concentrations of CGM. Varying concentrations of CGM (0, 1, 2.5, 5, 10, 25, and 50 μM) were applied to hDPSCs for 24–96 h, and cell viability was confirmed. It was confirmed that the lower concentration of CGM (1 to 10 μM) did not affect the hDPSC cells at all over time but rather increased the viability of the cells. In particular, in the CGM 5 μM (48 h, 122.5%; 72 h, 138.9%; 96 h, 157.7%) and 10 μM (48 h, 131.1%; 72 h, 140.6%; 96 h, 163.5%) treatment groups, the increase in cell viability after 48 h showed statistical significance. However, at relatively high concentrations of 25 μM (48 h, 98.3%; 72 h, 93.5%; 96 h, 67.0%) and 50 μM (48 h, 97.9%; 72 h, 85.5%; 96 h, 58.2%) of CGM, the cell viability decreased after 96 h of treatment (Fig. 1). Hence, concentrations of 5 and 10 μM CGM are proposed to stimulate cell proliferation without causing cytotoxic effects. Subsequently, we conducted FACS analysis to explore whether the heightened cell viability in hDPSCs due to CGM treatment was associated with cell cycle progression. In hDPSCs, treatment with less than 10 μM CGM increased the G1 phase (cell growth phase) and S phase (DNA synthesis and DNA replication phase) cell numbers; however, 25 μM and 50 μM CGM decreased the number of G1 and S phases (Fig. 2A, B). In addition, we confirmed the expression of cyclin proteins (G1, Cyclin A; G1/S, Cyclin E) related to G1 and G1/S phases and their partner CDK2 in CGM-treated hDPSCs through Western blot (Fig. 2C). Treatment with CGM at concentrations ranging from 1 to 10 μM significantly elevated the expression of Cyclin A, Cyclin E, and CDK2 in hDPSC cells. However, at concentrations exceeding 25 μM, the expression of these markers decreased. Collectively, these findings suggest that CGM fosters cell growth and DNA synthesis at concentrations of 10 μM or lower, maintaining elevated cell viability, even with prolonged exposure.

Figure 1. Cytotoxic effect of Chios gum mastic (CGM) on human dental pulp stem cells (hDPSCs). hDPSCs were treated with CGM at concentrations of 0–50 μM for 24–96 h. Cell viability was expressed as % based on the 24-h control group (0 μM). The results were expressed as mean ± SD and were derived from three independent experiments (*p < 0.05, **p < 0.01).

Figure 2. Effects of Chios gum mastic (CGM) on the cell cycle progression of human dental pulp stem cells (hDPSCs). Cells were treated with 0, 1, 2.5, 5, 10, 25, and 50 μM CGM for 48 h. (A) After staining the cells with propidium iodide, the cell cycle was analyzed using FACS. (B) The percentage ratios of G1 (green symbols), S (blue symbols), and G2 (purple symbols) of the histogram shown in A are shown as line category graphs. (C) The protein expression of cell cycle-related factors (Cyclin A, Cyclin E, and CDK2) was confirmed through Western blot analysis. β-actin was used as a loading control.

CGM promotes the activation of cell proliferation through the Wnt/β-catenin signaling in hDPSCs

Based on previous findings, we validated the notable impact of 5 and 10 μM CGM treatment on hDPSCs, particularly after 48 h. Therefore, we performed a transwell migration assay and a cell proliferation assay using crystal violet staining to confirm the effects of 5 and 10 μM CGM on cell proliferation and migration. The cell migration (Fig. 3A) of hDPSCs was assessed over a period of 48 h, and cell proliferation (Fig. 3B) was monitored for 7 days under treatment with 5 and 10 μM of CGM. CGM demonstrated clear cell migration and proliferation compared to the untreated group (control), with the 10 μM CGM treatment group showing approximately twice the cell migration and proliferation compared to the 5 μM CGM treatment group (Fig. 3A, B). The Wnt signaling pathway is known to play divergent roles during development, normal homeostasis, and disease, and its activation controls both cell proliferation and differentiation [17]. Thus, we verified the influence of CGM treatment on the Wnt signaling pathway in hDPSCs. The gene expressions of Wnt3a, β-catenin, and AXIN were significantly increased in the CGM 5 and 10 μM treatment groups. Protein expression was analyzed through Western blot, which revealed a marked increase in wnt3a with the 10 μM CGM treatment group. For GSK3β and p-GSK3β, a progressive increase in protein levels was observed from 1 μM to 5 μM CGM treatments, while a decrease was noted at concentrations above 10 μM CGM. AXIN protein expression was also significantly increased in the 5 and 10 μM CGM treatment groups (Fig. 3E). Immunofluorescent staining and Western blot analysis were used to examine the expression of Ki-67, a marker of cell proliferation, and a considerable increase was seen in the 5 and 10 μM CGM treatment groups (Fig. 3D, E). These results strongly indicate that treatment with 10 μM CGM stimulates cell proliferation in hDPSCs through the activation of the Wnt signaling pathway.

Figure 3. Effect of Chios gum mastic (CGM) on cell proliferation in human dental pulp stem cells (hDPSCs). (A) Using a transwell chamber, hDPSCs were treated with 5 and 10 μM of CGM, and cell migration was confirmed for 48 h (×40). (B) To determine the cell proliferation rate by CGM, hDPSCs were treated with 5 and 10 µM of CGM for 7 days. Cell proliferation was confirmed by staining the cells with 1% crystal violet (×40). (A, B) The cell migration and proliferation rates were expressed based on the control group and displayed as a histogram. The results were expressed as mean ± SD and were derived from three independent experiments (*p < 0.05, **p < 0.01). (C) mRNA expression patterns of Wnt3a, β-catenin, and AXIN in hDPSC cells following CGM treatment. Cells were treated with 0, 1, 2.5, 5, 10, 25, and 50 μM CGM for 48 h. The results were expressed as mean ± SD and were derived from three independent experiments (*p < 0.05, **p < 0.01, ***p < 0.001). (D) After treating the cells with 0, 5, and 10 μM CGM for 48 h, the expression of Ki-67 was observed using a confocal microscope through immunofluorescence. Actin was stained with rhodamine phalloidin (red), Ki-67 was stained with Alexa 488 (green), and nuclei were stained with DAPI (blue) (×400). (E) The protein expression of Wnt signal-related factors (Wnt3a, β-catenin, GSK3β, p-GSK3β, AXIN1, and Ki-67) was confirmed by Western blot analysis. β-actin was used as a loading control.

CGM increases the mineralization and odontogenic differentiation of hDPSCs

Subsequently, we investigated whether CGM influences the expression of collagen type I, a pivotal protein within the extracellular matrix encompassing the pulp matrix. To examine this, hDPSCs were exposed to 5 and 10 μM CGM, and after 48 h, collagen expression was evaluated through immunofluorescence (green fluorescence) using a confocal microscope. In the CGM-treated groups, a notable increase in cytoplasmic collagen expression was observed (Fig. 4A). Our investigation extended to assessing the impact of CGM on hDPSC mineralization during odontogenic differentiation. This assessment involved alizarin red staining and analysis of ALP activity. Following treatment with 5 μM and 10 μM CGM, hDPSC cells were cultured in a differentiation-inducing medium for 7 and 14 days. Remarkably, in the 14-day group, hDPSCs treated with 5 μM (1.48) and 10 μM (1.98) CGM exhibited significantly heightened ALP activity ratios (Fig. 4B). Using alizarin red staining, we confirmed the mineralization of hDPSCs. Upon treatment with 5 μM and 10 μM CGM, substantial deep red calcium deposition was observed in the 10 μM CGM group after 7 days and in both the 5 μM and 10 μM CGM groups after 14 days (Fig. 4C). Quantitative analysis of the staining unveiled a marked increase in the mineralization ratio within the 14-day CGM-treated group (5 μM: 4.2, 10 μM: 6.4) (Fig. 4D). Lastly, we scrutinized alterations in osteogenic-related genes within cells subjected to CGM treatment during hDPSC differentiation. After treating hDPSCs with CGM and culturing them in a differentiation medium for 7 days, qPCR analysis was conducted. The gene expression of osteogenic markers, including ALP, Bone Sialoprotein (BSP), Dentin Sialophosphoprotein (DSPP), Dentin Matrix Protein 1 (DMP1), Osteopontin (OP), and Runt-related transcription factor 2 (RUNX2), exhibited noteworthy elevation, particularly in the CGM 10 μM treatment group (Fig. 4E). These results indicate that CGM promotes the expression of collagen type I from the proliferation stage of hDPSCs and stimulates osteogenic genes after differentiation, leading to mineralization.

Figure 4. Osteogenic differentiation effect of Chios gum mastic (CGM) in human dental pulp stem cells (hDPSCs). (A) After treatment with 0, 5, and 10 μM of CGM in hDPSCs for 48 h, the expression of collagen type I was observed using confocal microscopy through immunofluorescence. Actin was stained with rhodamine phalloidin (red), collagen type I was stained with Alexa 488 (green), and nuclei were stained with DAPI (blue) (×400). (B) To determine the cell alkaline phosphatase (ALP) activity by CGM, hDPSCs were treated with 5 and 10 µM of CGM for 7 and 14 days. The ALP activity ratio was calculated in relation to the control group (for each 7 and 14 days), and the results were presented as a histogram. Results were expressed as mean ± SD and were derived from three independent experiments (*p < 0.05, **p < 0.01, ***p < 0.001). (C, D) To measure ALP activity, we treated CGM under the same conditions as alizarin red staining and cultured the cells for 7 and 14 days. The color development observed after alizarin red staining was quantified based on the control group (for each 7 and 14 days) and expressed accordingly. Results were expressed as mean ± SD and were derived from three independent experiments (*p < 0.05, ***p < 0.001) (×40). (E) In order to confirm the effect of CGM on the odontogenic differentiation process at the mRNA level, hDPSCs were treated with 5 and 10 µM of CGM for 7 days, and the expression of bone and dentine-related factors (ALP, BSP, DMP-1, DSPP, OP, and RUNX2) in the cells was quantified by qPCR. The results were expressed as the mean ± SD. Data were derived from three independent experiments (**p < 0.01, ***p < 0.001).

The present study has successfully demonstrated that CGM exhibits low cytotoxicity in hDPSCs while also significantly promoting cell proliferation and enhancing odontogenic differentiation. The pursuit of alternative treatments devoid of side effects for various disease models is a growing area of research. Notably, there has been a surge in the investigation of herbal medicine owing to its potential to offer effective remedies for various ailments [18,19]. For centuries, materials extracted from herbs or plants have been used for regenerative treatments, and recently, studies have been published that provide positive expectations for tooth regeneration from these natural materials [20-22]. Scientific interest in CGM has increased over the past decade, and several studies have reported CGM’s diverse biomedical and pharmacological properties, including bacterial eradication, peptic ulcer relief, dental plaque inhibition, cancer prevention, and cardiovascular protection [23]. Moreover, a prior study from our research group underscored CGM’s capacity for osteogenesis through experiments involving hFOB 1.19 and MC3T3-E1 osteoblasts [15]. Building upon this foundation, our current research aimed to corroborate the effects of CGM on hDPSCs by focusing on cell proliferation and the facilitation of odontogenic differentiation. This exploration is rooted in CGM’s established ability to foster bone regeneration and differentiation, as evidenced by previous studies.

To assess the potential toxic effects of CGM on hDPSCs, cell viability was evaluated across different concentrations and durations. At concentrations of 10 μM or lower, CGM exhibited no cytotoxicity; conversely, it displayed a pattern of promoting cell proliferation. Even when exposed to higher concentrations of CGM, 25 μM or beyond, after a 96-h treatment, low cytotoxicity was observed (Fig. 1). It is recognized that the cytotoxic effects of CGM can vary significantly based on the specific cell type. Among the tested cell lines, the promyelocytic leukemia cell line HL-60 exhibited the highest sensitivity to CGM-induced cytotoxicity. This sensitivity was followed by myeloblastic leukemia lines (ML-1, KG-1), erythroleukemia line (K-562), oral squamous cell carcinoma lines (HSC-2, HSC-3, HSC-4), hepatocellular carcinoma line (HepG2), and glioblastoma lines (T98G, U87MG). Notably, normal oral cells, including gingival fibroblasts, dental pulp cells, and periodontal ligament fibroblasts, demonstrated a higher degree of resistance to these cytotoxic effects [13]. The Wnt/β-catenin signaling pathway plays a crucial role in orchestrating various facets of cellular behavior throughout development. It achieves this by promoting cell cycle progression and proliferation through the transcriptional upregulation of target genes, including cyclins [24]. This pathway ensures that cells undergo two key processes during division: first, the replication of DNA, and second, the separation of replicated chromosomes into individual cells. The former process is known as the interphase, while the latter is called mitosis. The interphase cell cycle consists of stages like cell growth (G1 phase), DNA synthesis (S phase), and preparation for mitosis (G2 phase) [25]. This sequential process of cell growth, replication, and division is governed by complexes like cyclin-dependent kinases (CDKs), proline-directed serine/threonine kinases, and regulatory cyclin subunits [26]. In the group treated with CGM in hDPSC cells, the number of cells entering the G1/S and S phases increased. Additionally, there was an increase in the protein expression of cyclin A, cyclin E, and their CDK counterpart, CDK2, which function in the G1 and S phases. However, there was a decrease in cells treated with CGM at concentrations higher than 25 μM. The Wnt signaling pathway can be divided into canonical and noncanonical pathways. The canonical Wnt pathway focuses on cell growth and development, whereas the noncanonical pathway emphasizes cell movement and structural organization [27]. In the canonical Wnt pathway, odontogenic differentiation is influenced by gene transcription regulation mediated by β-catenin [28]. When Wnt ligands bind to Frizzled receptors, this leads to the stabilization and accumulation of β-catenin in the cytoplasm and its subsequent translocation into the nucleus. In the nucleus, β-catenin acts as a coactivator with TCF/LEF transcription factors, leading to the transcription of specific genes that promote odontogenic differentiation. This pathway plays a critical role in the development and regeneration of dental tissues by regulating the differentiation of DPSCs into odontoblasts, the cells responsible for forming dentin [29]. β-catenin is an important component of the centrosome, which is the microtubule organizing center during mitosis, forming the spindle apparatus. In addition to β-catenin, other proteins, such as AXIN1, AXIN2/Conductin, APC, and GSK3, also participate in the formation of the spindle apparatus by binding to the microtubules [30-32]. In this study, the effect of CGM on cell proliferation in hDPSCs was examined. The expression of genes or proteins involved in the Wnt signaling pathway, such as Wnt3a, β-catenin, AXIN, and GSK3β, displayed significant increases at CGM concentrations of 10 μM or lower. These findings suggest that a low concentration of CGM stimulates Wnt signaling, subsequently regulating cell-cycle proteins and thereby promoting cell proliferation in hDPSCs (Figs. 2 and 3).

In this study, we investigated the mineralization potential of hDPSCs by analyzing alizarin red staining and ALP activity after treating them with CGM. Notably, CGM exhibited a remarkable ability to enhance the osteogenic differentiation of hDPSCs. Collagen type I plays a pivotal role as a fundamental fibrous protein within the extracellular matrix of human dentin. It is among the initial components synthesized during dentin formation, contributing significantly to dentin mineralization [33]. Our findings further solidified that CGM treatment significantly upregulated the expression of collagen type I within hDPSCs. Furthermore, ALP activity plays a crucial role in the development of mineralized tissues [34]. With its distribution pattern similar to that of all hard tissues, ALP is found within blood vessels and cellular membranes of cells, contributing to hard tissue formation [35]. Consequently, our findings suggest that CGM not only boosts the expression of collagen type I within hDPSCs but also enhances intracellular ALP activity, ultimately fostering the mineralization process of hDPSCs (Fig. 4A–D).

We also investigated whether odontogenic markers were activated by the CGM treatment-induced differentiation and mineralization of hDPSCs. Mature dentin predominantly consists of about 70% minerals, 20% organic matrix (mainly collagen), and 10% water [36]. Specifically, within the organic matrix, around 90% comprises collagen types I and III, accompanied by some lipids and noncollagenous components [37]. Matrix proteins present in bone are also found in dentin, and some matrix proteins present in dentin are also present in bone [38]. DSPP and DMP-1, which are matrix proteins of the noncollagenous matrix, fill the space between collagen fibers and accumulate along the dentin tubules [39]. In this study, we utilized qPCR to quantify the expression patterns of odontogenic markers (DMP-1 and DSPP) and osteogenic markers (OP, RUNX2, and BSP). CGM treatment significantly upregulated the gene expression of DMP-1, DSPP, OP, RUNX2, and BSP within hDPSCs.

In our study, we observed that treatment with a specific concentration of CGM, 10 μM, enhanced odontogenic differentiation of DPSCs. This enhancement was evidenced by the upregulation of Wnt3a, β-catenin, and AXIN, along with the downregulation of GSK3β. While the Wnt/β-catenin pathway plays a crucial role in stem cell self-renewal and differentiation, its effects on osteogenic differentiation in DPSCs have remained a subject of debate [40]. Most studies have reported that Wnt/β-catenin signaling increases bone mass and promotes osteogenic differentiation in DPSCs [41]. However, Scheller et al. [42] reported that the activation of Wnt/β-catenin and overexpression of β-catenin inhibit bone formation in DPSCs. Furthermore, Zhou et al. [43] demonstrated that although the Wnt/β-catenin signaling pathway actively participates in various biological processes, its activity is suppressed during the final mineralization process. Aberrant activation of the Wnt/β-catenin pathway at the terminal stage disrupts the mineral crystal structure and reduces the bonding force between minerals and the organic matrix. One possible explanation for these discrepancies is that our results primarily focused on early-stage changes in Wnt/β-catenin signaling, while only observing changes in odontogenic differentiation and bone- and dentin-related factors (ALP, BSP, DMP-1, DSPP, OP, and RUNX2) at the final stage.

Taken together, our findings indicate that hDPSCs exhibited minimal sensitivity to CGM toxicity. Furthermore, CGM treatment at low concentrations induced cell proliferation by activating cell-cycle proteins. Additionally, our study demonstrated that CGM stimulates ALP activation by augmenting the expression of collagen type I, a representative matrix protein in dentin. This activation, in turn, triggers odontogenic and bone differentiation markers, ultimately fostering the mineralization of hDPSCs. Although further research is necessary to unveil the diverse molecular mechanisms and CGM-induced odontogenic differentiation in hDPSC cells, our study’s outcomes suggest that CGM holds substantial potential as a novel natural agent. Beyond tooth regeneration, this potential could extend to applications within the realm of regenerative medicine.

The authors extend their deepest appreciation to all the participants for their invaluable support in this study.

This study was financially supported by a National Research Foundation of Korea (NRF) grant funded by the Korean government (No. NRF-2019R1A2C108405712).

The authors declare no conflicts of interest.