SH-SY5Y (neuroblastoma cell line) (CRL-2266), HepG2 (liver cancer cells) (HB-8065), HT29 (colorectal cancer cells) (HTB-38), and 4T1 (breast cancer cells) (CRL-2539) were purchased from the American Type Culture Collection (ATCC) and authenticated using short tandem repeat analysis (also conducted by the ATCC). The cells were maintained in Eagle's minimum essential medium (Gibco; Thermo Fisher Scientific, Inc.) supplemented with 10% fetal bovine serum (Amresco, LLC), 100 units/ml penicillin and 100 µg/ml streptomycin (Gibco; Thermo Fisher Scientific, Inc.) at 37°C in 5% CO₂. LNCaP cell were maintained in RPMI-1640 (Gibco; Thermo Fisher Scientific, Inc.) supplemented with 10% fetal calf serum, 100 units/ml penicillin, and 100 µg/ml streptomycin at 37°C in 5% CO₂.

SH-SY5Y cells, HepG2, HT29, 4T1, and LNCaP cells were plated in 96-well microplates at 4×10⁵ cells/wells and then incubated for 48 h. Cells were treated with 7-MH at different concentrations and reference compound for 24, 48 and 72 h. Then, 10 µl of MTT reagent (5 mg/ml) (Sigma-Aldrich; Merck KGaA) was added. The cells were incubated for 2 h until purple precipitate was visible after addition of dimethyl sulfoxide. The absorbance at 570 nm was measured. The percentage calculation of cell viability was carried out using the following formula: % Cell viability=(Absorbance of treated cells ×100)/Absorbance of control (untreated cells). Cell morphology was examined using a phase-contrast microscope by having 36 µM doxorubicin (Sigma-Aldrich) as positive control.

SH-SY5Y cells were plated in 96-well microplates at a density 4×10⁵ cells/wells and then incubated for 48 h. The cells were treated with various concentrations of 7-MH or 100 µM NAC for 2 h. Then, the cells were treated with 250 µM H₂O₂ for 4 h to induce oxidative stress. Cell viability was determined by MTT colorimetry. Absorbance was measured at 570 nm.

Apoptosis occurs as a result of G0/G1 phase arrest. Apoptosis as a protective mechanism ensures homeostasis of host cells through cell shrinkage, fragmentation of cellular DNA and formation of ‘apoptotic bodies’ subsequently leading to cell death. For cell cycle analysis, the cells were treated with various concentrations of 7-MH for 2 h. Then, the cells were treated with H₂O₂ for 4 h to induce oxidative stress. The cells were fixed by ethanol for 2 h at 4°C and stained with 50 mg/ml propidium iodide (PI) for 30 min in the presence of RNase before analysis. The percentage of apoptotic cells was quantitated using an FACScan flow cytometer with BD FACSDivaTM software (v. 6.1.3) (BD FACSAria; BD Biosciences). Late apoptotic cells were distinguished from non-apoptotic, intact cells by their decreased DNA content, which was determined by their low PI staining intensity.

In order to investigate the mechanisms of interaction with the apoptotic pathway in cancer cells, the cells were plated in six-well plates at a density of 1×10⁶ cells/wells and then incubated for 48 h. The cells were treated with various concentrations of 7-MH at 30 min, and cell death was induced with H₂O₂ at 15 min for SH-SY5Y cells. In HepG2, HT29, 4T1, and LNCaP cells, the cells were treated with various concentrations of 7-MH at the indicated time. Whole-cell lysates were prepared with lysis buffer [25 mM HEPES, pH 7.7, 0.3 mM MgCl₂, 0.2 mM EDTA, 10% Triton X-100, 20 mM β-glycerophosphate, 1 mM sodium orthovanadate, 1 mM phenylmethylsulfonyl fluoride (PMSF), 1 mM dithiothreitol (DTT), 10 µg/ml aprotinin, and 10 µg/ml leupeptin] (Gibco; Thermo Fisher Scientific, Inc.). The cell lysates were collected from the supernatant after centrifugation at 2,500 × g for 10 min 4°C.

The total protein concentration was measured by using the Bradford dye-binding method (Bio-Rad). The cell lysates (15 µl)were loaded and resolved by 7.5-12.5% SDS-PAGE and transferred to an Immobilon-P-nylon membrane (MilliporeSigma). The membrane was treated with BlockAcc (Dainippon Phamaceutical Co., Ltd.) and probed at room temperature for 2 h with the following primary antibodies: anti-caspase-3 (cat. no. 9662), GSK-3 (cat. no. 5558), phospho-p38 (cat. no. 4511), p38 (cat. no. 54470), Mcl-1 (cat. no. 94296), Bcl-xL (cat. no. 2764), BAX (cat. no. 5023), phospho-Akt (cat. no. 4060), Akt (cat. no. 4691), phospho-ERK (cat. no. 9911), phospho-P65 (cat. no. 3031), P65 (cat. no. 3033), Bcl-2 (cat. no. 4223), survivin (cat. no. 2808), MAPK13 (cat. no. 4511), and anti-actin antibodies (cat. no. 3700), all diluted at 1:1,000 and obtained from Cell Signaling Technology, Inc. The antibodies were detected using horseradish peroxidase-conjugated anti-rabbit (cat. no. 14708), anti-mouse (cat. no. 14709), and anti-goat IgG (cat. no. 98164) secondary antibodies (1: 5,000; Cell Signaling Technology, Inc.) and visualized using the enhanced chemiluminescence system (Life Science Technology). Densitometric analysis of western blot bands was performed using ImageJ software (version IJ 1.46 r; National Institutes of Health.).

The cancer cell migration and invasion capacities were determined using a Transwell assay. A total of 1×10⁶ cells/well in serum-free DMEM were added into the top chambers of a 24-well insert (pore size, 8-mm; Corning Life Sciences) and incubated for 4 h to allow the cells to attach. The cells were treated with candidone and incubated for 24 h at 37°C. For the invasion assays, the membranes were pre-coated with collagen for 4 h at 37°C. The lower chambers were filled with DMEM supplemented with 10% fetal bovine serum as a chemoattractant. After 4 h, the migratory/invasive cells were fixed with a 10% formalin solution at room temperature for 30 min and then stained with 0.05% crystal violet solution at room temperature for 30 min, after which the stained cells were counted under an inverted light microscope (magnification, ×20).

4T1 cells were transfected with a luciferase reporter plasmid (Promega Corporation) using Lipofectamine^® 2000 (Invitrogen; Thermo Fisher Scientific, Inc.) under the control of NF-κB and STAT3 sites containing a neo-resistance gene. The luciferase activity of transfected cells was compared with 4T1 CMV control cell (the 4T1 cells stably expressing luciferase with a CMV-promoter). A stable clone was isolated in medium containing 500 µg/ml G418 (Thermo Fisher Scientific, Inc.). Cells were seeded in a 96-well plate and treated with DMSO (control) and 0.1, 1, 10, or 100 µM of 7-MH for 24 h and compared with resveratrol as a positive control. Luciferase activity was measured using the Dual-Luciferase reporter assay system (Promega Corporation).

Female BALB/c mice (6 weeks old) were separated into groups of 6–7 mice at temperature and humidity of 23±2°C and 50±10%, respectively, and were administered food and water ad libitum. The experiment was carried out according to the standard guidelines of the National Institutes of Health and was approved (approval no. A2017INM-7) by the Animal Care and Use Committee of the University of Toyama (Toyama, Thailand). The lung metastasis model was performed by culturing mouse mammary carcinoma 4T1 cells with the luciferase gene (4T1-Luc2 cells), and then the cells were harvested and resuspended in cold phosphate-buffered saline (PBS). The cell suspensions (5×10⁵/50 µl/mouse) were inoculated in mice by intravenous injection (the number of mice was n=6 each group and repeated twice so at least 24 mice as total from 30 prepared mice). After 6 days, D-luciferin (150 mg/kg) was injected, and 20 min later, the lungs were harvested from the mice. Lung luminescence was then determined using an imaging system (IVIS Spectrum; Caliper Life Science). The method of euthanasia was cervical dislocation and death was confirmed before disposal of the animal by evaluating consciousness including lack of a heartbeat, lack of respiration, lack of corneal reflex and presence of rigor mortis. In accordance with the Guide for the Care and use of Laboratory Animals of the National Institutes of Health, the humane endpoint was set based on the percentage of body weight loss (20% body-weight reduction).

The TAK1 kinase template was prepared from 4GS6 and validated by redocking with the irreversible inhibitor (5Z)-7-oxozeaenol. All hydrogens were added, water molecules were deleted, and Gasteiger charges were assigned to the TAK1 kinase template and all ligands by using AutoDockTools (ADT). The AutoGrid was used to generate grid maps. The grids were designated to include the active site of TAK1 kinase. The grid box dimensions were defined as 100×100×100 Å, and the grid spacing was set to 0.375 Å. All ligands were docked using the Lamarckian genetic algorithm via the Autodock 4.2.6 auxiliary program. The Lamarckian genetic algorithm protocol was set to a population size of 150 individuals with 150 ligand orientation runs. Additionally, the energy evaluation was 1,000,000, which was as the maximum number of evaluations. The docking complex poses were analyzed for their interactions by using BIOVIA Discovery Studio 2017.

The statistical technique used for the analysis was a one-way analysis of variance (ANOVA) followed by Tukey's post hoc test for comparison between 2 groups and between 3 or more groups. The data were analyzed using SPSS software (version 24; IBM Corp.). The analysis was performed in triplicate, and the values are presented as the mean ± SDs. P<0.05 was considered to indicate a statistically significant difference.

To investigate the effect of 7-MH on H₂O₂-induced neuronal cell death, neuronal cells were treated with various concentrations of 7-MH or 100 µM NAC (reference compound) for 2 h before switching to 250 µM H2O2 for 4 h. Cell viability was assessed using MTT assay. The results showed significantly increased cell viability when compared with H₂O₂-insulted samples. The values obtained with 7-MH treatment at a concentration of 100 µM showed a stronger neuroprotective effect than that achieved by NAC treatment (Fig. 2A). A decrease in morphologically-confirmed cell death was observed as a result of 7-MH, compared with 100 µM NAC (Fig. 2B). In consistency with previous findings, 7-MH showed a neuroprotective effect on NG108-15 cells (mouse neuroblastoma and rat glioma cell lines) (35).

To verify the 7-MH inhibition of apoptosis by H₂O₂, the cells were labeled with PI and analyzed using flow cytometry (There was a limitation to stain with Annexin V). The results revealed the DNA content histograms obtained after the PI staining of cells that had been treated with various concentrations of 7-MH for 2 h and had been eliminated by treatment with an H₂O₂ concentration of 250 µM for 4 h. When the cells were incubated in the medium alone (control), a single peak of nuclei with diploid DNA content was observed. By contrast, when the cells were incubated in H₂O₂ alone (negative control), an increase in apoptotic cells in sub-G₀/G₁ was observed. When the cells were incubated in NAC and H₂O₂ (positive control), the results were similar to those of the control group. In the presence of 7-MH, inhibiting apoptosis with H₂O₂, apoptotic cells with increased DNA content were distinguishable. A characteristic hypodiploid DNA content peak, which shows sub-G₀/G₁ apoptotic populations, was observed following the treatment of neuronal cells with 7-MH in a concentration-dependent manner (Fig. 3).

To further evaluate the protective molecular mechanisms of 7-MH in neuronal cells, the cells were treated with various concentrations of 7-MH for 2 h, and cell death was induced with H₂O₂ for 4 h. Key proteins involved in apoptosis regulation were examined, including GSK-3, p-p38, Mcl-1, Bcl-2, and BAX, using an immunoblot assay. As demonstrated in Fig. 4, 7-MH markedly inhibited p-p38, BAX, and cleaved caspase-3 compared with the control; and induced Mcl-1, Bcl-2, and Bcl-xL in a concentration-dependent manner. The results indicated that 7-MH efficiently inhibits the apoptotic effect of H₂O₂ induced in neuronal cells.

In order to elucidate the molecular mechanism of cancer cellapoptosis, the GSK-3 signaling pathways were assessed. Neuroblastoma cells were treated with various concentrations of 7-MH for 24 h. Cell viability was assessed using MTT assays. These results revealed that 7-MH at a concentration of 100 µM significantly induced cancer cell death with morphological changes, including cell rounding, shrinkage, and detachment (Fig. 5A and B). 7-MH activated the cleaving of caspase-3 by increasing the level of Bax and decreasing the levels of Mcl-1 and Bcl-xl, which are regulated by GSK-3 (Fig. 5C). This indicated that 7-MH induced apoptosis in SH-SY5Y cells via the GSK-3 pathway.

To test the effect of 7-MH on cancer migration and invasion, HT29 and HepG2 cancer cells were treated with various concentrations of 7-MH for 24 h. Cell viability was assessed using an MTT assay. The results showed that 7-MH at a concentration of 100 µM significantly induced cancer cell death in a time-dependent manner, due to its effect on HT29 being more potent than on HepG2 cells (Fig. 6A). The morphological changes, including cell rounding, shrinkage, and detachment, were observed in cells treated with 100 µM of 7-MH compared with 36 µM doxorubicin as a positive control (Fig. 6B). Moreover, 7-MH at concentrations of 1 and 10 µM inhibited the migration and invasion of HT29 cancer cells (Fig. 7A and B). The western blot results revealed that 7-MH activated the cleaving of caspase-3 (marker of apoptosis) and decreased the levels of phospho-Erk, phospho-p38, Bcl-2, survivin, and matrix metalloproteinase-9 (marker of metastasis) (Fig. 8A-C). The results indicated that 7-MH-induced cell death, inhibited migration, and invasion of HT29 cancer cells.

To examine the effect of 7-MH on the viability of LNCaP cells, the cells were treated with various concentrations of 7-MH for 24, 48 and 72 h, and the cell viability was examined using MTT assay. The result showed that 7-MH significantly inhibited cell growth at concentrations of 1, 10, and 100 µM for 24 h; and at concentrations of 10 and 100 µM for 48 h and 72 h (Fig. 9A). To confirm the effects of 7-MH on LNCaP cell proliferation, cells were treated with various concentrations of 7-MH for 24 h. 7-MH was observed to significantly inhibit cell growth in a dose dependent manner (Fig. 9B). In previous studies, 7-MH was observed to be a carbazole isolated from the roots of C. harmandiana, which exhibited cytotoxicity against NCI-H187 (human small-cell lung cancer cells), KB (human epidermoid carcinoma of oral cavity cell lines) and HT29 (human colorectal adenocarcinoma cell line) cells (36–38).

To understand the mechanism by which 7-MH induced cell death, multiple potential signaling pathways that have been demonstrated to be engaged in 7-MH-induced apoptosis were screened. Western blotting showed that 7-MH incubation leads to activation of Akt and p38, whereas the expression of p65, pERK, and MAPK13 was inhibited in a time-dependent manner (Fig. 10). This result indicated that 7-MH induced apoptosis by inducing Akt and p38 activation and inhibiting the p65, pERK, and MAPK13 pathways.

7-MH significantly reduced the viability of 4T1 cells when compared with resveratrol as a positive control (Fig. 11). The effect of 7-MH on the activation of NF-κB and STAT3 was examined in 4T1 cells stably transfected with an NF-κB- and STAT3-dependent reporter plasmid. Cells were treated with 7-MH for 6 h. It was found that 7-MH inhibited NF-κB and STAT3 activation (Fig. 12).

In 4T1 cancer cell metastasis, the Transwell assay showed that 10 µM of 7-MH significantly inhibited 4T1 cancer cell migration (Fig. 13). In vivo assay showed that 7-MH reduced the luminescence signal of 4T1-Luc2 cell metastasis in the lungs of mice (Fig. 14).

To understand the binding interactions between 7-MH and TAK1 kinase, a molecular docking study utilizing the Autodock 4.2.6 program was performed. The 4GS6 PDB code, which is bound with the irreversible inhibitor (5Z)-7-oxozeaenol, was used as the TAK1 template. The binding modes and interaction diagrams of 7-MH bound to TAK1 kinase are represented in Fig. 15. The results of docking revealed that 7-MH exhibited multiple binding sites with TAK1 kinase, which are likely to be Lys63, Met104, Tyr106, Ala107, Leu163, Pro160, Cys174, ASP175, Val42, Val50, Val90, Ala61, Gly43, Gly45, Gly110 and Ser111; and its binding energy (ΔG) is-7.72 kcal/mol.