Prostate cancer tissues and para-cancer tissues (>2 cm from the surgical edge) confirmed by pathology after urologic surgery at The First Affiliated Hospital of Jinan University from September 2018 to March 2020 were selected. All 10 patients were pathologically confirmed and were not previously treated with chemoradiotherapy. Benign prostatic hyperplasia and other urinary problems were ruled out. Pathological grading was based on Gleason scoring in accordance with the World Health Organization histopathological classification standard for prostate cancer (24,25). Among the samples, 10 were from cases of prostate cancer and 10 were from adjacent tissues. All tissues were stored in a refrigerator at −80°C. The patients ranged in age from 49 to 73 years with an average age of 62.3±8.9 years. This study was performed at The First Affiliated Hospital of Jinan University and approved by the Ethics Committee of The First Affiliated Hospital of Jinan University. It was in line with the Declaration of Helsinki. All patients provided signed informed consent.

Human prostate cancer cell lines BPH-1, DU145, PC3, TRAMP-C2, and normal prostate cell line (RWPE-1) were purchased from the American Type Culture Collection (ATCC, Washington, DC, America). The cells were cultured in RPMI-1640 medium containing 10% fetal bovine serum (FBS) (Gibco; Thermo Fisher Scientific, Inc.) and incubated in an incubator (Thermo Fisher Scientific, Inc.) at 37°C with 5% CO₂. The cell culture was digested with 0.25% trypsin. After counting and digestion, cells in logarithmic growth phase were inoculated into a 6-well plate at the density of 2×10⁵ cells per well.

The cells were transfected with miR-367-3p mimics and mimic-NC (NC: Non-targeting), and were divided into the miR-367-3p mimic group (miR-367-3p upregulated group, 5′-UAGCUUAUCAGACUGAUGUUGA-3′ and 5′-AACAUCAGUCUGAUAAGCUAUU-3′) and mimic-NC control group (NC, 5′-UUCUCCGAACGUGUCACGUTT−3′ and 5′-ACGUGACACGUUCGGAGAATT−3′). Meanwhile, the cells were transfected with miR-367-3p inhibitor and inhibitor NC, which were divided into the miR-367-3p inhibitor group (the group with downregulated miR-367-3p expression, 5′-UCAACAUCAGUCUGAUAAGCUA-3′) and inhibitor NC control group (5′-GUGGAUAUUGUUGCCAUCA-3′). In addition, the cells were also transfected with si-NC, si-Rab23, vector-NC and vector-Rad23. Transient transfection of cells was performed according to the instructions for Lipofectamine 2000 (Thermo Fisher Scientific, Inc.). After transfection, cell RNA and proteins were extracted for real-time quantitative PCR (qPCR) and western blot experiments.

The prostate cancer cells were inoculated into a 12-well plate. A day later, BrdU was added at a concentration of 10 µM. The experiment was performed 24 h later. The cells were washed with PBS three times for 3 min each time and fixed with 4% paraformaldehyde for 20 min. The cells were washed with PBS three times for 3 min each time. HCl (2 M) was added at 37°C for 30 min. The cells were washed three times with 0.1 mol/l boric acid solution and then passed through a Triton X-100 permeable membrane for 20 min. The cells were blocked with 5% BSA serum at 37°C for 30 min. BrdU primary antibody was added at the dilution of 1:1,200. The cells were incubated at 4°C overnight, mixed with red fluorescent secondary antibody (1:200), and incubated for 45 min. They were stained with DAPI for 1 min. An anti-quenching agent was added to block the plate. The cells were imaged and counted under a fluorescence microscope (magnification, ×200, Nikon, Japan).

Vertical wounds were scratched into prostate cancer cells by using a 100-µl pipette tip. The cell culture medium was discarded. The wounded plate was flushed with PBS three times. After the scratch, the cells were cultured in serum-free medium. The cultured cells were photographed at 0 and 24 h. Image Pro PLUS 6.0 software (Media Cybernetics) was used to analyze and calculate the cell migration distance (26). The migration distance of each group was represented by the ratio of the migration distance and original scratch distance (9).

Cells were selected from each group 48 h after transfection. After trypsin digestion, Matrigel-Transwell chambers (Millipore, USA) were inoculated with 5×10⁴ cells per well (27). Serum-free DMEM medium was used for culture in the laboratory. A total of 500 µl of 10% FBS culture medium was added to the lower chamber. After 24 h, the upper layer without invaded cells was wiped with a cotton swab and fixed with 4% poly(methanol) (Sigma-Aldrich; Merck KGaA) for 15 min. The invasive cells were stained with 1% crystal violet for 5 min and washed with PBS for three times. Five fields were randomly selected under a Nikon Eclipse TE2000-U fluorescence microscope (magnification, ×100, Nikon) to observe and count and record the number of invasive cells.

The target gene of Rab23 for miR-367-3p was predicted by using TagetScan (http://www.targetscan.org/vert_71/) bioinformatics software. The Rab23 mRNA 3′-untranslated region (UTR) fragment containing the miR-367-3p binding site and the Rab23 3′-UTR mutation fragment mutated at the miR-367-3p binding site were cloned into pmIR-Reporter Luciferase Vector (designed by Shanghai Gemma Biological Co.). The recombinant plasmids were named Rab23-WT and Rab23-MUT. DU145 cells in the logarithmic growth phase were collected and inoculated into a 6-well plate at a density of 1×10⁶ cells per plate. After 90% cell confluence, the miR-367-3p mimic and recombinant plasmid were cotransfected with Lipofectamine™ 2000 for backup in accordance with the manufacturer's specifications (Invitrogen; Thermo Fisher Scientific, Inc.). After cotransfection for 24 h, reporter cell lysis buffer was added for 10 min at room temperature in accordance with the instructions of the dual-luciferase reporter assay kit. A total of 50 µl of firefly luciferase assay reagent was added. Relative light units (RLUs) were detected after blending. After 10 min, 100 µl of Renilla luciferase assay reagent was added. The RLU of the internal reference plasmid pRL-TK was measured after blending, and relative luciferase activity was calculated.

The mirVana miRNA separation kit and the TaqMan miRNA kit were purchased from Applied Biosystems. Reverse transcription kits (Prime Script™ RT Reagent Kit with gDNA Eraser) and real-time PCR kits (SYBR Premix II ExTaq™) were procured from TaKaRa (Japan). After digestion and counting, well-grown cells were inoculated into a 10-cm Petri dish at the density of 1×10⁶ cells per plate and incubated at 37°C and 5% CO₂ under saturated humidity. The cells were collected when their confluence reached 90%. The SYBR Green II fluorescent dye method and an IQ5™ Real-Time PCR Detection System (Bio-Rad Laboratories, Inc.) were used for data analysis. The primer sequences were as follows: U6 F, 5′-CACTGTTCCACCCCTCAGAGC−3′ and R, 5′-GCCACTTGTCGGCGATAAGG-3′ and GAPDH F, 5′-ATATCGCTGCGCTGGTCGTC-3′ and R, 5′-AGGATGGCGTGAGGGAGAsGC-3′. The reaction conditions were 94°C (15 min), 94°C (30 sec), 60°C (30 sec), and 72°C (30 sec) for a total of 40 cycles with a final extension at 72°C for 8 min. miRNA results for U6 were corrected. Rab23 mRNA expression was corrected on the basis of GAPDH expression. Relative expression was determined through the 2^−ΔΔCq method (28). Three independent replications were conducted.

The cells were collected through centrifugation and resuspended with RIPA (50 mmol/l Tris-HCl, pH 7.5, 150 mmol/l NaCl, 1% NP-40, 0.5% sodium deoxycholate, and 0.1% SDS). They were ultrasonicated at 12,000 r/min and centrifuged at 4°C for 10 min. Total protein concentration was determined in accordance with the instructions of the BCA kit. SDS-PAGE was performed with 30 µg of each sample. Protein was transferred to a PVDF membrane and blocked with 5% skimmed milk at room temperature for 1 h. The samples were incubated with the primary antibodies of Gli1 (ab217326, 1:1,000 dilution; Abcam), Gli2 (ab277800, 1:1,000, Abcam), and Rab23 (ab230200, 1:1,000, Abcam) separately. GAPDH was used as an internal reference. The samples were kept at 4°C overnight. The membranes were washed with TBST and incubated with secondary horseradish peroxidase-conjugated antibody for 1 h at room temperature. Relative protein expression after ECL development was analyzed by using QuantityOne software (v4.6.7) (Bio-Rad Laboratories, Inc.) after internal reference correction.

The tumor tissues were fixed with 40% formalin (volume fraction). The tissues (approximately 2-mm in thickness) were cut into the appropriate sizes, embedded, and microwaved with citrate buffer (pH=6.0) for antigen repair. The tissues were blocked with normal goat serum for 20 min and incubated overnight with the primary antibody (Rab23, ab230200, 1:100 dilution; Abcam) at 4°C then with the corresponding secondary antibody. The samples were then subjected to DAB color development treatment, hematoxylin redyeing, dehydration, and sealing.

A total of 12 male SPF-grade BALB/C nude mice with weights of 15–20 g/mouse and ages of 4–6 weeks were used in the experiment. Six mice in each group were inoculated with PC3-miR-NC and PC3-miR-367 prostate cancer cells under the armpit at the injection volume of approximately 0.2 ml/piece. The animal experiments were performed in Feb. 2019. The tumor volume was measured with a micron caliper when the tumor became visible (100 mm³) after inoculation. The living conditions of nude mice were observed daily. The length and width of the tumor were measured. All nude mice were sacrificed at the end of the third week of the experiment. Euthanasia method was as follows. The mice were put into a euthanasia chamber without pre-filled CO₂. Next, the cylinder was opened and 100% carbon dioxide was added. The filling rate was about 15% CO₂/min of the chamber volume. After 10 min, the nude mice were examined for death. The surviving mice continued to be treated with CO₂ for 5 mins. When the animals were determined to be not moving, breathing, and the pupils were dilated. The CO₂ was closed and another 2 min passed to confirm the animal death. The exfoliated transplanted tumor was weighed. And the maximum diameter of the tumor tissue we observed was no more than 1.3 cm. Tumor volume (TV) was calculated in accordance with the following formula: TV (mm³)=0.5 × long diameter (mm) × short diameter² (mm²). Experimental animal welfare followed the guidelines for Welfare Ethics Review (GB/T 35892-2018). Animal experiments were approved by the Ethics Committee of The First Affiliated Hospital of Jinan University (Guangzhou, Guangdong, China) (no. IRB-JN-2019-023).

Each experiment was repeated independently three times. SPSS 17.0 statistical software (SPSS Inc.) was used to analyze relevant data (29). The results are expressed as mean ± SD. Comparison between groups was performed by the unpaired Student's t-test. One-way ANOVA followed by Tukey's multiple comparison tests were selected for multiple group comparisons. Pearson correlation coefficient was used to analyze coexpression correlation. Here, P<0.05 was indicative of a statistically significant result (30).

The role of miR-367-3p in prostate cancer can be inferred by detecting its expression level. We collected 10 pairs of prostate cancer tumor tissues and adjacent control tissues. The expression of miR-367-3p was detected through qRT-PCR. The experimental results showed that miR-367-3p expression in prostate cancer tumor tissues was downregulated relative to that in the adjacent control tissues (Fig. 1A). Furthermore, the expression levels of miR-367-3p in normal human prostate epithelial cells (RWPE-1) and prostate cancer cells (TRAMP-C2, BPH-1, PC3, and DU145) were analyzed. The experimental results showed that compared with that in RWPE-1 cells, miR-367-3p expression in BPH-1, DU145, PC3, and TRAMP-C2 cells was downregulated and was the lowest in the DU145 and PC3 cells (Fig. 1B). Therefore, the PC3 and DU145 cells were selected for subsequent experiments.

After analyzing the expression level of miR-367-3p in prostate cancer tissues and cell lines, the effects of miR-367-3p on the malignant evolution of prostate cancer cells were evaluated through cell proliferation, wound healing and Transwell assays. First, we measured the expression efficiency of miR-367-3p in DU145 and PC3 cells. The experimental results showed that miR-367-3p mimics upregulated the expression of miR-367-3p, whereas the miR-367-3p inhibitor inhibited the expression of miR-367-3p (Fig. 2A and B). Subsequently, the proliferation capability of the cells was measured via the BrdU assay. The experimental results showed that the overexpression of miR-367-3p inhibited the proliferation of DU145 and PC3 cells, whereas the inhibition of miR-367-3p enhanced the proliferation of DU145 and PC3 cells (Fig. 2C and D).

We performed wound healing experiments to evaluate the migratory capability of prostate cancer cells. The results showed that the overexpression of miR-367-3p inhibited the migration of DU145 and PC3 cells, whereas the inhibition of miR-367-3p increased the migration of DU145 and PC3 cells (Fig. 3A and B). Subsequently, we performed the invasion assay to evaluate the invasive capability of prostate cancer cells. We found that the invasive capability of the DU145 and PC3 cells in the miR-367-3p mimic group was significantly lower than that in the blank control group. Compared with that of the cells in the inhibitor group, the invasion capability of DU145 and PC3 cells in the miR-367-3p inhibitor group was enhanced (Fig. 3C and D). We performed animal experiments to verify the antitumor effect of miR-367-3p. The results of the animal experiments showed that miR-367-3p inhibited tumor growth and reduced tumor weight (Fig. 3E).

We used TargetScan to predict the target genes of miR-367-3p to investigate the function of miR-367-3p. The prediction results showed that Rab23 may be a target gene of miR-367-3p. Fig. 4A shows the binding site information of Rab23 and miR-367-3p. We conducted the luciferase reporter gene experiment to further verify our conjecture in DU145 cell lines. The experimental results showed that luciferase activity in the miR-367-3p mimic + Rab23-WT (wild-type) group was lower than that in the mimic-NC + Rab23-WT group. However, after the mutation of the Rab23 binding site, the miR-367-3p mimics could no longer inhibit the luciferase activity of Rab23 (Fig. 4B). The qRT-PCR results showed that Rab23 expression in the miR-367-3p mimic group was significantly decreased compared with that in the miR-NC group. However, Rab23 expression was significantly upregulated in the miR-367-3p inhibitor group (Fig. 4C). Furthermore, we analyzed the expression level of Rab23 in prostate cancer tissues. The experimental results showed that Rab23 was upregulated in the prostate cancer tissues (Fig. 4D). In addition, we quantified the expression of Rab23 in the para-cancer control and tumor tissues. Our immunohistochemical test results showed that Rab23 was highly expressed in the prostate cancer group relative to that in the control group (Fig. 4E). miR-367-3p and Rab23 coexpression levels were highly and negatively correlated (Fig. 4F).

We studied the effect of Rab23 on the malignant evolution of prostate cancer cells through Transwell, wound healing, and cell proliferation assays. First, the expression efficiency of Rab23 in DU145 and PC3 cells was detected. The experimental results showed that siRNA Rab23 reduced the expression of Rab23 (Fig. 5A and B). Subsequently, the proliferation capability of the cells was measured via the BrdU assay. The experimental results showed that Rab23 knockdown inhibited the proliferation of DU145 and PC3 cells (Fig. 5C and D). We performed wound healing experiments to evaluate the migratory capability of prostate cancer cells. The results showed that the knockdown of Rab23 significantly inhibited the migration of DU145 and PC3 cells (Fig. 5E and F). Subsequently, we conducted the invasion assay to evaluate the invasive capability of prostate cancer cells. We found that the invasive capability of DU145 and PC3 cells in the Rab23 knockout group was significantly decreased compared with that in the blank control group (Fig. 5G and H).

Next, we examined the effects of miR-367-3p on the Hedgehog signaling pathway. RT-PCR results showed that Gli1 and Gli2 mRNAs were expressed in the prostate cancer cell lines DU145 and PC3. Expression levels were statistically analyzed in accordance with the ratio of the absorbance value of the amplified products. The expression of Gli1 mRNA in DU145 and PC3 cells was significantly decreased after transfection with miR-367-3p mimics and was statistically different than that in the normal control group (Fig. 6A and B). After transfection with miR-367-3p mimics, the expression of Gli2 mRNA in DU145 and PC3 cells was also significantly decreased (Fig. 6C and D). Changes in the expression levels of Gli1 and Gli2 were detected through western blot analysis and are shown in Fig. 6E. The experimental results showed that miR-367-3p mimics inhibited the expression of Gli1 and Gli2. However, Gli1 and Gli2 were upregulated after treatment with the miR-367-3p inhibitor. The above experimental results indicated that miR-367-3p downregulated Rab23 expression and inhibited the Hedgehog signaling pathway.

Rab23 was overexpressed, and the anticancer effect of miR-367-3p was analyzed again to validate the interaction between Rab23 and miR-367-3p. First, we detected Rab23 expression in DU145 and PC3 cells. The results of qRT-PCR and western blot analyses showed that miR-367-3p mimics could inhibit the expression of Rab23, whereas Rab23 overexpression could reverse the inhibition of miR-367-3p (Fig. 7A and B). CCK-8 assay results of DU145 and PC3 cells showed that Rab23 overexpression could promote cell proliferation, whereas miR-367-3p mimics could inhibit cell proliferation. The overexpression of Rab23 reversed the miR-367-3p mimic-mediated inhibition of proliferation (Fig. 7C and D). The invasion detection results of DU145 and PC3 cells showed that the overexpression of Rab23 could promote cell invasion, whereas miR-367-3p mimics could inhibit cell invasion. Rab23 overexpression reversed the miR-367-3p mimic-mediated inhibition and invasion of the DU145 and PC3 cells (Fig. 7E and F).