Wogonin was purchased from Sigma-Aldrich (St. Louis, MO, USA) and dissolved in dimethylsulfoxide (DMSO; Sigma-Aldrich) as a stock solution of 100 mM. Wogonin was further diluted in RPMI-1640 (Invitrogen, Big Cabin, OK, USA) plus 10% fetal bovine serum (FBS; Invitrogen, Grand Island, NY, USA) to the appropriate final concentrations. The primary polyclonal rabbit anti-human antibodies: PI3K, phosphorylated (p)-PI3K (Tyr458), AKT, p-AKT (Ser473), PARP-1, pancreatic ER stress kinase (PERK), eukaryotic initiation factor 2α (eIF2α), activating transcription factor 6 (ATF6), inositol-requiring enzyme 1α (IRE1α) and β-actin were obtained from Cell Signaling Technology (Beverly, MA, USA). The secondary horseradish peroxidase (HRP)-labeled mouse anti-rabbit IgG polyclonal antibodies for western blot analysis were provided by Merck Millipore (Beijing, China). Annexin V-fluorescein isothiocyanate (FITC) and propidium iodide (PI) were purchased from BD Biosciences (Palo Alto, CA, USA), SH-6 was provided by Santa Cruz Biotechnology (Dallas, TX, USA).

The adenoviral constructs were generated using a protocol from Qbiogene’s AdEasy vector system (Carlsbad, CA, USA) with the following modifications. The constitutively active AKT constructs were amplified by PCR and cloned into the pShuttle-CMV adenovirus transfer vector (BD Clontech Laboratories, Palo Alto, CA, USA). HA-AKT was cloned into the HindIII and EcoRV sites and was transformed into BJ5183 bacteria, resulting in Ad-HA-AKT. All constructs were purified using Qiagen Plasmid Maxiprep kits according to the manufacturer’s instructions. The constructs were confirmed by restriction enzyme digestion and sequencing analysis.

The adenoviruses were amplified by infecting human acute promyelocytic leukemia HL-60 (ATCC^® CCL-240™) cells. Infected cells were harvested and centrifuged at 1,250 × g at 4°C for 10 min, and the resulting cellular lysate pellet was resuspended in Dulbecco’s modified Eagle’s medium (DMEM) without supplements. The cells were disrupted using three freeze/thaw cycles and the suspension was subsequently spun at 1,250 × g at 4°C for 15 min to release the virus particles. The supernatant containing the viral particles was spun at 100,000 × g (Beckman SW28 rotor) for 16 h at 4°C through an isopycnic CsCl gradient. The viral band was isolated and dialyzed against 10% glycerol in phosphate-buffered saline (PBS; 137 mM NaCl, 2.7 mM KCl, 1.47 mM KH₂PO₄ and 8.1 mM Na₂HPO₄, pH 7.6) for 16–24 h at 4°C. The viral particle concentration (titer) was determined by absorbance at 260 nm. viruses were aliquoted and stored at −80°C until used.

The HL-60 suspension cells were purchased from the Peking Union Medical College Cell Library (Beijing, China). The cells were cultured in RPMI-1640 supplemented with 100 U/ml of penicillin, 100 μg streptomycin and 10% FBS at 37°C in a humidified atmosphere containing 5% CO₂. The media were changed every 2–3 days and subcultured when the cell population density reached 70–80% confluency. Cells were seeded at an appropriate density according to each experimental design.

Cell lysates (30 μg) from the HL-60 cells obtained after treatment with wogonin at the desired doses and time periods, were analyzed for caspase acivity spectrophotometrically at 405 nm using a microtiter plate reader. The assays were performed by incubating the cell lysates with 0.2 mM of the caspase-specific colorimetric tetrapeptide substrates, Ac-LEHD-p-nitroaniline (pNA) (for caspase-9), Ac-IETD-pNA (for caspase-8) or Ac-DEVD-pNA (for caspase-3) for 1 h at 37°C as described in a previous study (31). The increase in the absorbance at 405 nm which corresponds to the amount of pNA liberated from the peptide substrates was converted into units of enzyme activity using a standard curve generated with free pNA. One unit of caspase-3, 8 or 9 activity corresponded to the amount of enzyme that will release 1 pmol of pNA from 0.2 mM DEVD-pNA, IETD-pNA or Ac-LEHD-pNA/min, respectively. Lysates from the HL-60 cells treated with DMSO were also used in these assays as the control group.

Cell viability was assessed using the MTT assay. HL-60 cells were plated into 96-well clusters at a density of 5×10⁴ cells/well. After a 24-h incubation under group-specified experimental conditions, the clustered HL-60 cells were processed for detection of cell viability by MTT assays. Spent medium was removed and 10 μl MTT solution (5 mg/ml) was added to 100 μl of respective growth medium without phenol red, and the plates were incubated at 37°C for 4 h in a humidified 5% CO₂ atmosphere. Then, the formazan crystals formed by mitochondrial reduction of MTT were solubilized in DMSO (100 μl/well), and the absorbance was read at 540 nm using a microplate reader (Bio-Rad, Hercules, CA, USA). Percent inhibition of cytotoxicity was calculated as a fraction of the control group and expressed as a percentage of cell viability.

Double-staining for Annexin V-FITC and PI was performed to estimate the apoptotic rate of the HL-60 cells. Briefly, after incubation with various doses of wogonin for 48 h, the HL-60 cells were trypsinized and washed twice with PBS, and centrifuged at 800 rpm for 5 min. Then, 1×10⁶ cells were suspended in binding buffer and double-stained with Annexin V-FITC and PI for 30 min at room temperature. Subsequently, the fluorescence of each sample was quantitatively analyzed by a FACSCalibur flow cytometer and CellQuest software. The results were interpreted as follows: PI-positive and Annexin V-FITC-positive stained cells were considered as apoptotic.

Total messenger RNA from the HL-60 cells was isolated using the RNeasy Mini kit (Qiagen, Valencia, CA, USA) according to the manufacturer’s instructions. The amount and quality of the RNA were determined using a NanoDrop 2000 spectrophotometer (Thermo Scientific, Wilmington, DE, USA). First-strand cDNA was synthesized from 300 ng of RNA with SuperScript^® III First-Strand Synthesis system (Applied Biosystems, Foster City, CA, USA) and used as the template for PCR with the FastStart HiFi PCR system (Roche Applied Science, Indianapolis, IN, USA). The fold-change in gene expression was obtained by dividing the treated group signal by that of the base expression level signal of the corresponding gene in the untreated cells. Results were normalized using qRT-PCR signal from β-actin of the respective samples.

The primers of the target genes were: glucose-regulated protein 78 (GRP78) forward, 5′-TCT GCT TGA TGT GTG TCC TCT T-3′ and reverse, 5′-GTC GTT CAC CTT CGT AGA CCT-3′; C/EBP-homologous protein (CHOP) forward, 5′-GGA GAA GGA GCA GGA GAA TGA-3′ and reverse, 5′-AGA CAG ACA GGA GGT GAT GC-3′; GRP94 forward, 5′-TCG GGA AGC AAC AGA GAA GG-3′ and reverse, 5′-TCA TCT TCC TTA ACC CTC CGC-3′; Bcl-2 forward, 5′-GCT GAG GCA GAA GGG TTA TG-3′ and reverse, 5′-GCC CCC TTG AAA AAG TTC AT-3′; Bax forward, 5′-AGG GTT TCA TCC AGG ATC GAG CAG-3′ and reverse, 5′-ATC TTC TTC CAG ATG GTG AGC GAG-3′; β-actin forward, 5′-GGC GGA CTA TGA CTT AGT TG-3′ and reverse, 5′-AAA CAA CAA TGT GCA ATC AA-3′.

For the western blot analysis, HL-60 cells were harvested, washed once in ice-cold PBS, gently lysed in ice-cold lysis buffer (250 mM sucrose, 1 mM EDTA, 0.05% digitonin, 25 mM Tris, pH 6.8, 1 mM dithiothreitol, 1 μg/ml leupeptin, 1 μg/ml pepstatin, 1 μg/ml aprotinin, 1 mM benzamidine and 0.1 mM phenylmethylsulphonyl fluoride) for 30 min, and centrifuged at 12,000 rpm at 4°C. The protein concentration was measured using the BioRad Bradford protein assay reagent, and subjected to SDS-PAGE. Proteins were transferred to polyvinylidene fluoride membranes and incubated successively in 5% bovine serum albumin in Tris-buffered saline Tween-20 buffer (25 mmol/l Tris, pH 7.5, 150 mmol/l NaCl and 0.1% Tween-20) for 1 h, then incubated overnight at 4°C with PI3K, p-PI3K (Tyr458), AKT, p-AKT (Ser473), PARP-1, PERK, eIF2α, ATF6, IRE1α or β-actin, followed by reaction with HRP-labeled mouse anti-rabbit IgG polyclonal antibodies for 1 h. After each incubation, the membranes were washed extensively in TTBS and the immunoreactive band was detected using ECL-detecting reagents.

Results are expressed as the mean ± SEM. Statistical comparisons were performed using a Student’s t-test, one-way analysis of variance (ANOVA), or two-way repeated measures ANOVA with the Student’s t-test for post hoc analyses. In all cases, values of P<0.05 were considered to indicate statistically significant results.

In the pre-experiment, we applied wogonin at doses of 10, 25, 50, 75, 100 and 150 μM. The results showed that wogonin inhibited the viability of HL-60 cells in a dose-dependent manner (Fig. 1a) and the effect was predominant at 75–150 μM. In addition, HL-60 cells were treated with 75 μM wogonin for 6, 12, 24, 48, 72 and 96 h. We found that 75 μM wogonin inhibited the viability of the HL-60 cells in a time-dependent manner (Fig. 1b) and the effect was predominant at 48–96 h. Therefore, HL-60 cells were treated with wogonin for 48 h in the subsequent experiments.

To further investigate whether wogonin induces the apoptosis of HL-60 cells, the cells were treated with 75 μM wogonin for 48 h and the apoptotic rate of the HL-60 cells was detected using Annexin V-FITC/PI staining. The results revealed that 75 μM wogonin induced apoptosis in the HL-60 cells (Fig. 1c).

Apoptosis induced by various cytotoxic agents is highly dependent on the activation of caspases, which play pivotal roles in cleaving specific target proteins (32). As shown in Fig. 1c, wogonin caused strong apoptotic death of the HL-60 cells. Therefore, we assessed whether wogonin activates caspase pathways in the HL-60 cells. Cells were exposed to increasing doses of wogonin for 48 h, and then analyzed for caspase activation by colorimetric assays. As shown in Fig. 2a-c, wogonin triggered a dose-dependent increase in the specific activities of caspase-3, -8 and -9. A mean specific activity peak was noted after incubation of the HL-60 cells with doses of 75–150 μM wogonin for 48 h.

We next performed western blot analysis to determine the level of cleaved PARP-1 that induces the enhanced apoptosis. The cleavage of PARP-1 was significantly higher in the HL-60 cells following wogonin treatment in a dose-dependent manner. The maximum activation of cleaved PARP-1 was detected following treatment of wogonin at doses of 75–150 μM for 48 h (Fig. 2d and e). This result was in accordance with the activities of caspase-3, 8 and 9. In addition, wogonin treatment also resulted in significant reduction in total PARP-1 levels (Fig. 2d and e).

The activation of caspase-9 by wogonin suggested that the mitochondrial apoptotic pathway was triggered in the HL-60 cells. Since Bcl-2 family proteins are known to control the mitochondrial-mediated apoptosis pathway by maintaining a balance between pro- and anti-apoptotic members (33), we examined the effects of wogonin on the expression levels of Bcl-2 family proteins in the HL-60 cells. Our results showed that wogonin increased the mRNA level of pro-apoptotic Bax (Fig. 2f), but decreased the mRNA level of Bcl-2 (Fig. 2g) in HL-60 cells. In addition, the Bax/Bcl-2 ratio was increased following treatment with wogonin in the HL-60 cells (Fig. 2h). These data demonstrated that wogonin induced HL-60 cell apoptosis through mitochondrial-mediated mechanisms.

Treatment with wogonin has been shown to induce accumulation of misfolded nascent glycoproteins in the ER lumen in various types of cancer cells, leading to ER stress and UPR activation (23,24,26,34). The expression of ER stress markers was analyzed in the HL-60 cells following treatment with wogonin. In the HL-60 cells, wogonin increased the mRNA expression of GRP94 (Fig. 3a), GRP78 (Fig. 3b) and CHOP (Fig. 3c) confirming induction of ER stress, and their increased expression was correlated with cleaved PARP-1 and activity of caspase-3, -8 and -9, markers of apoptosis.

When ER stress occurs, GRP78 is released from three key branches: PERK, IRE1 and ATF6 and binds misfolded proteins, thereby activating the UPR (35,36). Next, we analyzed the expression levels of the following key UPR signal transduction molecules: PERK, eIF2α, ATF6 and IRE1α. Exposure of the HL-60 cells to wogonin resulted in the upregulated expression of p-PERK, p-eIF2α and cleaved ATF6 and activation of IRE1α (Fig. 3d-f). These data suggested that wogonin induced HL-60 cell apoptosis through ER stress-mediated mechanisms.

To further understand the molecular mechanisms by which wogonin induces HL-60 cell apoptosis, we examined the expression of PI3K-AKT, critical signaling proteins associated with cell apoptosis. The levels of p-PI3K and p-AKT were detected, and the results demonstrated that wogonin significantly blocked the constitutive phosphorylation of PI3K at Tyr458 and AKT at Ser473 in a dose-dependent manner (Fig. 4a).

To determine the role of PI3K-AKT in wogonin-induced apoptosis and ER stress in HL-60 cells, constitutively active HA-tagged AKT constructs were made in AKT at Ser473, which were mutated to an aspartic acid residue to mimic phosphorylated AKT. The expression of HA-AKT significantly increased the phosphorylated AKT activity (Fig. 4b) after wogonin treatment. Active AKT significantly increased the cell viability (Fig. 5a) and reduced the percentage of HL-60 apoptotic cells (Fig. 5b) when compared with the cells in the wogonin only treatment group. Moreover, active AKT also reduced wogonin-induced caspase-3, -8, -9 (Fig. 5c) and PARP-1 (Fig. 5d) activities and the Bax/Bcl-2 ratio (Fig. 5e-g) when compared with cells in the wogonin only treatment group. In addition, upregulated expression of p-PERK, p-eIF2α and cleaved ATF6 and activation of IRE1α induced by wogonin were reduced in the HL-60 cells expressing active AKT (Fig. 6a-c). These results indicated that the pro-apoptotic effects of wogonin in HL-60 cells were associated with inhibition of the activation of the PI3K/AKT signaling cascade.