The LN229 human glioblastoma cell line was purchased from the American Type Culture Collection (ATCC, USA). Cells were plated into T25 flasks and maintained in Dulbecco's Modified Eagle's Medium (DMEM) supplemented with 10% fetal bovine serum (FBS), 100 U/mL penicillin and 100 μg/mL streptomycin (all from Gibco, USA). Cells were incubated at 37°C in an incubator (SANYO, Japan) with humidified 5% CO₂/95% air. Cells were passaged every 3–5 days, and cryopreserved with liquid nitrogen tank when necessary. The drug VU0155041 sodium was obtained from Tocris (UK) and dissolved in sterile 0.9% normal saline (NS). The recombinant human sonic Hedgehog protein (shh) was purchased from R&D Systems (USA) and reconstituted in sterile 0.01 M phosphate buffered saline (PBS) containing 0.1% bovine serum albumin (BSA).

LN229 cells were plated onto coverslips and fixed with 4% paraformaldehyde (PFA) at room temperature (RT) for 20 min followed by washing three times in PBS. Before blocking with BSA (5% in PBS) for 1 h, the cells were permeabilized in 0.1% Triton X-100 for 10 min. The primary rabbit anti-mGluR4 polyclonal antibody (1:200, Abcam, UK) was added and incubated overnight at 4°C. The cells were then incubated with Alexa Fluor 594 conjugated anti-rabbit IgG (1:800, Invitrogen, USA) at RT for 2 h. Cell nuclei were counterstained using 4′,6-diamedino-2-phenylindole (DAPI, 1 μg/mL). Coverslips were mounted on slides by antifade mounting medium (Molecular probes, USA). The immunostaining negative control was generated by replacing the primary antibody with 5% BSA, and neural stem cells, which show functional expression of mGluR4 (Saxe et al., 2007; Zhang et al., 2016), were used as a positive control. Images were acquired using a fluorescence microscope equipped with a digital camera (BX51 + DP71, Olympus, Japan) and analyzed with ImageJ software (NIH, USA).

All siRNA duplexes were designed and synthesized by Genepharma Corporation (China). The siRNA sequences targeting Gli-1 and mGluR4 were as follows: siGli-1-1, 5′-GGCUCAGCUUGUGUGUAAUTT-3′; siGli-1-2, 5′-CUCCACAGGCAUACAGGAUTT-3′; simGluR4-1, 5′-GCAUGUCACCAUAAUUUGCTT-3′; simGluR4-2, 5′-GGUCAUCGGCUCAUGGACATT-3′; A non-specific siRNA (siNC, 5′-CGTACGCGGAATACTTCGATT-3′) was used as a negative control for the siRNA transfection experiments. siRNA transfection was performed as previously described with minor modifications (Zhang et al., 2015). In brief, LN229 cells were cultured in 24-well plates. Before siRNA transfection, cells were rinsed three times using pre-warmed Opti-MEM medium (Invitrogen). The cells were incubated with 100 nM of siRNA duplexes in the presence of Lipofectamine 2000 (Invitrogen, USA) and Opti-MEM for 6 h at 37°C. Then, the medium was replaced, and the cells were cultured for 24 h before further processing. siRNA transfection efficiency was evaluated by a fluorescence-labeled non-specific siRNA. Depletion of mGluR4 and Gli-1 expression was confirmed by western blot analysis at 24 h post-transfection.

Cell viability was assessed using a 3-(4, 5-dimethylthiazol-2-yl)- 2,5-diphenyl tetrazolium bromide (MTT)-based assay. LN229 cells were cultured in 96-well plates for 24 h prior to the experiments. At the indicated time points, 20 μL of MTT (5 mg/mL, Sigma-Aldrich, USA) was added to each well and incubated for 2 h at 37°C. Then, the medium was aspirated and the resulting insoluble purple crystals were dissolved using 150 μL dimethyl sulfoxide (DMSO, Sigma-Aldrich). Absorbance at 490 nm was read using a multi-microplate spectrophotometer (BioTek, USA). Triplicate wells were subjected to each treatment and at least three independent experiments were performed. The assay data were presented as the percentage of A490 of each treatment to that of the control cells.

Cell cycle and apoptosis were analyzed by FACS assay. LN229 cells were plated into T25 flasks and subjected to treatments. At the predetermined time points, the adherent cells were collected by trypsin digestion. Cell cycle was measured as previously described (Chen et al., 2010). Briefly, cells were fixed in ice-cold 75% ethanol overnight at 4°C. After three washes in PBS, the cells were incubated with 100 μg/mL propidium iodide (PI) containing 100 μg/mL RNase A for 30 min at 37°C in the dark. Then, FACS assay was performed using a FACS Calibur (BD Biosciences, USA) with excitation at λ488 nm and emission at λ630 nm. For each sample, 20 000 cells were detected. Data were collected and analyzed using FACSort CellQuest software and Modfit LT software (both from BD Biosciences), respectively. The changes in cell cycle distribution were demonstrated by the proliferation index. The following formula was used: proliferation index = [(S+G2/M)/(G0/G1+S+G2/M)] × 100%. Apoptosis analysis was conducted with Annexin V-FITC and PI staining using a Cell Apoptosis Kit (Invitrogen) according to the manufacturer's instructions. In brief, cells were suspended in 100 μL PBS, mixed with 5 μL of Annexin V-FITC and 1 μL of PI (100 μg/mL) and incubated at RT for 20 min. Then 400 μL of 1x Annexin-binding buffer was added, and the cells were immediately placed on ice. Cell apoptosis was analyzed with a BD FACS Calibur. The percentage of early apoptotic cells (Annexin V⁺/PI⁻, lower right quadrant) plus later apoptotic cells (Annexin V⁺/PI⁺, upper right quadrant) was calculated. At least three independent experiments were performed in each assay.

Cell apoptosis was detected using a terminal-deoxynucleotidyl transferase-mediated dUTP nick end labeling (TUNEL) assay kit (Roche Diagnostics, USA) according to the manufacturer's instructions. In brief, LN229 cells were plated onto coverslips. After treatments, the cells were fixed with 4% PFA for 20 min, followed by permeabilization using 0.1% Triton X-100 in 0.1% sodium citrate buffer for 2 min on ice. The cells were further stained with 50 μL TUNEL reaction mixture for 1 h at 37°C and extensively rinsed. Counterstaining was performed with DAPI (1 μg/mL) and coverslips were mounted onto the glass slides. Images were collected using a fluorescence microscope with a 40 × objective (BX51 + DP71, Olympus) and processed with ImageJ software. For cell death analysis, TUNEL stained cells in 10 randomly selected fields of each sample were counted, and triplicate samples were used for each treated group. At least three independent experiments were carried out. The percentage of TUNEL-stained cells among the total cells (DAPI-stained cells) was determined to evaluate cell death.

LN229 cells were cultured on 6-well plates and treated. Cells were harvested with RIPA lysis buffer (Pierce, USA) containing anti-protease cocktail (Roche, USA) and cleared by centrifugation at 4°C. Protein concentrations were measured by BCA reagents (Pierce). Twenty to forty micrograms of each sample was subjected to 8 or 12% SDS-PAGE and transferred to polyvinylidene fluoride membranes (Bio-Rad, USA). Membranes were blocked with 5% non-fat milk for 1 h, followed by incubation with specific primary antibodies. The primary antibodies and dilutions used in the experiments were as follows: rabbit anti-mGluR4 (1:1,000, Abcam); rabbit anti-Gli-1 polyclonal (1:1,000, Cell Signaling Technology); rabbit anti-caspase 3 polyclonal (1:1,000, Cell Signaling Technology); rabbit anti-caspase 8 polyclonal (1:1,000, Cell Signaling Technology); rabbit anti-caspase 9 polyclonal (1:1,000, Cell Signaling Technology); mouse anti-Bcl-2 monoclonal (1:1,000, Millipore); mouse anti-Bax monoclonal (1:1,000, Millipore); mouse anti-cyclin D1 monoclonal (1:1,000, Cell Signaling Technology); mouse anti-β-actin monoclonal (1:10,000, Sigma-Aldrich). Then, membranes were incubated with horseradish peroxidase-conjugated anti-rabbit or anti-mouse secondary antibodies (1:100,000, Sigma-Aldrich) for 1 h, and processed for visualization with enhanced chemiluminescent substrate (Pierce) followed by development on X-ray films (Fuji, Japan). Immunoblot bands were imaged by a gel imaging system (G: Box, Syngene, UK) and analysis of band intensity was performed with ImageJ software. The expression levels of investigated proteins were determined and normalized to the internal control, β-actin. All western blot data represent at least three separate experiments.

Data are shown as the mean ± SD of at least three independent experiments (n = 3–6, which always refers to independent experiments). Each experiment was run in triplicate or quadruplicate. Statistical comparisons were carried out by one-way ANOVA followed by Tukey's test with SPSS software (Version 23.0). P < 0.05 was considered as the standard for statistical significance.

Expression of mGluR4 in LN229 cells was determined by a specific primary antibody using immunofluorescence staining. The results showed that 95 ± 5% of the LN229 cells expressed mGluR4 (Figure 1A, Figure S1). To identify the effect of mGluR4 activation on cell viability, LN229 cells were treated with serial concentrations of a specific mGluR4 agonist, VU (1, 10, 30, and 50 μM) for 12, 24, 48, and 72 h. MTT assay showed that VU treatments decreased viability of LN229 cells in a time- and dose-dependent manner. Treatments with 30 or 50 μM of VU induced significant reduction of cell viability at 24, 48, and 72 h, compared that of controls (Figure 1B). Because there was no significant difference in cell viability between 30 and 50 μM VU treatments, the lower dose of 30 μM VU was selected for further experiments.

To observe the effect of mGluR4 on proliferation of LN229 cells, mGluR4 gene expression was downregulated using a small interfering RNA technique. Transfection efficiency was determined using a fluorescence-labeled non-specific control siRNA. Western blot analysis revealed that mGluR4 protein expression in LN229 cells was effectively reduced by transfection with gene-targeted siRNAs (simGluR4-1 and simGluR4-2), compared with that following siNC transfection, while transfection with Lipofectamine 2000 only (vehicle) and siNC had no obvious influence on mGluR4 expression, compared with that of non-transfected cells (Figures 2A,B). High expression levels of mGluR4 were found in cerebellar tissue, which was used as a positive control (Figures 2A,B).

Utilizing the mGluR4-downregulated LN229 cells, we examined the influence of VU on the expression of cyclin D1, one of the key regulators of the cell cycle. In the siNC transfected LN229 cells, cyclin D1 was significantly decreased after treatment with 30 μM of VU for 24 h, while VU had no significant influence on cyclin D1 expression in the LN229 cells transfected with mGluR4-targeted siRNAs (Figures 2C,D). Furthermore, transfection with mGluR4-targeted siRNAs rescued the reduction in cyclin D1 induced by VU, compared with that of the siNC-transfected group (Figures 2C,D). These data suggested that mGluR4 activation inhibit the expression of Cyclin D1, which may result in cell cycle arrest and decrease of LN229 cell proliferation.

We further determined the effect of mGluR4 activation on LN229 cell apoptosis. Expression levels of pro-apoptotic precursors, pro-caspase-8/9/3, and the anti-apoptotic/apoptotic protein pair Bcl-2/Bax were investigated in LN229 cells by western blot analysis. Participation of these potential intracellular moderators in cellular apoptosis was previously estimated by other investigators by determining their intracellular levels (Elmore, 2007; Ouyang et al., 2012; Zhang et al., 2015). Expression of mGluR4 in LN229 cells was depleted following transfection with mGluR4-specific siRNAs (simGluR4-1 and simGluR4-2), and treatment with the vehicle or 30 μM of VU for 24 h. In siNC-transfected cells, VU treatment led to a significant decrease in pro-caspase-8/9/3 levels. However, in the LN229 cells transfected with mGluR4-specific siRNAs, the levels of these proteins were not markedly influenced by VU treatment. Compared with that of the siNC-transfected cells, the inhibitory action of VU on pro-caspase-8/9/3 expression was significantly attenuated by mGluR4 knockdown (Figures 3A,C–E, Figure S2).

We also assessed the effect of the mGluR4 agonist on expression of the anti-apoptotic/apoptotic protein pair Bcl-2/Bax. Western blot analysis revealed that VU treatment decreased the level of anti-apoptotic protein Bcl-2 and increased the level of apoptotic protein Bax. Hence, the ratio of Bcl-2/Bax was significantly reduced, as compared with that in the control. However, the effect of VU on Bcl-2/Bax expression was markedly prevented by mGluR4-specific siRNA transfection (Figures 3B,F–H).

LN229 cell apoptosis was further analyzed using Annexin V-FITC and PI staining followed by FACS assay. In siNC transfected cells, agonist VU significantly induced apoptotic cell death, compared with that of the control. However, after transfection with mGluR4-specific siRNAs, the cell apoptosis induced by VU was abated, as compared with that of untreated counterparts. By comparison with that of the siNC-transfected cells, the enhancement of cell apoptosis by VU was significantly attenuated by mGluR4 downregulation (Figures 4A,B). These results suggested that the effect of mGluR4 activation on apoptosis-related proteins tended to induce the apoptosis of LN229 cells.

To explore the potential link between mGluR4 and regulation of Gli-1 level in LN229 cells, the cells were treated with 30 μM of VU for serial durations (0.5, 1, 4, 12, and 24 h) or with serial concentrations of VU (1, 3, 10, 30, and 50 μM) for 24 h. Then, the expression levels of Gli-1 protein were determined using western blot analysis. As shown in Figure 5, expression of Gli-1 was significantly downregulated by mGluR4 activation, and the agonist VU functioned in a time-dependent manner (Figures 5A,C). Moreover, Gli-1 expression was inhibited by VU in a dose-dependent manner; 10, 30, and 50 μM of VU significantly suppressed Gli-1 expression, while 1 and 3 μM did not (Figures 5B,D).

We further confirmed the inhibitory role of mGluR4 in Gli-1 expression in LN229 cells transfected with mGluR4-targeted siRNA. In the siNC-transfected LN229 cells, Gli-1 expression was decreased by VU treatment, while VU had no significant influence on Gli-1 level in the cells transfected with mGluR4-targeted siRNA. Compared with that of siNC-transfected cells, mGluR4 downregulation significantly attenuated the contribution of VU to repression of Gli-1 expression in LN229 cells (Figures 5E,F). These data suggested that activation of mGluR4 inhibits the expression of Gli-1 and thereby may block the activation of the SHH pathway.

Involvement of Gli-1 in regulation of LN229 cell growth was further investigated. Expression of Gli-1 was knocked down using a small interfering RNA technique. Western blot analysis showed that Gli-1 expression in LN229 cells was significantly blocked by transfection with Gli-1-targeted siRNA (siGli-1-1 and siGli-1-2), compared with that following siNC transfection. A high expression level of Gli-1 was found in a HepG2 cell line, which was used as a positive control in the experiments (Figures 6A,B). MTT assay demonstrated that Gli-1 downregulation significantly reduced LN229 cell viability. However, in the presence of the shh protein (shh, 5 μg/mL), one of the Hedgehog ligands that can increase the intracellular level of Gli-1, the viability of LN229 cells was remarkably increased (Figure 6C). Furthermore, the expression of cyclin D1 was suppressed in the Gli-1-targeted siRNA-transfected cells compared with that in siNC-transfected cells. Treatment with shh protein increased cyclin D1 expression (Figures 6D,E).

To examine the effect of Gli-1 downregulation on cell apoptosis, expression levels of the aforementioned apoptosis-related proteins were estimated in Gli-1-downregulated LN229 cells. The results revealed that the expression of pro-caspase-8/9/3 was reduced in Gli-1-targeted siRNA-transfected cells. Gli-1 knockdown also led to decrease of Bcl-2 level and increase of Bax level, which caused a decline in the Bcl-2/Bax ratio. Conversely, shh protein administration significantly increased pro-caspase-8/9/3 expression and the Bcl-2/Bax expression ratio (Figure 7). These findings were consistent with a recent investigation of medulloblastoma cells (Lin et al., 2016) and, implied an important role of Gli-1 in regulation of apoptosis and cell cycle progression in LN229 cells.

To further identify whether the reduction in Gli-1 expression is involved in the growth inhibition and apoptosis of LN229 cells induced by mGluR4 activation, the cells were treated with 5 μg/mL of shh protein, which subsequently increased the expression level of Gli-1 and growth of the tumor cells. Then, 30 μM of VU was used to eliminate the effect of shh on Gli-1 expression and growth of LN229 cells. Cell proliferation and apoptosis were then assessed. Cell cycle analysis revealed that the percentage of proliferating cells (S + G₂/M) was significantly increased by shh treatment as compared with that of the control, while VU treatment remarkably abolished the effect of shh on cell proliferation. VU in the absence of shh also played an inhibitory role in cell proliferation (Figure 8A). Similarly, the effect of shh on increasing expression of cyclin D1 was also attenuated by VU treatment (Figures 8B,C).

Cell apoptosis was assessed using TUNEL staining. As shown in Figure 9, shh treatment significantly decreased the number of TUNEL-positive cells, and VU treatment eliminated the inhibitory effect of shh on LN229 cell apoptosis, which enhanced cell apoptosis, compared with that of shh-treated cells. Consistently, VU treatment without shh remarkably increased apoptosis of LN229 cells. Correspondingly, treatment with shh substantially suppressed cleaved activation of pro-caspase-8/9/3, decreased the Bax level, and increased the Bcl-2 level and Bcl-2/Bax ratio, which imply a decrease of cell apoptosis. However, the regulatory role of shh in the expression of these apoptosis-related proteins was antagonized by VU administration (Figure 10), which suggests the potential involvement of Gli-1 downregulation in mGluR4-mediated inhibition of proliferation and enhancement of apoptosis of LN229 GBM cells.

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest. The reviewer KKS and handling Editor declared their shared affiliation.