Quinidine, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) and tetrodotoxin (TTX) were products of the Sigma-Aldrich Corp. (St. Louis, MO, USA). RPMI-1640 medium and fetal bovine serum (FBS) were obtained from Life Technologies (Carlsbad, CA, USA). Annexin V-fuorescein isothiocyanate/propidium iodide (Annexin V-FITC/PI) apoptosis detection kit was procured from Wuhan Antgene Biotechnology Co., Ltd. (Hubei, China). Caspase-3, -8 and -9 activity assay kits were obtained from Beyotime Institute of Biotechnology (Jiangsu, China). All other chemicals were of standard analytical grade.

Human glioma U87-MG cells were purchased from American Type Culture Collection (ATCC) (Manassas, VA, USA) and grown in RPMI-1640 supplemented with 10% FBS and 100 U penicillin/streptomycin in 5% CO₂ at 37°C. Cells were passaged every three days and maintained at exponential growth to ~80% confluence for later experiments.

Briefly, cells were seeded at 5,000 cells/well into a 96-well plate and incubated overnight. After exposure to different concentrations of quinidine for 48 h, MTT solution (final concentration 0.5 mg/ml) was added to each well and the samples were incubated for another 4 h. Subsequently, the supernatant was removed and cells were dissolved in 150 μl DMSO. Finally, absorbance at 570 nm was measured by using a 96-well microplate reader (Thermo Fisher Scientific, Inc., Waltham, MA, USA).

Cells were double-stained by using an Annexin V-FITC/PI apoptosis detection kit. Briefly, cells were seeded at 5×10⁶ cells per flask into cultured flasks and incubated overnight. After exposure to different concentrations of quinidine for 48 h, cells were harvested and washed with cold PBS twice, and resuspended in Annexin V binding buffer. Then 5 μl of FITC-labeled Annexin V and 5 μl of PI were added. Cells were gently oscillated and incubated for 15 min at room temperature in the dark. After addition of 200 μl binding buffer to each tube, cells were analyzed on a flow cytometer (Becton-Dickinson, Franklin Lakes, NJ, USA) within 1 h.

The activities of caspase-3, -8 and -9 were determined using the activity assay kits. To evaluate the activity of caspase, cell lysates were prepared after treatment with different concentrations of quinidine. Assays were performed on 96-well microtitre plates by incubating 10 μl protein of cell lysate per sample in 80 μl reaction buffer [1% NP-40, 20 mmol/l Tris-HCl (pH 7.5), 137 mmol/l Nad and 10% glycerol] containing 10 μl caspase substrate (Ac-DEVD-pNA for caspase-3, Ac-IETD-pNA for caspase-8, Ac-LEHD-pNA for caspase-9) (2 mmol/l). Lysates were incubated at 37°C for 4 h. Samples were measured with a microplate reader at an absorbance of 405 nm. The detail analysis procedure is described in the manufacturer’s protocol.

Membrane currents were recorded using a whole-cell voltage clamp and borosilicate glass pipettes (outer diameter, 1.5 mm; direct current resistance, 3–6 MΩ) constructed using a two-step puller (P-97; Sutter Instrument, Novato, CA, USA). To investigate the voltage-gated K⁺ currents, the pipette solution consisted of 140 mmol/l KCl, 2.5 mmol/l MgCl₂, 10 mmol/l HEPES, 11 mmol/l EGTA and 5 mmol/l ATP, and the pH was adjusted to 7.2 (18). The cells were bathed in an extracellular solution containing 135 mmol/l NaCl, 5.4 mmol/l KCl, 1.0 mmol/l MgCl₂, 0.33 mmol/l NaH₂PO₄, 1.8 mmol/l CaCl₂, 10 mmol/l HEPES and 10 mmol/l D-glucose with 1 μmol/l TTX. The osmolarity was adjusted to 330 mosmol/l with sucrose, and the pH was adjusted to 7.4. Whole-cell patch clamp recordings were performed at room temperature using a patch clamp amplifier (Axon-200B; Axon Instruments, Union City, CA, USA) (19). Adjustments of capacitance and series resistance compensation were performed before the membrane currents were recorded. The membrane currents were filtered at 10 kHz (−3 dB), and the data were processed using Clampfit (Axon Instruments).

After treated with 100 μmol/l quinidine for 48 h, cells were harvested in PBS, collected by centrifugation, and total RNA extracted using the miRNeasy kit (Qiagen, Hilden, Germany) according to manufacturer’s instructions. The experiment was repeated three times. Quantity and quality of RNA was determined by absorbance (260 and 280 nm). LC Sciences (Houston, TX, USA) performed microarray assay using 5 mg total RNA which was size fractioned using a YM-100 Micron centrifugal filter (Millipore, Billerica, MA, USA) and the small RNAs (<300 nt) isolated were 3′-extended with a poly(A) tail using poly(A) polymerase. Three biological samples of RNA were pooled and array was performed using quadruplicate internal repeats of pooled RNA as previously described (12). Data were analyzed by first subtracting the background and normalization of array was to statistical mean of all detectable transcripts. System related variation of data was corrected using LOWESS filter (locally-weighted regression) method (12). Probes were single channel and detected signals greater than background plus thre times the standard deviation was derived.

To confirm the miRNA levels obtained from the microarray results, expression of miRNAs was assessed using quantitative real-time PCR. Total cellular RNA was extracted from each of the experimental groups using Qiagen miRNeasy RNA purification system according to the manufacturer’s instructions. Quantity and quality of RNA was determined by absorbance (260 and 280 nm). Reverse transcription was performed using gene specific primers. The sequences of the primer pairs are shown in Table I. RNU6B was taken as an internal control. Quantitative real-time PCR was performed using Platinum SYBR-Green qPCR SuperMix-UDG (Invitrogen Life Technologies Carlsbad, CA, USA). The reactions were carried out in a 96-well optical plate at 95°C for 10 min and then amplified for 15 sec at 90°C, followed by 30 sec at 60°C for 40 cycles. The relative value of each miRNA was calculated using the 2^−ΔΔCt method, where Ct is the number of cycles at which the application reaches a threshold, as determined by SDS software v1.2 (Applied Biosystems, Inc., Foster City, CA, USA). Thermal denaturation was administered at the conclusion of the qPCR to determine the number of the products that were present in the reaction. Each reverse transcription and qPCR assay was performed in triplicate.

Changes in expression of miRNAs between the control and quinidine-treated groups were selected and their putative cellular target genes were predicted using TargetScan (http://www.targetscan.org/). Using gene ontology (GO) (http://geneontology.org/) and Kyoto Encyclopedia of Genes and Genomes (KEGG) analysis tool (http://www.genome.jp/kegg/), the target genes were categorized into the following two groups: apoptosis and cell proliferation.

Data were expressed as mean ± standard error (mean ± SE). Statistical significance was assessed by using paired Student’s t-test. P<0.05 was considered to be statistically significant.

To measure the effect of quinidine on the proliferation of U87-MG cells, cells were treated with various concentrations of quinidine (12.5, 25, 50, 100, 200, 400 and 800 μmol/l) for 48 h, and MTT assay was used to determine the number of the live cells remaining after the drug treatment. As shown in Fig. 1A, quinidine significantly inhibited the cell proliferation of U87-MG cells in a dose-dependent manner (P<0.01). The value of IC₅₀ was calculated to be 58.96 μmol/l at 48 h.

To determine whether the reduced cell viability by quinidine was related to apoptotic cell death, the effect of quinidine on apoptosis of U87-MG cells was studied by Annexin V-FITC/PI staining. As illustrated in Fig. 1B, quinidine increased the percentage of apoptotic cells in a concentration-dependent manner, and the apoptotic rates of U87-MG cells treated with 25, 50 and 100 μmol/l quinidine were 9.9, 29.6 and 53.1%, respectively. However, the effect of quinidine on necrosis was slight. At 100 μmol/l, quinidine only induced necrosis in ~2.5% of U87-MG cells, indicating that quinidine could induce apoptosis of U87-MG cells without causing obvious necrosis.

Apoptosis is always executed in a caspase-8-regulated plasma membrane extrinsic pathway and/or caspase-9-regulated mitochondrial intrinsic pathway (20). To determine whether caspases were activated by quinidine, we examined the activities of caspase-3, -8 and -9 in quinidine-treated U87-MG cells. As shown in Fig. 1C, quinidine treatment (25, 50 and 100 μmol/l) resulted in a dose-dependent activation of the initiator caspases-9, and the executor caspase-3. Additionally, activation of caspase-8 was also observed, but the activities of caspase-8 in quinidine-treated cells were not significantly different from that of control cells. These data implied that both mitochondrial and death receptor pathways were involved in the apoptotic process induced by quinidine in U87-MG cells, and the mitochondrial pathway was the main mode.

To determine whether quinidine inhibited cell proliferation via blockage of voltage-gated K⁺ currents (21), we next studied the dose-dependent effects of quinidine on voltage-gated K⁺ currents in U87-MG cells. The voltage-gated K⁺ currents consisted of two components (transient and sustained K⁺ currents) in U87-MG cells, and representative recordings of the voltage-gated K⁺ currents were evoked with a step-up depolarization protocol (Fig. 2). Briefly, the membrane potential was pre-hyperpolarized from −50 to −110 mV for 100 msec, depolarized from −50 to +60 mV (10 mV increment per step, duration 200 msec), and subsequently restored to the original depolarizing potential of −50 mV (Fig. 2A). Depolarizing the voltage from −110 to −50 mV and maintaining the level of −50 mV for 100 msec inactivated transient K⁺ currents, and a further depolarization voltage step from −50 to +60 mV evoked sustained K⁺ currents (Fig. 2B). The transient component (Fig. 2C) was then visualized in isolation using point-by-point subtraction of the sustained component (Fig. 2B) from the total outward current (Fig. 2A). As illustrated in Fig. 2D and E, quinidine shifted the current-voltage curves of transient and sustained K⁺ currents downward at different depolarization potentials in a dose-dependent manner. Quinidine was more potent in suppressing sustained than transient K⁺ currents. For instance, at the depolarizing voltage of +60 mV, the percentage inhibition by 100 μmol/l quinidine of transient and sustained K⁺ currents were ~46.64 and 82.04%, respectively (Fig. 2D and E).

To examine the miRNAs involved in quinidine-induced cytotoxicity, total RNA was extracted from U87-MG cells treated with 100 μmol/l quinidine for 48 h. The μParaflo^® miRNA microarray containing 2,042 mature human miRNA probes was used to identify the cellular miRNA expression profiles. We found that the expression of specifc miRNAs in quinidine-treated U87-MG cells was signifcantly altered when compared to that in untreated (control) cells. The results of quinidine-regulated miRNAs and their chromosomal locations are summarized in Table II. Of the 2,042 human miRNAs tested, 11 miRNAs were upregulated and 16 miRNAs were downregulated in our experiment.

Based on previous published reports (22,23), we selected the following five miRNAs: the upregulated miR-149-3p, the downregulated miR-25-3p, miR-100-5p, miR-424-5p and miR-365a-3p and measured their expression levels by quantitative real-time PCR. After individual miRNA level in each sample was quantified and normalized to U6 expression, quantitative PCR data confirmed that the expressions of miR-149-3p and miR-424-5p were up- and downregulated significantly in the quinidine-treated U87-MG cells, respectively (P<0.05). Although the expressions of miR-25-3p, miR-100-5p and miR-365a-3p were downregulated in microarray analysis, there were no statistically significant differences by real-time PCR analysis (Fig. 3).

Since the cellular functions of miRNAs are directly mediated by controlling their target gene expression, we further analyzed the putative target genes of the miRNAs and the functional relationship between the gene and anticancer properties using bioinformatic tools. First, the miRBase target database tool, MicroCosm, revealed that 3,803 genes were potentially targeted by quinidine-specific miRNAs. Moreover, since quinidine promotes the death of cancer cells through its anti-proliferative and pro-apoptotic activities, we selected genes with functions related to cell proliferation and apoptosis by using the GO and KEGG analysis tool. Consistent with the previous finding, the GO and KEGG analysis results showed that several target genes of the quinidine-responsive miRNAs were functionally involved in anticancer pathways. These sets of genes are shown in Fig. 4.