Cancer RNA-Seq Nexus (CRN) is a database of phenotype-specific transcriptome profiling in cancer cells. Using this database, we systematically obtained RNA-seq datasets concerning cervical cancer tissues from The Cancer Genome Atlas (TCGA), Sequence Read Archive (SRA) and NCBI Gene Expression Omnibus (GEO) (28). The expression data of AFAP1-AS1 in cervical cancer was downloaded from this database. The DNA methylation data of cervical cancer was downloaded from the MethHC database (http://methhc.mbc.nctu.edu.tw/php/index.php), data which came from TCGA (29). The data used to draw the survival curve were all from the UCSC XENA database (http://xena.ucsc.edu/).

The HeLa cell line (ATCC no. CCL-2) is the most widely usedcervical cancer cell line. Due to its strong ability to proliferate, invade and migrate, we selected it for further research. The HeLa cells were purchased from the American Type Culture Collection (ATCC; Manassas, VA, USA) and cultured in RPMI-1640 medium supplemented with 10% fetal bovine serum (FBS; Thermo Fisher Scientific, Inc., Waltham, MA, USA), penicillin and streptomycin (both from Sigma-Aldrich; Merck KGaA, Darmstadt, Germany) in a humidified incubator with 5% CO₂ at 37°C. For gene knockdown, the cells were seeded and cultured overnight. Subsequently, 20 nM AFAP1-AS1 siRNA or scramble control (NC) siRNA (Guangzhou RiboBio Co., Ltd., Guangzhou, China) were transfected into the cervical cancer cells using Lipofectamine 3000 (Thermo Fisher Scientific, Inc.) according to the manufacturer's instructions. The two siRNAs used in this manuscript had no off-target effect and exhibited the optimal knockdown efficiency in previous studies by us, as well as other groups (19,30). The cycling conditions of PCR were as follows: First, 10 min at 95°C. Denaturation at 95°C for 15 sec, annealing at 60°C for 30 sec, and elongation at 70°C for 10 sec. After 40 cycles, the PCR tubes were incubated 5 min at 70°C. The sequences were as follows: AFAP1-AS1 siRNA1, 5′-GGGCTTCAATTTACAAGCATT-3′ and AFAP1-AS1 siRNA2, 5′-CCTATCTGGTCAACACGTATT-3′. The abovementioned nucleotide sequences were synthesized by Guangzhou RiboBio Co., Ltd.

The cervical cancer cells were seeded and cultured in 6-well culture plates following transfection for 24 h. When the cells grew to 90% confluence, a 10 µl tip was used to create a scratch. Images were captured (magnification, ×20) at different time points (0, 24 and 48 h) using a microscope (The Cell Culture Laboratory Solution CKX53; Olympus, Tokyo, Japan). The ocular ruler was performed to measure the gap width in each group at the identified time point. All the wounds in the experimental group had the same width at the 0 h (31–33).

The invasive capacity was detected by Transwell invasion assays (34–36). The HeLa cells (1×10⁵) were transfected with AFAP1-AS1 siRNA or NC siRNA and were plated on the top chamber of a 24-well Transwell insert (8-µm pore size; BD Biosciences, San Jose, CA, USA). Matrigel (20 µl) (2 mg/ml; BD Biosciences) was placed on the top well. The bottom well contained 800 µl completed RPMI-1640 medium (containing with 15% FBS). Following 48 h of incubation at 37°C, the cells on the upper surface were removed using a cotton swab, while the cells that had invaded through the matrigel were fixed in 4% paraformaldehyde for 20 min and stained with 0.1% crystal violet (Merck KGaA, Darmstadt, Germany) for 10 min at room temperature. Invasive tumor cells were observed and calculated under a light microscope (The Cell Culture Laboratory Solution CKX53; Olympus) and 5 randomly low power (×20) fields were selected for each test.

MTT assay was used to assess cell viability (37–40). The transfected cells were seeded into 96-well plates at a density of 5×10³ cells/well. Subsequently, 100 µl complete RPMI-1640 medium were added per well and cultured for 0, 1, 2, 3, 4 and 5 days. MTT reagent (20 µl of 1 mg/ml; Sigma-Aldrich; Merck KGaA) was then added to each well followed by incubation for 4 h. The absorbance was detected at 450 nm using a spectrophotometer (Beckman Coulter, Inc., Brea, CA, USA).

The HeLa cells were harvested after being transfected with the siRNAs for 36 h and total RNA was extracted using TRIzol reagent (Invitrogen/Thermo Fisher Scientific, Inc.) according to the manufacturer's instructions. The RNA was reversed into cDNA using the 5X All-In-One RT Master Mix Reagent kit (Abm Canada Inc., Milton, ON, Canada) according to the kit protocol. cDNA was subjected to RT-qPCR using the SYBR Premix Ex Taq II kit (Takara Bio, Inc., Shiga, Japan). Reactions were carried out on the CFX96 Real-Time PCR Detection System (185–5196; Bio-Rad Laboratories, Inc., Hercules, CA, USA). Fold changes of lncRNA were calculated using the 2^−ΔΔCq method, with β-actin as an internal control (41). The sequences of the primers were as follows: AFAP1-AS1 forward, 5′-AATGGTGGTAGGAGGGAGGA-3′ and reverse, 5′-CACACAGGGGAATGAAGAGG-3′; β-actin forward, 5′-TCACCAACTGGGACGACATG-3′ and reverse, 5′-GTCACCGGAGTCCATCACGAT-3′.

Cell cycle distribution was detected by flow cytometry. When the cells were transfected with the siRNAs for 48 h, the cells were collected and fixed in 75% ethanol at −20°C overnight. The fixed cells were then washed thrice with phosphate-buffered saline (PBS) before being incubated at room temperature with RNase A for 20 min. These cells were stained using a propidium iodide (PI) staining kit (BD Biosciences) and incubated in the dark for 30 min at 4°C. A Beckman flow cytometer (Beckman Coulter, Inc.) was used for cell cycle analysis. Three independent experiments were conducted.

Total protein in the HeLa cells was extracted using RIPA extraction reagent (Beyotime Institute of Biotechnology, Shanghai, China) supplemented with a protease inhibitor cocktail (Roche Diagnostics, Basel, Switzerland). Protein from the HeLa cells was quantified by the BCA method. To separate the protein, 30–50 µg protein per lane was loaded and electrophoresed in 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE). Following electrophoresis, the protein bands were transferred onto PVDF membranes. The membranes were blocked with 5% skim milk for 1 h at room temperature and then incubated with primary anti-Rho GDP-dissociation inhibitor 1 (RHOGDI, 1:1,000; cat. no. 14282-1-AP), anti-Rac Family Small GTPase 2 (RAC2, 1:1,000; cat. no. 10735-1-AP), anti-RAB1B (1:800; cat. no. 17824-1-AP), anti-RAB11A (1:800; cat. no. 20229-1-AP), anti-profilin 1 (PFN1, 1:1,000; cat. no. 11680-1-AP) and anti-LIM and SH3 protein 1 (LASP1, 1:1,000; cat. no. 10515-1-AP) antibodies (ProteinTech, Inc., Rosemont, IL, USA) or E-cadherin (1:1,000; cat. no. 3195), Zonula occludens-1 (ZO-1, 1:1,000; cat. no. 8193), Vimentin (1:1,000; cat. no. 5741), β-catenin antibodies (1:1,000; cat. no. 8480) (Cell Signaling Technology, Inc., Danvers, MA, USA) overnight at 4°C. The following day, the membranes were incubated with a horseradish peroxidase-conjugated secondary antibody (goat anti-mouse IgG-HRP; cat. no. SC-2005; and goat anti-rabbit IgG-HRP, cat. no. SC-2005; 1:2,000; Santa Cruz Biotechnology, Inc., Santa Cruz, CA, USA) for 30 min at 37°C. Immunoreactive bands were visualized using an ECL detection reagent (Amersham; GE Healthcare Life Sciences, Chalfont, UK). β-actin (1:1,000; cat. no. 4970; Cell Signaling Technology, Inc.) expression was used as a housekeeping gene control.

All the experimental data are represented as the means ± standard error (SE) and processed using SPSS statistical software, version 19.0 (SPSS, Inc., Chicago, IL, USA). Overall survival (OS) was analyzed using the Kaplan-Meier method and the results of the analysis were considered significant if log-rank test yielded a value of P<0.05. Significance between 2 groups was evaluated by a Student's t-test. Pearson's correlation analysis was used to assess the correlation between AFAP1-AS1 expression and its methylation. For the comparison of multiple groups, analysis of variance (ANOVA) with Dunnett's post hoc t-test was performed. A P-value <0.05 was considered to indicate a statistically significant difference.

Firstly, the expression level of AFAP1-AS1 in cervical cancer samples was analyzed using TCGA data from the CRN database. The data suggested that AFAP1-AS1 was more abundant in the cervical cancer tissues compared to the normal tissues (P<0.001; Fig. 1A). Moreover, a high AFAP1-AS1 expression was positively associated with the TNM stage (P<0.05; Fig. 1B). To validate whether the expression levels of AFAP1-AS1 were associated with the OS of patients with cervical cancer, we analyzed the patient outcome data from the UCSC XENA database. This analysis revealed that compared to patients with a low expression of AFAP1-AS1, patients with a high AFAP1-AS1 expression had a shorter survival time (P<0.05; Fig. 1C).

DNA methylation is an important part of epigenetic inheritance and is involved in the transcriptional expression of lncRNAs (42,43). In the present study, to investigate whether the expression of AFAP1-AS1 is influenced by DNA methylation, we downloaded the methylation data of AFAP1-AS1 in cervical cancer from the human pan-cancer method database, MethCH. We found that the average methylation level of the AFAP1-AS1 promoter region was lower in the cervical cancer samples when compared with the non-tumor cervical cancer samples (P<0.01; Fig. 2A). In addition, the results of Pearson's correlation analysis revealed that the expression level of AFAP1-AS1 negatively correlated with its methylation level (r=−0.33, P<0.001; Fig. 2B). The survival analysis was constructed using the UCSC XENA database, and we identified that the hypomethylation of AFAP1-AS1 was associated with a poor overall survival of patients with cervical cancer (P=0.05; Fig. 2C). These data suggested that the high expression of AFAP1-AS1 may be related to the hypomethylation of its promoter region. In addition, the hypomethylation of the AFAP1-AS1 promoter region and the high expression levels of AFAP1-AS1 may act as independent prognostic indicators of patients with cervical cancer.

To elucidate the role of AFAP1-AS1 in cervical cancer, the AFAP1-AS1 expression was downregulated by transfection of the cervical cancer cell line, HeLa, with two validated siRNAs targeting AFAP1-AS1 (siRNA1 and siRNA2). We examined the knockdown efficiency of these two siRNAs by RT-qPCR which yielded satisfactory results. AFAP1-AS1 expression was knocked down by at least 65% in the HeLa cells (both siRNAs, P<0.01; Fig. 3A). The functional experiments were performed to examine the phenotypic alterations induced by the silencing of AFAP1-AS1 in the cervical cancer cells. The scratch test results revealed that the migratory capacity of the AFAP1-AS1-silenced cells was suppressed at 24 and 48 h following transfection with siRNA compared to the si-control group (24 h, siRNA1 vs. control, P<0.01; siRNA2 vs. control, P<0.05; 48 h, siRNA1 vs. control, P<0.05; siRNA2 vs. control, P<0.001; Fig. 3B and C). Moreover, as shown in Fig. 4, the number of invaded cells was also decreased in the AFAP1-AS1 siRNA-transfected cancer cells compared with the si-control group (both siRNAs, P<0.001; Fig. 4). However, we noted that there was no significant association between the expression of AFAP1-AS1 and cell viability and cell apoptosis. By contrast, AFAP1-AS1 knockdown slightly, yet significantly increased the number of cells in the G0/G1 phase in the present study (P<0.05; Fig. 5).

To investigate the potential mechanisms of AFAP1-AS1 regarding its promoting effect on the migration and invasion of cervical cancer cells, we examined the expression levels of several key molecules of the Rho/Rac signaling pathways by western blot analysis. The results suggested that the knockdown of AFAP1-AS1 increased the expression of RHOGDI, PFN1, RAB11A and RAC2, while it decreased the expression of RAB1B and LASP1 (Fig. 6).

In order to further investigate the mechanisms underlying the role of AFAP1-AS1 in cervical cancer development, we also detected changes in the protein levels of some EMT-related genes. The results revealed that following the knockdown of AFAP1-AS1, the expression of ZO-1 increased, while the expression of Vimentin and β-catenin decreased. However, E-cadherin expression was not markedly altered (Fig. 7).