Human fetal osteoblasts (hFOB 1.19) were obtained from the American Type Culture Collection (CRL-11372, Rockville, MD, USA) and maintained in Dulbecco's modified Eagle's medium-Ham's F-12 (1:1) basal medium supplemented with 10% fetal bovine serum (FBS) and 1% penicillin/streptomycin/glutamine (Gibco, 15140-122, Invitrogen Corporation, Grand Island, NY, USA) in a standard cell-culture incubator at 37°C and 5% CO₂. Differentiation was induced by transfer to osteogenic medium (OM) containing 100 µM ascorbic acid, 2 mM β-glycerophosphate, and 10⁻⁷ M dexamethasone, as previously described [21].

Human PDLCs and cementoblasts [22], immortalized by transfection with the telomerase catalytic subunit of the human telomerase reverse transcriptase, were kindly provided by Professor Takashi Takata (Hiroshima University, Hiroshima, Japan). Cells were cultured in α-minimum essential medium (α-MEM; Invitrogen Corporation) supplemented with 10% FBS, 100 U/mL of penicillin, and 100 μg/mL of streptomycin in a humidified atmosphere of 5% CO₂ at 37°C.

Primary osteoblasts were enzymatically isolated from the calvaria of 1-day-old ICR mice (Samtako, Osan, Korea). Briefly, the calvaria were aseptically dissected, and the fragments were incubated in phosphate-buffered saline (PBS) solution with 0.2% collagenase-dispase solution (Sigma-Aldrich, St. Louis, MO, USA) for 2 hours at 37°C. The cells were dissociated from the bone fragments, counted, and plated at 4×10⁴ cells/cm². The cells were grown in α-MEM supplemented with 10% FBS, 100 U/mL of penicillin, and 100 μg/mL of streptomycin in a humidified atmosphere of 5% CO₂ at 37°C.

All animal experimental procedures in the current study were approved by the Institutional Animal Care and Use Committee of Kyung Hee University (approval number: KHMC-IACUC 2015-002). Primary human PDLCs were cultured using an explant technique, as described previously [23]. Briefly, impacted third molars were obtained from 3 healthy patients of the Department of Oral and Maxillofacial Surgery, Kyung Hee University Dental Hospital, Seoul, Korea. The experiments were performed with approval from the Ethical Committee of Kyung Hee University Dental Hospital, and informed written consent was obtained from all patients.

Freshly extracted teeth were immediately placed in PBS supplemented with antibiotics (100 U/mL of penicillin, 100 μg/mL of streptomycin, and 0.25 μg/mL of Fungizone). In order to culture PDLCs, the PDL was carefully removed from the middle third of the root by a scalpel. Tissues were cultured in Dulbecco's modified Eagle's medium supplemented with 10% FBS and antibiotics. Cultures were maintained at 37°C in a humidified atmosphere of 5% CO₂ and 95% air. Confluent cells were detached with 0.25% trypsin and 0.05% EDTA for 5 minutes, and aliquots of separated cells were subcultured. Cell cultures between the third and eighth passages were used in this study.

A neodymium (Nd₂Fe₁₄B) disc magnet (1 mm thick and 15.0 mm in diameter, Qingdao Qiangsheng Magnets Co., Ltd., Shandong, China) and 4-well plastic culture plates were used to create an SMF exposure system (Figure 1A). A magnet was placed below the wells horizontally, and the north (N) side of the magnet was oriented towards the well. The desired field intensities (3, 15, and 50 mT) were established by controlling the distance between the magnet and the culture plates, as previously described [17]. The distance was fixed by the space between the magnet and bottom wall of the well. We measured the SMF intensity with a Gauss meter (TM-701, Kanetec, Tokyo, Japan) that could detect either axial or transverse fields. To eliminate the interference of adjacent SMFs, we attached only 1 magnet disc on 1 culture plate and provided adequate space between adjacent culture plates (at least 30 mm).

Cell viability was confirmed using a 3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium (MTT) assay kit (CellTiter 96 AQueous One Solution, Promega, Madison, WI, USA) following the protocol suggested by the manufacturer.

Intracellular and soluble ALP, a marker of osteoblast activity, was assayed by the release of p-nitrophenol from p-nitrophenylphosphate. Cells were washed with PBS and sonicated with a cell disruptor. ALP activity was measured using p-nitrophenyl phosphate (final concentration, 3 mM) in 0.7 M 2-aminomethyl-1-propanol (pH 10.3) and 6.7 mM MgCl₂. Absorbance was measured at 415 nm using an enzyme-linked immunosorbent assay reader (Beckman Coulter, Fullerton, CA, USA).

Alizarin red staining was used to detect mineralization. Cells were fixed in 70% ice-cold ethanol for 1 hour and rinsed with distilled water. Fixed cells were stained with 40 mM Alizarin red (pH 4.2) for 10 minutes with gentle agitation. All Alizarin red staining images were photographed.

Total RNA was extracted using Trizol reagent (Life Technologies, Gaithersburg, MD, USA). Then, 1 mg of RNA was reverse transcribed using oligo (dT) 15 primers (Roche Diagnostics, Mannheim, Germany) and AccuPower RT PreMix (Bioneer Corporation, Daejeon, Korea). Thereafter, the reverse transcriptase-generated DNA (2–5 mL) was amplified. Primer sequences are detailed in Table 1. The polymerase chain reaction (PCR) products were subjected to electrophoresis on 1.5% agarose gels, and stained with ethidium bromide.

Aliquots of 1×10⁶ cells were solubilized in ice-cold 1% Triton X-100 lysis buffer (Boston BioProducts, Inc., Ashland, MA, USA). After 30 minutes on ice, the lysates were clarified by centrifugation. Proteins (20 µg) were resolved by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (10% acrylamide), transferred to nitrocellulose membranes, and probed with specific antibodies (diluted to 1:1,000), followed by incubation with secondary horseradish peroxidase-conjugated antibody (1:5,000). Products were detected using an enhanced chemiluminescence system (Amersham-Pharmacia, Piscataway, NJ, USA).

To measure calcium content, cells were harvested by centrifugation at 13,500 rpm for 1 minutes, resuspended in lysis buffer (0.1% Triton X-100, Boston BioProducts, Inc.) and sonicated on ice for 1 minute to destroy the cell membranes. The resulting supernatant was used for calcium measurements made with a QuantiChrom™ Calcium Assay Kit (DICA-500, BioAssay Systems, Hayward, CA, USA). Calcium content was estimated by measuring absorbance at 612 nm in a microplate reader.

Cells were fixed with freshly prepared 3% paraformaldehyde for 15 minutes and permeabilized with 0.5% Triton X-100 (Boston BioProducts, Inc.) for 15 minutes. After 1 hours of incubation with 10% normal goat serum/PBS, cells were incubated with anti-p65 antibody (Santa Cruz Laboratories, Santa Cruz, CA, USA) diluted 1:50 in PBS for 2 hours, washed and incubated with fluorescein isothiocyanate (FITC)-conjugated immunoglobulin G, and diluted at 1:100 in PBS for 1 hour. In order to identify the nuclei, the FITC-labeled samples were counterstained with 10 μg/mL of propidium iodide for 2 minutes. To acquire the immunofluorescence images, a confocal microscope (Cell Voyager, Yokogawa, Japan) was used.

Descriptive statistics (mean±standard deviation) for each parameter were evaluated in all the groups. The data distributions satisfied normality as tested by the Shapiro-Wilk test, meaning that intergroup comparisons could be made by using 1-way analysis of variance and the Duncan multiple range test. P values <0.05 were considered to indicate statistical significance.

To determine the cytotoxic potential of SMFs, its effect on viability of the human osteoblasts was evaluated. Up to a magnitude of 50 mT, no cytotoxic effects could be observed using the MTT assay (Figure 1B). To determine the effects of SMFs on osteoblastic differentiation, we evaluated its effects on ALP activity (an early marker of osteoblastic differentiation) and mineralization. ALP activity was significantly higher (P<0.05) and calcified nodules more frequent after SMF treatment than when the cells were simply maintained in normal osteogenic supplement (OS) or OM (Figure 1C). Osteoblasts exposed to an SMF of 15 mT had significantly higher ALP levels than the other groups (P<0.05). Because the endpoint of osteoblast formation is the production of a calcified, mineralized matrix, calcium content was measured as an indicator of osteodifferentiation. The SMF-exposed osteoblasts had significantly larger mineralized nodules (Figure 1D) and extracellular calcium deposits than their control counterparts (Figure 1E) (P<0.05). The largest number of mineralized nodules and the greatest calcium content was observed in the 15 mT group. In order to explore the effects of SMFs on the osteoblastic differentiation of osteoblasts, we examined the expression of specific markers of osteoblastic differentiation. Treating the osteoblasts with SMFs increased the expression of mRNA encoding the OS- and OM-induced differentiation markers runt-related transcription factor 2 (Runx2), osteopontin (OPN), and osteocalcin (OCN) (Figure 1F and G). Maximal upregulation of the osteoblast marker genes was observed at 15 mT.

Since SMFs of 15 mT maximally stimulated the osteoblastic differentiation of human osteoblasts, 15 mT was used in further experiments. Cytotoxicity of PDLCs and cementoblasts in response to SMFs was not observed on days 3, 7, or 14 (Figure 2A and B). To investigate whether SMFs stimulated PDLCs and cementoblasts to differentiate toward a cementoblast/osteoblast phenotype, we measured ALP activity and the expression of osteoblast- and cementum-related genes, calcium content, and calcified nodule formation. Treatment with SMFs enhanced ALP activity and mineralized nodule formation in PDLCs and cementoblasts, compared with OS and OM controls (Figure 2C-F). Similar increases in calcium concentration were also observed in SMF-treated PDLCs and cementoblasts (Figure 3A and B). In addition, SMFs upregulated the mRNA expression of osteoblast markers in PDLCs, including Runx2, OPN, and OCN (Figure 3C), as well as the expression of cementoblast markers, such as cementum protein 1 (CEMP-1) and cementum-derived attachment protein (CAP), in cementoblasts (Figure 3D).

To further assess the osteoblastic activity of the SMFs, we examined primary cultured osteoblasts and PDLCs. Primary cultured PDLCs and osteoblasts showed no cytotoxicity in response to SMFs, but did show increased ALP activity and mineralization (Figures 4A-F, 5A and B). In addition, the treatment of primary cultured PDLCs and osteoblasts with SMFs increased the expression of mRNA encoding the differentiation markers Runx2, OPN, and OCN (Figure 5C and D).

To determine the basis of the effects of SMFs on osteoblast differentiation, their effects on Wnt/β-catenin, MAPK, and NF-κB signaling were examined. As shown in Figure 6A and B, SMFs increased Wnt1 and Wnt3a expression, the phosphorylation of glycogen synthase kinase-3β (GSK-3β), and total β-catenin protein levels in osteoblasts. Moreover, treatment with SMFs increased OM-induced p38 and JNK phosphorylation, but not that of ERK (Figure 6C). In addition, it enhanced the OM-induced nuclear translocation of p65 (Figure 6D). Immunocytochemical observations showed that the nuclear content of NF-κB p65 increased markedly in SMF-treated osteoblasts compared with the OM control (Figure 6E).

To confirm the involvement of Wnt/β-catenin, MAPKs, and NF-κB pathways in SMF-induced osteoblastic differentiation, inhibitors of Wnt (DKK1), ERK (PD98059), JNK (SP6100126), p38 (SB203580), MAPK, and NF-κB (PDTC) signaling were used. SMF-induced ALP activity, mineralization nodule formation, calcium concentration, and the mRNAs of osteoblast markers were effectively inhibited by DKK1, SP600126, SB203580, and PDTC, but not by PD98059 (Figure 7A-D).