Journal of Cancer Prevention 2016; 21(4): 249-256
Published online December 30, 2016
© Korean Society of Cancer Prevention
You Na Oh1, Soojung Jin1,2, Hyun-Jin Park1, Hyun Ju Kwon1,2, and Byung Woo Kim1,2
1Blue-Bio Industry Regional Innovation Center, Dong-Eui University, Busan, Korea, 2Department of Life Science and Biotechnology, College of Natural Sciences and Human Ecology, Dong-Eui University, Busan, Korea
Correspondence to :
Byung Woo Kim, Blue-Bio Industry Regional Innovation Center, Dong-Eui University, 176 Eomgwang-ro, Busanjin-gu, Busan 47340, Korea, Tel: +82-51-890-2900, Fax: +82-505-182-6951, E-mail: email@example.com
This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (http://creativecommons.org/licenses/by-nc/4.0) which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited.
To examine the antioxidant activity of EESR, 2,2-diphenyl-1-picrylhydrazyl radical scavenging activity assay was performed. Inhibitory effect of EESR on cancer cell growth and proliferation was determined by water-soluble tetrazolium salt assay. To investigate the mechanism of EESR-mediated cytotoxicity, HT29 cells were treated with various concentrations of EESR and the induction of cell cycle arrest and apoptosis was analyzed by flow cytometry, 4,6-diamidino-2-phenylindole staining, and Western blot analysis. EESR showed significant antioxidant activity and inhibitory effect on HT29 cell growth in a dose-dependent manner. EESR induced cell cycle arrest at G2/M phase in a dose-dependent manner by modulating cyclin B, cyclin-dependent kinase 1 (CDK1), and CDK inhibitor p21 expression. EESR-induced apoptosis was associated with the upregulation of p53, a death receptor Fas, and a pro-apoptotic protein Bax and the activation of caspase 3, 8, and 9, resulting in the degradation of PARP. EESR possessing antioxidant activity efficiently inhibits proliferation of HT29 cells by inducing both cell cycle arrest and apoptosis. EESR may be a possible candidate for the anticancer drug development.
To examine the antioxidant activity of EESR, 2,2-diphenyl-1-picrylhydrazyl radical scavenging activity assay was performed. Inhibitory effect of EESR on cancer cell growth and proliferation was determined by water-soluble tetrazolium salt assay. To investigate the mechanism of EESR-mediated cytotoxicity, HT29 cells were treated with various concentrations of EESR and the induction of cell cycle arrest and apoptosis was analyzed by flow cytometry, 4,6-diamidino-2-phenylindole staining, and Western blot analysis.
EESR showed significant antioxidant activity and inhibitory effect on HT29 cell growth in a dose-dependent manner. EESR induced cell cycle arrest at G2/M phase in a dose-dependent manner by modulating cyclin B, cyclin-dependent kinase 1 (CDK1), and CDK inhibitor p21 expression. EESR-induced apoptosis was associated with the upregulation of p53, a death receptor Fas, and a pro-apoptotic protein Bax and the activation of caspase 3, 8, and 9, resulting in the degradation of PARP.
EESR possessing antioxidant activity efficiently inhibits proliferation of HT29 cells by inducing both cell cycle arrest and apoptosis. EESR may be a possible candidate for the anticancer drug development.
Keywords: Anticancer, Antioxidant, Apoptosis, Cell cycle arrest,
Colorectal cancer is the second most prevalent malignancy and the fourth leading cause of cancer death globally, with 694,000 mortalities in 2012.1 Colorectal cancer is the third most common cancer in men and the second most common in women, representing almost 10% of the global cancer incidence.1 Due to its difficulty in early diagnosis, strong invasion, metastasis, and poor prognosis, colorectal cancer is highly lethal and aggressively malignant.1,2 As the outcome of surgical resection, the main treatment for colorectal cancers, is not always satisfactory, chemopreventive and chemotherapeutic approaches of novel natural compounds and extracts have received increasing attention in recent years.3–5
The abnormal cell growth is one of the critical characteristics of cancer for invasion or spreading out to other parts of the body. Cancer cells can grow unusually via limitless number of cell division and can avoid apoptosis.6,7 Therefore, the inhibition of cancer cell proliferation by inducing cell cycle arrest and apoptosis is being an important preventive strategy against cancer.8,9 Cell cycle program is regulated by cyclin/cyclin-dependent kinase (CDK) complexes.10,11 The major molecules responsible for G2/M transition are cyclin A /CDK1 and cyclin B/CDK1.12 The activation of cyclin/CDK complex accelerates cell cycle progression, which are regulated by phosphorylation. Whereas damage signals of cells activate p53 via CHK2 to repair damaged DNA, leading to cell cycle arrest.13 The expression of CDK inhibitor p21 is upregulated by activated p53, resulting in the suppression of G2/M transition by the inactivation of cyclin/CDK complex.14,15
Apoptosis, a type of programmed cell death, is characterized by cell membrane blebbing, cell shrinkage, nuclear fragmentation, chromatin condensation, and chromosomal DNA fragmentation. 16,17 There are two basic apoptotic signaling pathways: the extrinsic and the intrinsic pathways.18,19 The extrinsic pathway can be mediated by one of several death receptors (TNF receptor superfamily) when bound by the appropriate ligand (e.g., Fas ligand, TNF-related apoptosis inducing ligand, and TNF-α).20 The interaction between death receptors and their ligands leads to the assembly of the death-inducing signaling complex, followed by the activation of an initiator caspase, caspase-8.21 The intrinsic apoptotic pathway is activated by various intracellular stimuli, including DNA damage, growth factor deprivation, and oxidative stress. It has been reported that the modulation of pro-apoptotic Bcl-2 family proteins can cause the mitochondrial membrane permeabilization, leading to the release of cytochrome
The ethanol extract of
Human colon adenocarcinoma HT29 cells, human hepatocellular carcinoma HepG2 cells, and human lung adenocarcinoma A549 cells were purchased from American Type Culture Collection (ATCC; Manassas, VA, USA). Cells were cultured in Dulbecco’s modified Eagle’s medium supplemented with 10% FBS, penicillin, and streptomycin at 37°C and 5% CO2.
Antioxidant activity of EESR was determined using a 2,2-diphenyl-1-picrylhydrazyl (DPPH) radical scavenging assay. Various concentrations (2.56–64 μg/mL) of EESR and standard compound (ascorbic acid) were added to DPPH solution. Absorbance was measured at 520 nm using a microplate reader (Beckman Coulter, Fullerton, CA, USA), after reacting for 30 minutes at room temperature. The assay was replicated three times and results are expressed as mean ± SD. The percentage of inhibition was assessed using the following formula:
A: sample absorbance 520 nm
B: color control absorbance 520 nm
C: control absorbance 520 nm
The concentration of sample inhibiting 50% of free radical DPPH (IC50) was determined.
The cell viability was measured using the EZ-Cytox cell viability assay kit (Daeillab, Seoul, Korea). Cells were plated at a density of 2–5 × 104 cells/mL in 24-well plate and treated with media containing DMSO as a control or various concentrations of EESR for 48 hours. EZ-Cytox assay reagent (10 μL) was added to each cell culture well, and the mixture was incubated for 30 minutes at 37°C. The absorbance was measured at 450 nm using a microplate reader. For morphological study, HT29 cells were treated with EESR for 48 hours and directly photographed with an inverted microscope using Axio Vision program.
The effect of EESR on the cell cycle in HT29 cells was examined using the MuseTM Cell Cycle kit (Merck Millipore, Darmstadt, Germany) according to the manufacturer’s instructions. Briefly, cells (2 × 105 cells/well) were plated in 6-well plate and treated with 0.1% DMSO as a vehicle control or with various concentrations of EESR for 48 hours. The cells were then harvested, washed once with PBS, and fixed in cold 70% ethanol for 3 hours at −20°C. Fixed cells were centrifuged at 300 ×
EESR-treated cells were lysed with lysis buffer (20 mM Tris-HCl [pH 7.5], 150 mM NaCl, 1 mM ethylenediaminetetraacetic acid, 1 mM ethylene glycol bis(2-aminoethyl ether) tetraacetic acid, 1% Triton X-100, 1 μg/mL leupeptin, 1 mM phenylmethanesulfonyl fluoride) for 1 hour at 4°C and centrifuged for 30 minutes at 13,000 rpm. Total soluble proteins in the supernatant were collected and the concentration of protein was determined by Bradford method. For Western blot analysis, 30 to 50 μg/mL of proteins were resolved by sodium dodecyl sulfate-polyacrylamide gel electrophoresis and blotted onto nitrocellulose membranes. Blots were incubated at 4°C overnight with specific primary antibodies followed by horseradish peroxidase-conjugated secondary antibodies and visualized by an enhanced chemiluminescence detection system (FluoChem®FC2; Alpha-Innotech, San Leandro, CA, USA) using Western blotting luminol reagent (Santa Cruz Biotechnology, Dallas, TX, USA). CDK1, cyclin A, cyclin B, cell division cycle 25C (Cdc25C), p-Cdc25C, Wee1, p53, Fas, Fas-associated protein with death domain (FADD), Bax, caspase-3, caspase-8, caspase-9, PARP, actin primary antibodies and peroxidase-conjugated secondary antibodies were from Santa Cruz Biotechnology. Primary antibodies against CHK2, p-CHK2, p21, p-CDK1, and Bcl-2 were purchased from Cell Signaling Technology (Beverly, CA, USA).
Cells (5 × 104 cells/well) were plated in 8-well chamber slide and treated with 0.1% DMSO as a vehicle control or with various concentrations of EESR for 48 hours. EESR-treated cells were fixed with 4% formaldehyde for 20 minutes at room temperature. Fixed cells were permeabilized with 0.5% Triton X-100 in PBS for 10 minutes at room temperature. After washing with PBS, cells were incubated with 1 μg/mL of 4,6-diamidino-2-phenylindole (DAPI) for 10 minutes and then washed three times in PBS. Apoptotic nuclei (condensed chromatin) were examined by fluorescence microscopy (Carl Zeiss, Jena, Germany).
Data are presented as the mean ± SD from at least three independent experiments. Statistical comparisons between groups were performed by SPSS program followed by Student
Since it has been reported that oxidative stress is one of the key components in the induction and the maintenance of cancer,27,28 we first examined the antioxidant effect of EESR using DPPH radical scavenging activity assay. As shown in Table 1, EESR inhibited DPPH activity in a dose-dependent manner showing 47.40%, 95.32% and 98.71% inhibition rate at 2.56 μg/mL, 12.80 μg/mL and 64.00 μg/mL, respectively. EESR showed the IC50 (concentration at 50% inhibition) value of 3.11 μg/mL, which were comparable to reference compound (ascorbic acid) with IC50 value of 1.49 μg/mL. These results indicate that EESR have significant antioxidant activity.
To investigate the anticancer activity of EESR, we examined the inhibitory effect of EESR on cancer cell growth using human colon adenocarcinoma HT29 cells, human hepatocellular carcinoma HepG2 cells, and human lung adenocarcinoma A549 cells. Cells were treated with various concentrations of EESR for 48 hours and cell viability was measured by water-soluble tetrazolium salt assay. As shown in Figure 1A, EESR inhibited the growth of HT29 cells in a dose-dependent manner, whereas EESR did not show any significant inhibitory effect on cell proliferation in HepG2 and A549 cells at the highest concentration (200 μg/mL). In addition, the morphological changes of EESR-treated HT29 cells were observed using a phase contrast microscope. Increasing concentrations of EESR caused inducing the morphological change of HT29 cells and decreasing viable adherent HT29 cells (Fig. 1B). Therefore, we decided to examine the molecular mechanism of cytotoxicity of EESR in HT29 cells.
To clarify the mechanisms involved in the cytotoxic effect of EESR in HT29 cells, cell cycle analysis was performed by flow cytometry. As shown in Figure 2, cell population at G2/M phase increased by EESR in a dose-dependent manner up to 100 μg/mL, accompanied by a decrease of cell population at G1 and S phases. Furthermore, EESR treatment induced significant accumulation of cells at SubG1 phase (apoptotic cell population) in a concentration-dependent manner, raising the possibility that EESR may induce apoptosis in HT29 cells. Figure 2 also shows the distribution of cells in various stages of the cell cycle. These results suggest that EESR may induce G2/M arrest and apoptosis in HT29 cells.
As we found that the cell population in G2/M phase increased in a dose-dependent manner by EESR, the expression of G2/M transition-related proteins was examined using Western blot analysis. As shown in Figure 3A, the expression of p53 and p21 increased, whereas the expression of cyclin B decreased significantly in EESR-treated HT29 cells. However, the expression of cyclin A did not show any big change after EESR treatment. In addition, EESR activated CHK2 and inactivated Cdc25C via phosphorylation (Fig. 3B). Figure 3B also showed that the expression of Wee1 kinase increased by EESR treatment. These results indicate that EESR modulates the checkpoint proteins and blocked the G2/M transition.
Since HT29 cell accumulation at SubG1 phase was detected by flow cytometry, we next observed the formation of apoptotic bodies, one of the hallmarks of apoptosis, in EESR-treated HT29 cells by DAPI staining. As shown in Figure 4A, the apoptotic bodies increased, whereas the number of cell decreased in a dose-dependent manner by EESR treatment. To investigate the molecular mechanisms involved in the apoptosis induction of EESR in HT29 cells, the expression of apoptosis-related molecules in the protein levels were determined by Western blot analysis. As shown in Figure 4B, exposure to EESR led to a significant increase of p53, a death receptor Fas, FADD, and Bax expression in HT29 cells. However, there was no big difference in Bcl-2 expression between vehicle control-treated and EESR-treated cells (Fig. 4B). As shown in Figure 4C, EESR activated caspases, including caspase-8, caspase-9, and caspase-3 in a dose-dependent manner, leading to the cleavage of PARP, one of the target proteins of caspase-3 in apoptosis.
In the present study, the antioxidant and anticancer activities of
G2/M transition is regulated by cyclin A/CDK1 and cyclin B/CDK1 complexes. In detail, cyclin A can associate with CDK1 in the late S phase and the cell will progress through the S phase to G2 phase.31 In the late G2 phase, cyclin A is replaced by cyclin B, leading to the formation of cyclin B/CDK1 complex, and cyclin B/CDK1 complex triggers the progression of the cells into and out of M phase.32 In this study, we found that EESR downregulated cyclin B expression, whereas expression of cyclin A did not show any big difference by EESR, suggesting that EESR may induce cell cycle arrest at the transition from G2 to M phase.
The negative regulation of the cyclin B/CDK1 complex occurs via p53-dependent and p53-independent pathways, resulting in cell cycle arrest.14 p53, a multifunctional tumor suppressor, is known to regulate various cellular events, such as cell cycle arrest, transcription, differentiation, and apoptosis.33 In the case of p53-dependent pathway, DNA damage signals trigger p53 upregulation, and then p53 induces the expression of p21, a CDK inhibitor, which can suppress G2/M transition by the inactivation of cyclin/CDK complex.34 In the p53-independent pathway, CHK1 and CHK2 can inhibit the function of Cdc25C phosphatase through phosphorylation, leading to increase of p-CDK1, an inactive form of CDK1.35,36 CDK1 is also negatively regulated by Wee1 kinase, which catalyzes the inhibitory phosphorylation of CDK1, resulting in G2/M arrest.37 Our data from Western blot analysis revealed that EESR induced the increase of p53 and p21 protein expression, and the inactivation of CDK1 via phosphorylation by inactivation of Cdc25C and upregulation of Wee1. These results suggest that EESR may induce G2/M arrest by both p53-dependent and p53-independent pathways.
It has been reported that p53 can activate the extrinsic pathway of apoptosis through the induction of Fas gene expression and the enhancement of cell surface levels of Fas.38,39 In addition, p53 is known to upregulate the expression of Bax, a pro-apoptotic Bcl-2 family, which governs the release of cytochrome
In conclusion, we found that EESR showed significant antioxidant activity and efficiently induced G2/M arrest and apoptosis in HT29 cells, followed by the suppression of HT29 cell proliferation. EESR-mediated cell cycle arrest at G2/M phase was associated with both p53-dependent and p53-independent pathways. In addition, EESR induced HT29 cell apoptosis associated with p53 upregulation through both extrinsic and intrinsic pathways. Our findings firstly show the antioxidant and anticancer activities of EESR and the mode of action, suggesting that EESR may be a possible nutraceutical candidate for chemoprevention and chemotherapy of cancer. Further studies will be needed to identify the bioactive compound(s) of EESR for its anticancer activity.
This work was supported by Blue-Bio Industry Regional Innovation Center (RIC08-06-07) at Dong-Eui University as a RIC program under Ministry of Trade, Industry and Energy (MOTIE) and Busan city.
DPPH radical scavenging activity by EESR.
|Concentration||Inhibition rate||Concentration||Inhibition rate|
|(μg/mL)||(%), mean ± SD||(μg/mL)||(%), mean ± SD|
|2.56||47.40 ± 0.63||0.51||22.70 ± 0.94|
|12.80||95.32 ± 0.13||2.56||79.66 ± 0.79|
|64.00||98.71 ± 0.25||12.80||97.57 ± 0.21|
IC50 (μg/mL): EESR, 3.11; ascorbic acid, 1.49. DPPH, 2,2-diphenyl-1-picrylhydrazyl; EESR, ethanol extract of
Soojung Jin, Hyun-Jin Park, You Na Oh, Hyun Ju Kwon, Jeong-Hwan Kim, Yung Hyun Choi, and Byung Woo KimJournal of Cancer Prevention 2015; 20(4): 241-249 https://doi.org/10.15430/JCP.2015.20.4.241
Ji-Young Hong, Hwa-Jin Chung, Song Yi Bae, Trinh Nam Trung, KiHwan Bae, and Sang Kook LeeJournal of Cancer Prevention 2014; 19(4): 273-278 https://doi.org/10.15430/JCP.2014.19.4.273
Seong-Joon Kim1, Yung-Hyun Choi2, Won-Ho Lee3
Eun-Young Kwon and Kun-Young Park