The UE7T-13 cells, a human bone marrow-derived MSC line infected with retroviruses expressing papillomavirus E7 and hTERT in order to prolong the life span of the cells (10–13), were purchased from the Health Science Research Resources Bank (JCRB1154; Japan Health Sciences Foundation, Tokyo, Japan) and cultured in α-modified minimum essential medium (α-MEM; Gibco-BRL, Gaithersburg, MD, USA) supplemented with 10% fetal bovine serum (FBS; Gibco-BRL). The MG-63 cells derived from human osteosarcoma (14) were purchased from the American Type Culture Collection (CRL-1427; ATCC, Manassas, VA, USA). The HuO9 human osteosarcoma cells (15) were purchased from the Japan Health Sciences Foundation (JCRB0427) and cultured in α-MEM containing 10% FBS. All cultures were maintained at 37°C in a humidified atmosphere of 5% CO₂.

The DP tissues were obtained from the crown and root of healthy human deciduous teeth (obtained from 3 donors, aged 6–8 years). Informed consent was obtained from the donors’ parents prior to tooth extraction, which was carried out at Tokushima University Hospital (Tokushima, Tokyo) during the course of orthodontic treatment. The study protocol was approved by the Ethics Committee of Iwate Medical University, School of Dentistry, Morkioka Japan (no. 01101) and Tokushima University Hospital, Tokushima, Japan.

The DP tissues were cut into sections using a surgical blade and were digested with collagenase (2 mg/ml) at 37°C for 30 min. The tissues were then washed with Dulbecco’s phosphate-buffered saline (DPBS), placed in culture dishes and maintained in α-MEM supplemented with 10% FBS. Fibroblastic cells that grew from the DP tissues were used as DPCs. When the cells reached confluence, they were detached with 0.2% trypsin and 0.02% ethylenediaminetetraacetic acid tetrasodium salt (EDTA-4Na) in DPBS (Gibco-BRL) and subcultured at a split ratio of 1:4, as previously described (1).

The DPCs were transfected with the pBABE-neo-hTERT plasmid, which harbored a neomycin-resistance gene (Addgene Inc., Cambridge, MA, USA) using Lipofectamine LTX (Invitrogen Life Technologies, Carlsbad, CA, USA) according to the manufacturer’s instructions. The cells were exposed to α-MEM containing 10% FBS and 200 µg/ml G418 (Gibco-BRL) for 12–15 days. The surviving cells were trypsinized and cultured further in 100-mm culture dishes, as previously described (16).

Single-cell clones were obtained using the limited dilution method as previously described (16). Clones obtained after single-cell cloning were named single cell clones derived from human deciduous tooth pulp cells (termed SDPCs). All the experiments were performed using clone no. 11 (SDP11 cells) cultured in α-MEM supplemented with 10% FBS in the absence or presence 10 ng/ml fibroblast growth factor (FGF)-2 (R&D Systems, Minneapolis, MN, USA) for 2 days. All cultures were maintained at 37°C in a humidified atmosphere of 5% CO₂ in air.

Total RNA was extracted from the cultured cells using TRIzol reagent (Invitrogen Life Technologies) as previously described (16). The total RNA pellet was washed briefly with 75% ethanol, resuspended in 30 µl diethylpyrocarbonate (DEPC)-treated water (Invitrogen Life Technologies) and stored at −80°C. The concentration of total RNA was determined spectrophotometrically by measuring the optical density at 260 nm, using a BioPhotometer (Eppendorf, Hamburg, Germany).

RNA (1 µg) was reverse transcribed into first-strand cDNA using a PrimeScript RT reagent kit (Takara Bio, Kyoto, Japan), according to the manufacturer’s instructions. cDNA samples were then amplified with specific primer pairs (listed in Table I) and as previously described: for SDF1 (30 cycles) (16), CXCR4 (35 cycles) (17) and glyceraldehyde 3-phosphate dehydrogenase (GAPDH, 28 cycles) (16). PCR was performed for the given number of cycles at 94°C for 1 min, 56°C for 1 min and 72°C for 1 min. Subsequently, 8 µl of the PCR product were resolved on a 4.0% agarose gel and stained with ethidium bromide. After staining, the bands were visualized using an imaging system (AE-6932GXCF; ATTO, Tokyo, Japan).

One microgram of the RNA sample was reverse transcribed into first-strand cDNA using a PrimeScript RT reagent kit (Takara Shuzo Co., Ltd., Kyoto, Japan), according to the manufacturer’s instructions and as previously described (1,16). A Thermal Cycler Dice real-time system (Takara Shuzo Co., Ltd.) was used for the two-step RT-PCR. The cDNA was amplified with SYBR Premix Ex Taq and specific oligonucleotide primers for the target sequences encoding parts of SDF1 and CXCR4. The primers (listed in Table I and as previously described) were designed based on the cDNA sequences of human mRNA for SDF1 (16), CXCR4 (17) and GAPDH (16). The amplification conditions were as follows: 10 sec at 95°C, followed by 40 cycles of 95°C for 5 sec and 60°C for 30 sec, with a final 15 sec at 95°C and 30 sec at 60°C in the Thermal Cycler Dice real-time system.

The cells were plated at a density of 1.5×10⁵ cells/dish (60 mm in diameter) and cultured for 2 days. The cells were then washed twice with DPBS and treated with lysis buffer (10 mM HEPES-KOH (pH 7.5), 100 mM KCl and 0.1% NP-40]. Conditioned medium (CM) was also collected. The protein concentrations in the cell lysates (CLs) and CM were measured using a Bio-Rad protein assay kit (Bio-Rad, Hercules, CA, USA). Equal amounts of protein (20 µg) from each sample were separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) on 10% gels and then transferred onto polyvinylidene difluoride membranes (Millipore, Bedford, MA, USA). After blocking with 5% skim milk in Tris-buffered saline containing 0.1% Tween-20 (TBST), the membranes were incubated with rabbit anti-human SDF1 antibody (Cat. no. 500-P87A; PeproTech, Rocky Hill, NJ, USA) and subsequently with anti-mouse secondary antibody (product no. 31432; Zymed Laboratories Inc., San Francisco, CA, USA). Specific protein bands on the membrane were detected using an enhanced horseradish peroxidase (HRP) conjugate substrate kit (Bio-Rad) as previously described (1).

The cells were fixed with cold 4% para-formaldehyde in 0.1 M DPBS (4) for 15 min. The cells were then rinsed in 1X Tris-buffered saline (TBS), blocked with 10% normal goat serum (NGS) in 1X Triton X-100 in TBS for 1 h, and incubated with primary antibodies overnight. The following primary antibodies were used: antibodies against SDF1 (1:1,000; PeproTech) and CXCR4 (1:1,000; Cat. no. MAB172-SP; R&D Systems). After washing in TBS, the cells were incubated for 1 h at room temperature with the appropriate secondary antibodies [anti-mouse Alexa 594 (Cat. no. R37115) at a dilution of 1:1,000 or anti-rabbit Alexa 488 (Cat. no. A-11034) at a dilution of 1:1,000 (both from Invitrogen Life Technologies)]. Finally, the nuclei were stained with 4′,6-diamidino-2-phenylindole (DAPI). The cells were then mounted on coverslips in ProLong anti-fade reagent (Invitrogen Life Technologies) and examined under an epifluorescence microscope (Nikon ECLIPSE TE2000-U; Nikon Corp., Tokyo, Japan).

Cell migration was evaluated using a Transwell system as previously described (10). Chambers with a 8-µm pore size and 6.4 mm in diameter were used (Corning Cell Culture Insert 24-well System; Sigma-Aldrich, St. Louis, MO, USA). The UE7T-13 cells were treated with DPBS or 0.1 µM AMD3100 (a specific antagonist of CXCR4; Millipore, Darmstadt, Germany) for 3 h prior to the assay. The cells were then dissociated and re-seeded into the upper chamber at a density of 1×10⁵ cells/ml. Subsequently, 700 µl of CM collected from the SDP11 cell culture were added to the lower well. Following a 6-h incubation at 37°C, the cells migrating through the membrane were fixed in 4% paraformaldehyde for 15 min, while the cells on the upper surface of the inserts were removed using cotton-tipped swabs. The Transwell chamber was immersed in 1 g/ml hematoxylin (Sigma-Aldrich) for 15 min. To quantify the migrating cells, 5 random microscopic fields per membrane were photographed using a Nikon inverted microscope system at ×40 magnification. All assays were performed independently 3 times.

In order to determine whether SDF1 expression can be modulated by FGF-2 (Cat. no. 233-FB-01M; R&D Systems), the SDP11 cells were cultured in the presence or absence of 20 ng/ml FGF-2 for 2 days. To confirm whether FGF-2 acts through the FGF receptor (FGFR), 1 µM AZD4547 (Cat. no. sc-364421, Santa Cruz Biotechnology, Inc., Dallas, TX, USA) was also used as a specific inhibitor of FGFR. Following culture, RNA was extracted and real-time PCR was performed as described above.

The results are expressed as the means ± SEM. Statistical significance was determined using one-way analysis of variance and the Bonferri correction method between pairs of groups. Differences with P-values of <0.01 were considered statistically significant.

We first confirmed the mRNA expression of SDF1 and CXCR4 in the SDP11, MG-63, HuO9 and UE7T-13 cells using RT-PCR (Fig. 1). SDF1 and CXCR4 mRNA was observed in all the cell lines examined in the present study (Fig. 1). No bands were obserbed for the water and RT(−) negative controls.

The SDP11 and UE7T-13 cells were stained using anti-SDF1 antibody (Fig. 2A–a and C–a). SDF1 presented as diffuse cytoplasmic fluorescent green signals, with no staining in the nuclei (Fig. 2A–a and C–a). No signals were observed when the primary antibody was omitted (Fig. 2A–d and C–d). CXCR4 was detected in both the cytoplasmic and cell surface compartments (Fig. 2B–a and D–a). No signal was detected when the primary antibody for CXCR4 was omitted (Fig. 2B–d and D–d).

To determine the protein expression of SDF1, whole CLs and CM were prepared from the SDP11 and UE7T-13 cells. The expression of SDF1 was detected in the CLs and CM from both cell lines using anti-SDF1 antibody (Fig. 3). The band representing SDF1 appeared at a higher molecular weight than the theoretical molecular weight, possibly due to either an association of the chemokine with other molecules or the aggregation of the chemokine, as SDF1 is able to form oligomers spontaneously (18). The highest levels of SDF1 protein in all samples tested were observed in the CLs from the SDP11 cells (Fig. 3B). SDF1 expression was significantly higher in the CLs from the SDP11 cells than in the CLs from the UE7T-13 cells (Fig. 3B); however, no significant differences (very slight difference) in SDF1 expression were observed between the CM of the SDP11 and UE7T-13 cells (Fig. 3C).

To determine whether the inhibition of CXCR4 in UE7T-13 cells inhibits cell migration, the cell cultures were treated with the CXCR4-specific inhibitor, AMD3100. As shown in Fig. 4, the migration of the UE7T-13 cells was significantly inhibited by treatment with AMD3100.

Subsquently, the effects of FGF-2 on SDF1 expression were evaluated by real-time PCR. The results of real-time PCR revealed that SDF1 expression was substantially inhibited by treatment with 20 ng/ml FGF-2 for 48 h in the SDP11 cells (Fig. 5). By contrast, treatment with AZD4547 (an inhibitor of the FGF receptor) alone significantly increased SDF1 expression, while combined treatment with FGF-2 and 1 µM AZD4547 did not significantly alter SDF1 expression (Fig. 5).