The human lung adenocarcinoma cell lines, A549 (Cancer Research Malaysia, Subang Jaya Medical Centre, Subang Jaya, Selangor Malaysia) and SK-LU-1 (AseaCyte Pte. Ltd., Subang Jaya, Selangor, Malaysia), were maintained in RPMI-1640 (HyClone/GE Healthcare Life Sciences, Pittsburgh, PA, USA) and MEM-α (Gibco/Thermo Fisher Scientific, Waltham, MA, USA), respectively, containing 10% (v/v) fetal bovine serum (FBS) (HyClone/GE Healthcare Life Sciences), and maintained in 5.0% CO₂ levels at 37.0°C in a humidified atmosphere.

The cells were seeded 24 h prior to transfection with 80.0 nM miRIDIAN microRNA human miR-361-5p mimics or hairpin inhibitors (GE Healthcare Dharmacon, Lafayette, CO, USA) using the DharmaFECT transfection reagent (GE Healthcare Dharmacon), as per the manufacturer’s instructions. miRIDIAN microRNA Mimic Negative Control #1 and miRIDIAN microRNA Hairpin Inhibitor Negative Control #1 (GE Healthcare Dharmacon) were used as negative controls.

The cells were plated 24 h prior to transfection with 100.0 nM Stealth RNAi (siBcl-xL) using Lipofectamine 2000 reagent (Invitrogen, Carlsbad, CA, USA), as per the manufacturer’s instructions. At 24 h post-transfection, the spent media were removed and the cells were transfected with miR-361-5p mimics or mimic negative control (NC).

Apoptosis was detected using the FITC Annexin V apoptosis detection kit (BD Biosciences, Franklin Lakes, NJ, USA) at 72 h post-transfection, as per the manufacturer’s instructions. Apoptosis was detected in 1.0×10⁴ cells using the BD FACSCanto™ II flow cytometer (BD Biosciences) and analyzed using BD FACSDiva™ software (BD Biosciences).

Caspase-3 and -7 activities were determined using the Caspase-Glo 3/7 assay kit (Promega, Madison, WI, USA), at 48 h post-transfection, as per the manufacturer’s instructions. Luminescence was detected using the GloMax Multi Luminescence Multimode Reader (Promega).

Experiments involving zebrafish were approved by the University of Malaya, Faculty of Medicine, Institutional Care of Use Committee (FOM IACUC) (Ethics reference no. 2015-181006/IBS/R/NO) and complied with all relevant animal welfare laws, guidelines and policies. Wild-type Danio rerio zebrafish embryos were cared for and maintained using standard husbandry practices.

Approximately 150 zebrafish embryos (Zebrafish Laboratory, Department of Biomedical Science, Faculty of Medicine, University of Malaya, Kuala Lumpur, Malaysia) were injected with A549 cells transfected with 80.0 nm miR-361-5p inhibitors or their corresponding negative controls. The A549 cells re-suspended in serum-free RPMI were injected at the superficial location of the yolk near to the perivitelline space of the embryos using a FemtoJet Microinjector (Eppendorf, Hamburg, Germany) and InjectMan NI 2 Micromanipulator (Eppendorf) with constant injection pressure and injection time. The injection volume and cell suspension was calibrated to be approximately 100-200 cells/injection in each embryo. Following microinjection, the embryos were immediately incubated at 37°C overnight.

At 12 h post-injection, the embryos were fixed in 4% paraformaldehyde overnight at 4°C, and then dehydrated in methanol for 2 h at −20°C. Following rehydration, the embryos were washed several times with 1% DMSO, 0.1% Triton in PBS (1X PDT) and blocked for 1 h at 25°C with gentle shaking in 10% FBS, 2% BSA in PBST (blocking buffer). The embryos were then stained with purified rabbit anti-active caspase-3 antibody (Cat. no. 559565, BD Biosciences) (1:500 dilution) for 2 h at 25°C followed by washes in PDT. The embryos were again blocked with blocking buffer, and then stained with anti-rabbit IgG Fab2 Alexa Fluor 647 Conjugate (Cat. no. 4414S, Cell Signaling Technology, Danvers, MA, USA) (1:500 dilution) overnight at 4°C. The following day, the embryos were washed with PDT prior to visualization and imaging using a Leica confocal laser-scanning microscope SPII and Leica Application Suite (LAS) software v5.0 (Leica, Wetzlar, Germany). Fluorescence was quantified using ImageJ Analyst [National Institutes of Health (NIH), Bethesda, MD, USA] software. The threshold was set to eliminate background fluorescence and embryos were analyzed to generate arbitrary fluorescence units.

The putative miRNA targets were identified using TargetScan Human v5.2, a database of conserved 3′UTR targets, found at http://www.targetscan.org/ (Whitehead Institute for Biomedical Research, Cambridge, MA, USA). TargetScan allows for the ranking of predicted miRNA targets based upon a total context+ score, which is the sum of the contribution of six targeting factors which include site type, site number, site location, local AU content, 3′-supplementary pairing, target site abundance, and seed-pairing stability. The total context+ score indicates the relative repression of mRNA, with low context scores being more favorable (18). Gene-annotation enrichment analyses of the predicted miRNA targets with total context+ scores of <0 were then performed using the web tool made up of an integrated biological knowledgebase and analytic tools (19) called Database for Annotation, Visualization and Integrated Discovery (DAVID) v6.7 at http://david.abcc.ncifcrf.gov/summary.jsp (SAIC Frederick, Inc. Frederick, MD, USA) using default parameters. Data from TargetScan 5.2 and DAVID were combined to generate a hypothetical pathway of the association between miR-361-5p and its gene targets.

The cells were plated 24 h prior to co-transfection with 40.0 ng of 3′UTR reporter constructs containing wild-type or mutated bindings sites and 80.0 nM of miR-361-5p mimic/hairpin inhibitor or mimic NC/hairpin inhibitor NC using DharmaFECT transfection reagent. Relative luciferase activity was assayed at 48 h post-transfection using the Dual Luciferase Reporter Assay System (Promega) and detected on the GloMax Multi Luminescence Multimode Reader (Promega). Luciferase activity was normalized to the internal control Renilla activity in each well.

Proteins were extracted at 48 h post-transfection using the NE-PER^® Nuclear and Cytoplasmic Extraction kit (Thermo Fisher Scientific), as per the manufacturer’s instructions. Size separation by electrophoresis in 12.0% (w/v) SDS-PAGE was performed prior to transfer to nitrocellulose membranes. The membranes were blocked with 5% (w/v) non-fat skimmed milk for 1 h at 25°C and incubated overnight at 4°C with primary monoclonal rabbit antibodies: Mothers against decapentaplegic homolog 2 (SMAD2) (Cat. no. 3122, Cell Signaling Technology) (1:1,000 dilution) and GAPDH (Cat. no. 2188, Cell Signaling Technology) (1:10,000 dilution). The membranes were subsequently washed and incubated with secondary goat anti-rabbit IgG HRP-linked antibody (Cat. no. 7074, Cell Signaling Technology) (1:1,000 dilution) and anti-biotin HRP-linked antibody (Cat. no. 7075, Cell Signaling Technology) (1:1,000 dilution). Bands were visualized using Western Bright Quantum (Advansta, Menlo Park, CA, USA) on the Fusion FX7 system (Vilber Lourmat, Eberhardzell, Germany) and quantified using ImageJ Analyst software with band intensities normalized to GAPDH.

The cells were plated 24 h prior to transfection with 80.0 nM miR-361-5p mimics or mimic NC. At 6 h post-transfection, the spent media were removed and the cells transfected with 50.0 ng of plasmid expressing the SMAD2 gene without its 3′UTR (pCMV6/SMAD2) (OriGene Technologies, Rockville, MD, USA). Empty pCMV6 (OriGene Technologies) was used as a control. The protein levels of SMAD2 were determined 48 h post-transfection by western blot analysis, while apoptosis was detected using the FITC Annexin V apoptosis detection kit and Caspase-Glo 3/7 assay kit as described above.

All in vitro experiments were performed in triplicate independent experiments and presented as the means ± standard deviation (SD). In vivo experiments were performed with sample size of 15 zebrafish embryos per treatment group. An unpaired Student’s t-test was used to determine the statistical significance of the differences between 2 groups of data, where a P-value ≤0.05 was considered to indicate a statistically significant difference. The analysis of statistical significance between 3 or more groups of data was performed using one-way analysis of variance (ANOVA), followed by Dunnett’s Multiple Comparison post-hoc test where a P-value <0.05 was considered to indicate a statistically significant difference.

The silencing of Bcl-xL in the A549 cells has been shown to downregulate miR-361-5p expression (7). In this study, to investigate the biological effects of miR-361-5p, miR-361-5p hairpin inhibitor and hairpin inhibitor negative control were transiently transfected into the lung adenocarcinoma cells, A549 and SK-LU-1. The results from RT-qPCR indicated that the transfection of miR-361-5p hairpin inhibitor successfully suppressed miR-361-5p expression levels, when compared to the hairpin inhibitor negative control (Fig. 1A), whereas transfection with miR-361-5p mimics significantly increased miR-361-5p expression (data not shown). Furthermore, the inhibition of miR-361-5p expression resulted in a significant increase in caspase-3/7 activities in both the A549 and SK-LU-1 cells (1.31±0.14 and 1.42±0.07 relative activity, respectively; Fig. 1B). Correspondingly, the detection of cell death at 72 h post-transfection, using flow cytometry after Annexin V-FITC/PI staining, indicated that the inhibition of miR-361-5p resulted in a significant increase in the percentage of apoptotic A549 and SK-LU-1 cells (25.10±2.94 and 43.93±1.53%, respectively) in comparison to the hairpin inhibitor negative control (Fig. 1C). However, no significant change in apoptosis and caspase activity was observed when miR-361-5p was overexpressed using mimics (data not shown).

To elucidate the association between Bcl-xL, miR-361-5p and apoptosis, a combination study was performed whereby the cells were first transfected with siBcl-xL followed by transfection with miR-361-5p mimics. The data revealed that the relative caspase-3/7 activity in the Bcl-xL-silenced cells was significantly decreased following miR-361-5p mimic transfection (Fig. 2A); a similar decrease in the percentage of apoptotic A549 and SK-LU-1 cells was observed (Fig. 2B), thus demonstrating that the overexpression of miR-361-5p was able to block siBcl-xL-induced cell death.

To analyze the in vivo effects of miR-361-5p on apoptosis, zebrafish were used as an animal model. A549 cells transfected with either miR-361-5p inhibitor or hairpin inhibitor negative controls were microinjected into zebrafish embryos. Following staining with anti-rabbit fluorophore-conjugated antibodies, the embryos were visualized using a Leica confocal microscope (Fig. 3A). The analysis of the fluorescent images using ImageJ software indicated that th caspase-3 levels were significantly increased by 6.41±1.04-fold in the embryos injected with miR-361-5p hairpin inhibitor-transfected cells in comparison to the embryos injected with hairpin inhibitor negative control-transfected cells (Fig. 3B). These results suggested that the downregulation of miR-361-5p expression was also able to induce apoptosis in vivo.

To elucidate the underlying molecular mechanisms responsible for miR-361-5p-mediated apoptosis, an in silico approach was used to identify the putative target genes of miR-361-5p using the TargetScan database, followed by gene-annotation enrichment analyses using the web tool, DAVID, which categorizes the predicted miR-361-5p targets into apoptosis-related pathways. The enrichment of genes involved in cancer pathways indicated the possible involvement of the transforming growth factor β (TGFβ), phosphatidylinositol 3-kinase/protein kinase B (PI3K/AKT) pathway and mitogen-activated protein kinase (MAPK) pathways, together with the intrinsic and extrinsic apoptotic pathways as potential targets of miR-361-5p (Fig. 4). SMAD2, a cytoplasmic signaling protein, was identified as a predicted target of miR-361-5p, with a total context+ score of −0.12.

Among the various target genes, SMAD2 was selected for further experimental validation as it contains two seed-recognizing miR-361-5p binding sites and is known to be involved in apoptosis and proliferation (20-22). The SMAD2 3′UTR and its corresponding mutant counterpart, were cloned into the 3′ end of the pmirGLO Dual-Luciferase miRNA Target Expression Vector (Fig. 5A). Relative Firefly luciferase activity, measured in the presence of miR-361-5p mimic/inhibitors or their respective negative controls were measured. The results indicated that the cells co-transfected with wild-type constructs (WT 3′UTR) and miR-361-5p mimics exhibited a substantial 0.35-fold decrease in relative luciferase activity in comparison to the cells in the control group (Fig. 5B). In the cells co-transfected with the mutant constructs (Mut 3′UTR), no significant differences were observed between the relative luciferase activities. This observation indicates that miR-361-5p can directly bind to the binding sequence on the 3′UTR of SMAD2, suppressing its gene expression; this was further verified by a decrease in SMAD2 protein levels as determined by western blot analysis (Fig. 5C and D).

The results of this study indicated that miR-361-5p suppresses the apoptosis of A549 and SK-LU-1 cells, and inhibits SMAD2 expression. Therefore, it was predicted that the miR-361-5p-mediated inhibition of apoptosis may in part be attributed to the function of SMAD2. To confirm that the effects of miR-361-5p on apoptosis are mediated by SMAD2, gene rescue experiments were performed, whereby a SMAD2 expression vector (pCMV6/SMAD2, lacking its 3′UTR) was co-transfected into the A549 and SK-LU-1 cells along with the miR-361-5p mimic. The SMAD2 protein expression levels were measured at 48 h post transfection and it was found that transfection with pCMV6/SMAD2 ‘rescued’ or reversed the decrease in protein expression that was observed in the cells transfected only with miR-361-5p mimics (Fig. 6A and B). Furthermore, transfection with pCMV6/SMAD2 was able to partially reverse the inhibition of apoptosis induced by miR-361-5p (Fig. 6C and D). Taken together, these observations suggest that the oncogenic role of miR-361-5p in lung adenocarcinoma cells is at least partially due to the inhibition of its target gene, SMAD2.