Apigenin, acridine orange, 3-methyladenine (3-MA), and monoclonal antibody against β-actin were purchased from Sigma-Aldrich (St. Louis, MO, USA). Apigenin was dissolved in dimethylsulfoxide (DMSO) and stored at −20°C before the experiments and dilutions were made in culture medium. 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT) was obtained from Amresco (Solon, OH, USA). Antibodies against beclin-1, Cdc2, Cdc25c, cyclin B1, p21, p53, poly (ADP-ribose) polymerase (PARP), caspase-3, caspase-8 and caspase-9 were purchased from Santa Cruz Biotechnology (Santa Cruz, CA, USA). Polyclonal antibody against LC3B was obtained from Cell Signaling Technology (Danvers, MA, USA). RPMI-1640, fetal bovine serum (FBS), and penicillin-streptomycin were purchased from HyClone (Logan, UT, USA).

The human colorectal cancer cell line HCT116 was purchased from the American Type Culture Collection (Manassas, VA, USA) and maintained at 37°C in a humidified 95% air and 5% CO₂ in RPMI-1640 supplemented with 10% FBS, 100 U/ml penicillin and 100 μg/ml streptomycin.

The cell growth was determined by MTT assay. Cells were seeded in 6-well culture plate and incubated at 37°C for 24 h. Then, cells were treated with different concentration of apigenin (0–50 μM) for 24 h. The cells were incubated in the dark with MTT reagent (0.5 mg/ml) at 37°C for 2 h. Medium was removed, formazan was dissolved in DMSO and the absorbance at 540 nm was measured by using an ELISA plate reader.

Cells were washed twice with phosphate-buffered saline (PBS) and fixed with 3.7% paraformaldehyde (Sigma-Aldrich) in PBS for 10 min at room temperature. Fixed cells were washed with PBS, and stained with 4 μg/ml Hoechst 33342 for 20 min at room temperature. The stained cells were washed twice with PBS and analyzed by a fluorescent microscope.

Cells were harvested with trypsinization and washed once with PBS. After centrifugation, the cells were fixed in 70% ethanol at −20°C overnight. Fixed cells were prepared for flow cytometry analysis by washing twice with PBS and then stained in propidium iodide (PI) (Sigma-Aldrich) solution (50 μg/ml in PBS) for 30 min at room temperature in the dark. Flow cytometry analysis was performed on a Cytomic FC500 (Beckman Coulter, Istanbul, Turkey).

After apigenin treatment, cells were harvested and washed with cold PBS. Total cells were lysed in lysis buffer [40 mM Tris (pH 8.0), 120 mMNaCl, 0.5% NP-40, 2 μg/ml aprotinin, 2 μg/ml leupeptine and 100 μg/ml phenymethylsulfonyl fluoride (PMSF)]. After centrifugation, the supernatant was collected and protein concentration was determined by protein assay reagents (Bio-Rad, Hercules, CA, USA). Equal amount of protein extracts were denatured by boiling at 100°C for 5 min in sample buffer (Bio-Rad). The proteins were separated on subject to 6–15% SDS-PAGE and transferred to polyvinylidenedi fluoride (PVDF) membrane. The membranes were blocked with 5% non-fat dry milk in Tris-buffered saline with Tween-20 buffer (TBS-T) (20 mM Tris, 100 mM NaCl, pH 7.5 and 0.1% Tween-20) for 1 h at room temperature. The membranes were washed 10 min, 4 times with TBS-T buffer each for 10 min and incubated for 1 h with horseradish peroxidase-conjugated secondary antibodies (Santa Cruz Biotechnology). The membranes were washed once for 10 min, 4 times with TBS-T buffer. Antigen-antibody complex were detected by the enhanced chemiluminescence (ECL) detection system (GE Healthcare, Piscataway, NJ, USA).

The Annexin V-FITC is used to quantitatively determine the percentage of cells within a population that are actively undergoing apoptosis. Cells harvested were washed twice with cold PBS, suspended the cells in 1X binding buffer (BD Biosciences, San Diego, CA, USA). The counted cells were stained in PI and Annexin V-FITC solution (BD Pharmingen FITC Annexin V Apoptosis Detection kit, BD Biosciences) for 15 min at room temperature in the dark, and then, the stained cells were analyzed by flow cytometry.

The formation of acidic vesicular organelles (AVOs) is a well-known feature of autophagy. Cells were seeded in 6-well culture plate and incubated at 37°C for 24 h. Then, cells were treated with different concentration of apigenin (0–50 μM) for 24 h, and stained with acridine orange (1 μg/ml) for 15 min. Subsequently, cells were washed with PBS and examined under a fluorescent microscope. Depending on intracellular acidity, autophagic lysosomes appeared as orange/red fluorescent cytoplasmic vesicles, while the nuclei were displayed bright green. Alternatively, to quantify the development of AVOs, apigenin-treated cells were stained with acridine orange (1 μg/ml) for 15 min, trypsinized, and then washed with PBS. The stained cells were then analyzed using a flow cytometer.

HCT116 cells were transfected with LC3-GFP plasmid using Lipofectamine 2000 reagent (Invitrogen, Grand Island, NY, USA) according to the manufacturer’s instructions. Cells were treated with 3-MA (5 mM), apigenin (25 μM) or apigenin + 3-MA for 24 h and the formation of punctuate LC3-positive structures was observed by confocal microscopy.

Results are expressed as the mean ± SD of three separate experiments and analyzed by Student’s t-test, and considered significantly different at p<0.05 or p<0.01.

To investigate the growth inhibitory effect of apigenin (Fig. 1A), the MTT assay was performed in HCT116 cells. As shown in Fig. 1B, apigenin inhibited effectively cell proliferation of HCT116 cells in a concentration-dependent manner. The lower concentrations of apigenin (6.25 μM) did not affect cell viability; however, the higher concentrations (25 and 50 μM) of apigenin significantly reduced the cell viability of HCT116 cells. Additional experiment was done to confirm the growth inhibitory activity of apigenin on HCT116 cells when analyzed by microscopic observations. As shown in Fig. 1C, cells treated with apigenin displayed distinct morphological changes compared with untreated cells. In particular, cells were rounded and cell number was decreased in a concentration-dependent manner after apigenin treatment.

To identify the mechanism responsible for apigenin-induced cell growth inhibition, cell cycle progression was evaluated by flow cytometry analysis, as shown in Fig. 2A, apigenin treatment caused a significant cell cycle arrest in the G2/M phase concentration-dependently. An accumulation of cells in G2/M phase of 9.99, 20.1, 27.2 and 29.5% was observed for 6.25, 12.5, 25 and 50 μM, respectively, when compared with untreated cells (7.12%). Next, we examined whether apigenin can modulate the expression of G2/M cell cycle regulators. Cells were treated with various concentrations of apigenin for 24 h and then the level of G2/M cell cycle regulatory proteins were examined by western blot analysis. As indicated in Fig. 2B, after the cells were exposed to apigenin, the levels of p53 and its downstream protein p21^CIP1/WAF1, a potent cyclin-dependent kinase (CDK) inhibitor in G1 and G2/M phases, were increased in a concentration-dependent manner. Therefore, in agreement with previous findings (Fig. 2A), apigenin treatment decreased Cdc25c and its regulatory protein Cdc2 as well as cyclin B1 (Fig. 2B). These data suggest the possibility that apigenin-induced growth inhibition of HCT116 cells was the result of G2/M arrest.

To determine whether the growth inhibitory effects of apigenin were due to apoptotic cell death, the morphological changes of cellular structures were assessed with Hoechst 33342 staining. The untreated control cells displayed intact nuclear structure, while cells were treated with apigenin resulted in chromosomal condensation and formation of apoptotic bodies, which are characteristics of programmed cell death (Fig. 3A, arrows). The effect of apigenin on apoptosis was further verified by western blot analysis. Apigenin treatment decreased the protein levels of procaspase-8, -9 and -3 in concentration-dependently. In accordance with the decrease of procaspases with consequent increase of the levels of active forms of caspases, the cleavage of PARP was increased in a concentration-dependent manner (Fig. 3B). These results suggested that apigenin induced apoptotic cell death in HCT116 cells.

Autophagy, an evolutionally conserved self-digestive process, is known as a cell survival mechanism during nutrient depletion and is essential to cellular homeostasis maintenance by facilitating the disposal of unfolded proteins and cellular constitutes (25). It has been reported that inhibition of autophagy has potential to promote apoptosis induced by several anticancer agents, which suggested that autophagy may play a protective role in cancer cells (26,27). Recently, apigenin has been reported to induce autophagy in human breast cancer cells (28). To determine whether apigenin induces autophagy in HCT116 cells, we examined the effect of apigenin on the formation of acidic vesicular organelles (AVOs) in HCT116 cells. For this, we used acridine orange (AO), which accumulates in acidic cell compartments and displays bright red fluorescence (29). The formation of AVOs can be detected by fluorescence microscopy and quantified by flow cytometry. As shown in Fig. 4A, apigenin treatment caused progressive increase of AVO formation in HCT116 cells. In agreement with microscopic observation, flow cytometry analysis also showed a concentration-responsive increase in AVOs in apigenin-treated cells as compared with their respective untreated control cells (Fig. 4B).

It is known that the beclin-1 level and LC3 conversion (LC3-I to LC3-II) are selective markers of autophagy. As shown in Fig. 4C, apigenin treatment markedly increased the expression of LC3-II, suggestive of autophagy induction. However, apigenin did not affect the expression of beclin-1 significantly. To further confirm the induction of autophagy by apigenin, we utilized fluorescence microscopy to examine autophagosome formation in HCT116 cells transiently expressing GFP-LC3 protein. Due to the conversion of LC3-I (a soluble form) to the LC3-II (a lipidized form), which associates with the membranes of autophagosomes (30), this conversion can be detected by observing the formation of punctate structures. Treatment of these cells for 24 h with apigenin resulted in relocalization of the GFP-LC3 protein to punctate cytoplasmic dots, indicators of autophagosome formation. Furthermore, co-treatment with apigenin and 3-MA, inhibits autophagy at initiation of double membrane encapsulation, significantly inhibiting GFP-LC3 dot formation in HCT116 cells (Fig. 4D).

Accumulating data suggest that inhibition of autophagy may enhance chemosensitization in human cancer cells (31). Thus, we determined whether inhibition of autophagy could potentiate apigenin-induced apoptotic cell death. For this, we first performed Annexin V/PI staining to evaluate the effects of 3-MA on apigenin-induced apoptosis. Results indicated that apoptosis induction was 25% by apigenin, 23% by 3-MA, and 45.3% by the combination of the two agents (Fig. 5A). Furthermore, the alteration in the protein levels of LC3 and apoptotic proteins procaspase-8, -9 and -3 and PARP cleavage were also observed by western blot analysis. The protein level of LC3-II was increased by apigenin, however, addition of 3-MA abrogated the effect of apigenin on LC3-II induction (Fig. 5B). We also found that the combination of 3-MA and apigenin resulted in a significant increase in the level of PARP cleavage and decrease in procaspase-3, -8 and -9 compared to apigenin treatment alone (Fig. 5B). Altogether, these results indicated that inhibition of autophagy by 3-MA enhanced apigenin-induced cell death in HCT116 cells.