HepG2, HepH3B, SK-Hep-1 and LO2 cell lines were obtained from the Cell Bank of The Chinese Academy of Sciences (Shanghai, China). [d-Ala2, d-Leu5] enkephalin (DADLE), naltrindole, GF109203X, U0126, and MTT were purchased from Sigma-Aldrich (St. Louis, MO, USA). The 5′,6,6′-tetrachloro-1,1′,3,3′-tetraethylbenzimidazolylcarbocyanine iodide (JC-1) Mitochondrial Membrane-Potential Assay kit was purchased from Abcam Plc (Cambridge, MA, USA). RPMI-1640 and fetal bovine serum (FBS) were purchased from Gibco-BRL (Carlsbad, CA, USA). An Annexin V-fluorescein isothiocyanate (FITC) apoptosis kit was purchased from Bio-Rad (Hercules, CA, USA). Phosphorylated PKC (rabbit, monoclonal), Bcl-2 (rabbit, polyclonal), and Bcl-2-associated X (Bax) antibodies (rabbit, polyclonal) were purchased from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA, USA). The following antibodies were used for western blot analysis: β-actin (sc-47778, diluted 1:1,000) and were obtained from Santa Cruz Biotechnology, Inc. Cytochrome c and phosphorylated ERK antibodies were purchased from Abcam Plc. The present study was approved by the Ethics Committee of The Second Affiliated Hospital of Dalian Medical University (Dalian, Liaoning, China).

Liver cancer cells were seeded at a density of 1×10⁶ cells/ml in T25 cell-culture flasks and cultured in Dulbecco’s modified Eagle medium (Gibco-BRL) supplemented with 10% FBS, penicillin and streptomycin in an incubator with 95% O₂ and 5% CO₂.

Liver cancer cells were cultured for 12 h. Except for those in the control group, cells were treated with various concentrations of H₂O₂ (10, 50, 100, 200 and 400 mM) for 12 h. While under H₂O₂ treatment, cells in the intervention groups were also treated with either the δ-opioid-receptor agonist DADLE (0.01, 0.1, 1.0 or 10 μM), the δ-opioid-receptor-specific inhibitor naltrindole, the PKC inhibitor GF109203X (10 μM) or the ERK inhibitor U0126 (10 μM) for 12 h.

The MTT assay was used to analyze cell viability. Human liver cancer cells were treated with 200 mM H₂O₂ and various concentrations of DADLE for 12 h followed by incubation with 20 μl MTT solution [5 mg/ml in phosphate-buffered saline (PBS), pH 7.4] for 4 h. The culture media was then removed and the formazan crystals in each well were fully dissolved in 200 μl dimethyl sulfoxide by vortexing for 10 min. The absorbance value of each well was measured and recorded using a microplate reader (FLx800™; Bio-Tek Instruments, Inc., Winooski, VT, USA) at a wavelength of 570 nm.

An Annexin V-FITC apoptosis kit was used according to the manufacturer’s instructions. Cell death was detected using flow cytometry (FACS Vantage SE flow cytometer: BD Biosciences, Franklin Lakes, NJ, USA). Cells positive for Annexin V and negative for PI were considered to be early apoptotic cells. In brief, cells were harvested using 0.25% trypsin, washed with PBS three times, stained with 10 μl Annexin V and 5 μl PI and incubated in the dark at room temperature for 15 min. Cells were then analyzed using flow cytometry.

Cells were suspended at a concentration of 1×10⁵ cells/ml, incubated with 10 μg/ml JC-1 staining solution, mixed thoroughly and incubated for 20 min. Non-conjugated JC-1 was removed using buffer, then cells were resuspended in buffer. Cells were analyzed using flow cytometry with an emission wavelength of 488 nm. The FL1-h and FL2-h values represent the intensities of red and green fluorescence, respectively. The results were quantitatively analyzed using CellQuest software (BD Biosciences, Franklin Lakes, NJ, USA).

In each group, liver cancer cells were collected and suspended in pre-chilled extraction buffer [0.2 mol/l mannitol, 50 mmol/l sucrose, 1 mmol/l EDTA, 1 mmol/l ethylene glycol tetraacetic acid, 10 mmol/l 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid, pH 7.4, 50 mmol/l dithiothreitol, 5 mmol/l protease inhibitor cocktail and 1 mmol/l phenylmethylsulfonyl fluoride (PMSF)]. Following homogenization for 5–6 repetitions, cell homogenates were centrifuged at 1,000 × g for 10 min at 4°C and the supernatant was collected. The supernatant was then centrifuged at 7,000 × g for 10 min at 4°C. The pellet was collected and the supernatant was centrifuged at 15,000 × g for 10 min at 4°C. The pellet was collected and exposed to density gradient centrifugation using Nycodenz^® (Axis-Shield, Oslo, Norway). The isolated mitochondria were then stored in a buffer solution at −80°C until required.

Cells were lysed in pre-chilled lysis buffer (50 mM Tris-HCl, 137 mM NaCl, 10% glycerol, 100 mM sodium orthovanadate, 1 mM PMSF (Gibco-BRL), 10 mg/ml aprotinin (Santa Cruz Biotechnology, Inc.), 10 mg/ml leupeptin (Amresco, Solon, OH, USA) and 1% Nonidet P-40 (Sigma; pH 7.4). Lysates were centrifuged at 12,000 × g for 20 min, after which the supernatant was collected and mixed with loading buffer (65 mM Tris-HCl, pH 6.8, 3% SDS, 10% glycerol and 6 M urea; Millipore, Billerica, MA, USA). Total protein concentrations were measured using a Pierce™ BCA protein assay kit (Pierce Chemical Co., Rockford, IL, USA). The electrophoresis buffer was supplemented with β-mercaptoethanol (Solarbio, Beijing, China) and bromophenol blue (Thermo Fisher Scientific, Rockford, IL, USA). Proteins were separated using 12% SDS-PAGE, transferred onto a polyvinylidene fluoride membrane (Bio-Rad), and incubated with primary antibodies at 4°C overnight. Protein bands were detected using an enhanced chemiluminescence method and the intensity of the protein bands were analyzed using a gel image-analysis system (Molecular Imager Gel DocTM XR+ system; Bio-Rad).

Total RNA was extracted from the cells in each group using an RNAiso™ Plus kit (Takara Bio, Inc., Shiga, Japan) according to the manufacturer’s instructions. Total RNA was subsequently quantified. The upstream primer for the δ-opioid receptor was 5′-ACCAAGATCTGCGTGTTCCT-3′, and the downstream primer was 5′-CGATGACGAAGATGTGGATG-3′. The upstream primer for the internal control, β-actin, was 5′-AAGGAAGGCTGGAAGAGTGC-3′, and the downstream primer was 5′-CTGGGACGACATGGAGAAAA-3′. qPCR was performed using a Takara RNA PCR kit (AMV) Version 3.0 (Takara Bio Inc.). The reaction conditions were as follows: Pre-denaturation at 94°C for 2 min followed by 31 cycles of denaturation at 94°C for 30 sec, annealing at 94°C for 30 sec and extension at 72°C for 30 sec, then a final extension at 72°C for 8 min. PCR amplicons were separated on 1.5% agarose and analyzed using a gel imaging analysis system (Molecular Imager Gel DocTM XR+ system; Bio-Rad).

The activity of caspase-3 was determined using the Caspase-3 activity kit (Beyotime Institute of Biotechnology, Haimen, China). To evaluate the activity of caspase-3, cell lysates were prepared following their respective treatment with various designated treatments. 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 mM Tris-HCl (pH 7.5), 137 mM Nad and 10% glycerol) containing 10 μl caspase-3 substrate (Ac-DEVD-pNA) (2 mM]. Lysates were incubated at 37°C for 4 h.

All results are presented as the mean ± standard error of the mean. The effects of the chemicals at different concentrations were analyzed using the analysis of variance method. Differences between groups were analyzed using the unpaired Student’s t-test. A value of P<0.05 was considered to indicate a statistically significant difference.

In human liver cancer cells cultured in media containing H₂O₂ for 12 h, the number of adhesive cells was observed to decrease and the morphology of the cells was observed to become round- or oval-shaped. These features were enhanced in a H₂O₂ concentration-dependent manner. Flow cytometry revealed that upon H₂O₂ treatment, the number of apoptotic cells increased in a concentration-dependent manner (Fig. 1A). The absorbance (A)_570nm value of the liver cancer cells was also found to decrease in a concentration-dependent manner following H₂O₂ addition (Fig. 1B).

Caspase family members have key roles in apoptosis. In the present study, caspase-3 and -8 expression were observed to increase in a concentration-dependent manner with H₂O₂ treatment, which was consistent with the increase in apoptosis observed (Fig. 1C). In order to investigate whether the H₂O₂-induced apoptosis was associated with the mitochondrial pathway in human liver cancer cells, mitochondrial and cytoplasmic levels of cytochrome c, as well as changes in mitochondrial membrane potential, were analyzed. H₂O₂ was found to significantly decrease the mitochondrial membrane potential (Fig. 1D). Furthermore, levels of cytochrome c in the cytoplasm were observed to significantly increase (Fig. 1E and G), while those in the mitochondria gradually decreased (Fig. 1F and G). These findings suggested that H₂O₂-induced human liver cancer cell apoptosis proceeded through the mitochondrial pathway.

Upon treatment with 200 mM H₂O₂, the addition of various concentrations of DADLE was found to increase the A_570nm value of liver cancer cells to various degrees. Increases in DADLE concentration from 0.01 to 1 μM were observed to increase the A_570nm value of liver cells in a dose-dependent manner. However, at concentrations >1 μM, no further increases were observed in the A_570nm value of liver cancer cells. These findings suggested that DADLE had a dose-dependent protective effect against H₂O₂-induced apoptosis in human liver cancer cells, with a maximum effect at 1 μM (Fig. 1H).

To study the effect of activated δ-opioid receptors on human liver cancer cell apoptosis, apoptosis was induced in liver cancer cells using H₂O₂. Cells were then treated with 1 μM DADLE, a specific δ-opioid receptor agonist. Annexin V-FITC/PI double-staining and flow cytometry revealed that the apoptosis rate of the cells treated with 200 mM H₂O₂ for 12 h was significantly increased compared with that in the control group. Furthermore, upon DADLE-induced δ-opioid receptor activation, the apoptosis rate was found to significantly decrease compared with the cells treated solely with H₂O₂. Moreover, when δ-opioid receptor activation was inhibited using 10 μM naltrindole, a δ-opioid receptor antagonist, the DADLE-induced protective effect was reverted (Fig. 2A and B). Caspase-3 and -8 expression was observed to increase rapidly following H₂O₂ treatment. δ-opioid receptor activation was found to significantly downregulate cytoplasmic caspase-3 and -8 levels, and naltrindole was observed to inhibit this protective δ-opioid receptor-induced effect (Fig. 2C). These findings suggested that δ-receptor activation significantly inhibits H₂O₂-induced apoptosis in human liver cancer cells.

To investigate whether the molecular mechanisms underlying the inhibition of liver cancer cell apoptosis by activated δ-opioid receptors are associated with the mitochondrial pathway, changes in mitochondrial membrane potential were analyzed. H₂O₂ treatment was found to gradually decrease the mitochondrial membrane potential in liver cancer cells. Concurrent δ-opioid receptor activation and H₂O₂ treatment had no significant effect on the mitochondrial membrane potential (Fig. 3A). Cytochrome c is released from mitochondria into the cytoplasm during apoptosis; therefore, western blot analysis was performed to investigate the cytoplasmic and mitochondrial cytochrome c levels. Compared with the cells treated with H₂O₂ alone, upon activation of the δ-opioid receptors with H₂O₂ treatment, cytoplasmic cytochrome c levels were observed to decrease and mitochondrial cytochrome c levels were found to increase (Fig. 3B and C). Furthermore, δ-opioid receptor activation was observed to increase cytoplasmic Bax and decrease Bcl-2 expression (Fig. 3D and E). These findings suggested that DADLE may activate δ-opioid receptors at the surface of the plasma membrane in liver cancer cells in order to stabilize mitochondrial membrane potentials and inhibit H₂O₂-induced apoptosis in human liver cancer cells.

In order to investigate whether δ-opioid receptors activate the PKC/ERK signaling pathway and whether inhibiting this signaling pathway alters the effect of δ-opioid receptors on human liver cancer cell apoptosis, the phosphorylation levels of PKC and ERK were investigated. Following δ-opioid receptor activation, phosphorylated PKC and ERK were observed to be significantly increased in the cytoplasm of human liver cancer cells (Fig. 4A and B), suggesting that δ-opioid receptor activation may lead to phosphorylation of PKC and ERK. It has previously been reported that δ-opioid receptor activation inhibits apoptosis in human liver cancer cells (23). However, in the present study, inhibiting the PKC pathway was found to increase apoptosis and inhibit cell proliferation, regardless of the δ-opioid receptor activation status. Inhibition of the ERK pathway was observed to have the same effect (Fig. 4C and D). These findings suggested that the PKC and ERK pathways are involved in mediating the protective effect of δ-opioid receptors on H₂O₂-induced human liver cancer cell apoptosis.

To assess whether δ-opioid receptors are expressed in human liver cancer tissues and cells, and whether they have a role in carcinogenesis, qPCR analysis was used to assess δ-opioid receptor mRNA expression in 50 liver cancer samples. δ-opioid receptors were found to be expressed in the 50 liver cancer samples, with the expression levels observed to be higher than those in the adjacent normal liver tissue. δ-opioid receptor mRNA expression was also analyzed in several liver cancer cell lines (Fig. 5A). Western blot analysis further revealed that the protein expression of the δ-opioid receptor was higher in liver cancer tissue than in the adjacent normal tissue. In addition, δ-opioid receptor protein expression was found to be higher in the liver cancer cell lines than in the normal liver cell lines (Fig. 5B). These findings indicate that δ-opioid receptors are highly expressed in human liver cancer tissues and cells and have an important role in liver cancer cell proliferation.