Sage Journals: Discover world-class research

Abstract

Tomentosin, a sesquiterpene lactone, is known to possess various biological activities. However, its anticarcinogenic activity against human hepatocellular carcinoma (HCC) cells has not been investigated in detail. Thus, this study aimed to elucidate the cytotoxic mechanism of tomentosin in human HCC cell lines HepG2 and Huh7. WST-1, cell counting, and colony formation assay results showed that treatment with tomentosin decreased the viability and suppressed the proliferation rate of HepG2 and Huh7 cells in a dose- and time-dependent manner. Cell cycle analysis revealed increased population of cells at the SubG1 and G2/M stage, and decreased population of cells at the G0/1 stage in HepG2 and Huh7 cells treated with tomentosin. Annexin V/propidium iodide double staining and TUNEL assay results showed increased apoptotic cell population and DNA fragmentation in HepG2 and Huh7 cells treated with tomentosin. Western blotting analysis results showed that tomentosin treatment significantly increased the expression level of Bax, Bim (short form), cleaved PARP1, FOXO3, p53, pSer15p53, pSer20p53, pSer46p53, p21, and p27, but decreased the expression of Bcl2, caspase3, caspase7, caspase9, cyclin-dependent kinase 2 (CDK2), CDK4, CDK6, cyclinB1, cyclinD1, cyclinD2, cyclinD3, and cyclinE in a dose-dependent manner. Taken together, this study revealed that tomentosin, which acted through cell cycle arrest and apoptosis, may be a useful therapeutic option against HCC.

Keywords

Tomentosin hepatocellular carcinoma FOXO3 p53 cell cycle arrest apoptosis

Introduction

Human hepatocellular carcinoma (HCC) is a malignancy arising from hepatocytes in the liver. HCC is the major cause of death in males worldwide.¹ In addition, its incidence and mortality rates have been rising in Eastern Asia, Southeastern Asia, and parts of Africa.² Moreover, the incidence of HCC has been rapidly increasing in Western countries.³ Several therapies, such as radiation and chemotherapy, are effective against cancer cells, but still deleterious to normal liver cells.⁴ Thus, the development of efficient therapeutic options against HCC without side effects is urgently needed.

Natural compounds originating from plants have been used to treat many diseases in humans.⁵ Biologically active phytochemicals are known to have anticancer, anti-inflammatory, anti-allergic, and antioxidant activities. Recently, numerous studies have showed the way for improving the human health with flavonoids and particularly the way to treat many cancer types and cardiovascular disease.^6
–8 Tomentosin, a kind of sesquiterpene lactones, is known to have a variety of biological activities. It can function as anti-inflammatory, antioxidant, and anticancer reagent.^9

–13 In addition, tomentosin has the activity for hypoglycemic, hypolipidemic, antibacterial, antifungal,^14,15 and inhibition of telomerase activity.¹⁶ However, there are no detailed studies on the potential anticancer activity of tomentosin in HCC cells. Therefore, in this study, we aimed to present results on the cytotoxic effect of tomentosin.

Cell cycle arrest in the G0/1 or G2/M cell division checkpoint phase occurs when cells are under stressful conditions, such as DNA damage.¹⁷ The G0/1 phase cell cycle arrest is caused by inhibition of cyclin and the cyclin-dependent kinase (CDK) complex by p21, an endogenous CDK inhibitor that transcriptionally regulates p53.^18,19 Likewise, the G2/M phase cell cycle arrest is activated by p53, which is responsible for the induction of p21, leading to blockade of DNA replication.²⁰ Apoptosis is a death process that occurs in damaged, abnormal, and aged cells, and its mechanism involves various complex signaling pathways.^21,22 For example, apoptosis causes cell morphology changes and DNA fragmentation.²³ The typical p53 and caspase pathways are also involved in apoptosis.^21,24 Both cell cycle arrest and apoptosis in cancer cells are the most targeted cellular process for the therapy of tumor patients. In this study, we examined the potential cytotoxic effect of tomentosin via induction of cell cycle arrest and apoptosis in HepG2 and Huh7 cells.

Materials and methods

Reagent

Tomentosin was purchased from EnsolBio (Daejeon, Korea). It was dissolved in DMSO, and 40 mM tomentosin stock solution was stored at −20°C. Primary antibodies against PARP1, cleaved PARP1, caspase3, caspase7, caspase9, Bax, Bim, Bcl2, p27, p21, FOXO3, p53, pSer15p53, pSer20p53, pSer46p53, Akt, pAkt, Erk, pErk, p38, pp38, Jnk, pJnk, CDK2, CDK4, CDK6, cyclinB1, cyclinD1, cyclinD2, cyclinD3, cyclinE, and β-actin were purchased from Cell Signaling Technology (Beverly, Massachusetts, USA). HRP-conjugated goat anti-mouse and anti-rabbit immunoglobulin G (IgG) secondary antibodies were purchased from Jackson ImmunoResearch (West Grove, Pennsylvania, USA).

Cell culture

HepG2 and Huh7 cells were purchased from ATCC (Manassas, Virginia, USA) and cultured in the recommended medium (Dulbecco’s modified Eagle’s medium) in a cell culture incubator at 37°C, 5% CO₂, and 95% humidity. Dulbecco’s modified Eagle’s medium was purchased from GIBCO (Grand Island, New York, USA). The medium contains 10% heat-inactivated fetal bovine serum and 1% of penicillin/streptomycin antibiotics (GIBCO).

WST-1 assay

HepG2 and Huh7 (4 × 103) cells were plated in six-well plates. After 24 h of culture, different concentrations of tomentosin (0, 10, 20, and 40 μM) were added, and the cells were cultured for another 24 or 48 h. After that, 15 µL of cell proliferation reagent (EZ-CYTOX; DoGenBio, Seoul, Korea) was added, and the cells were then placed in an incubator under standard conditions for 45 min. The absorbance level was measured at 450 nm using a microplate reader.

Colony formation assay

HepG2 and Huh7 cells (500 cells) were plated in six-well plates. After 24 h of culture, the cells were treated with tomentosin (0, 10, and 20 μM) for 24 h in an incubator. The media were then replaced with fresh media, and the cells were cultured for 2 weeks in an incubator. After incubation, the cells were rinsed with PBS and stained with freshly prepared 1% crystal violet solution for 25 min. After staining, the number of colonies was counted.

Cell counting assay

HepG2 and Huh7 (2 × 10⁴) cells were plated in six-well plates and cultured in an incubator for 24 h. The cells were treated with tomentosin (0, 10, and 20 μM) for 24, 48, or 72 h. After that, the number of cells was counted with a hemocytometer.

TUNEL assay

Tomentosin-induced double-stranded cleavage of DNA in HepG2 and Huh7 cells was visualized using an assay kit (Promega, Madison, Wisconsin, USA) following the manufacturer’s instructions. HepG2 and Huh7 (2 × 10⁵) cells were plated in six-well plates. After 24 h, the cells were treated with tomentosin (0, 10, 20, and 40 μM) for 24 or 48 h. The cells were then rinsed twice with PBS and fixed with 4% formaldehyde for 20 min at room temperature. After being washed with PBS, the cells were incubated in 0.5% Triton X-100 for 5 min at room temperature. After that, the cells were stained with 50 µL of TUNEL reaction mixture in an incubator for 1 h. The cells were counterstained with Hoechst staining solution (Sigma-Aldrich, St. Louis, MO, USA) for 10 min for visualization of the nuclei. The labeled cells were observed with a fluorescence microscope (Eclipse TE 2000-U; Nikon, Tokyo, Japan).

Annexin V/propidium iodide double staining assay

Apoptotic cell rates were measured by using an FITC Annexin V apoptosis detection kit (BD Biosciences, Franklin Lakes, New Jersey, USA). HepG2 and Huh7 (2.5 × 10⁵) cells were seeded in 6-cm dishes and cultured for 24 h. The cells were treated with tomentosin (0, 10, 20, and 40 μM) for 24 or 48 h, rinsed twice with PBS, resuspended in 1× binding buffer, and stained with FITC Annexin V in darkness for 15 min at room temperature. Apoptotic cell rates were measured by flow cytometry (Beckman Coulter, Brea, California, USA).

Cell cycle analysis

HepG2 and Huh7 (2.5 × 10⁵) cells were seeded in 6-cm dishes and incubated for 24 h. The cells were treated with tomentosin (0, 10, 20, and 40 μM) for 24 or 48 h. The cells were washed twice with PBS and resuspended in 70% EtOH for 24 h at −20°C. Next, the cells were rinsed twice with PBS and resuspended in propidium iodide (PI) (Sigma-Aldrich) buffer in darkness for 15 min at room temperature. Stained cells were analyzed by flow cytometry (Beckman Coulter).

Western blotting analysis

HepG2 and Huh7 (5 × 10⁵) cells were plated in 10-cm culture dishes and incubated. After 24 h, the cells were treated with tomentosin (0, 10, 20, and 40 μM) for 48 h. Total protein lysate was isolated using RIPA buffer (Sigma-Aldrich) containing protease inhibitors (Sigma-Aldrich). Protein concentration was determined using a Qubit™ Fluorocytometer (Invitrogen, Carlsbad, California, USA). The same amount of proteins was separated through SDS-PAGE for 2 h at 110 V. Proteins were transferred to an Immobilon-P PVDF membrane (Merck Millipore, Burlington, Massachusetts, USA) for 2 h at 50 V. The blots were blocked using 3% BSA (Bovogen, Australia) buffer at room temperature for 1 h. Membranes were incubated with primary antibodies at 4°C overnight, washed three times with TBST for 15 min, and finally incubated with anti-mouse or anti-rabbit IgG antibodies (1:10,000 dilution) in 3% BSA buffer for 1 h at room temperature. The fluorescent blots were visualized with an ECL kit (GenDEPOT, Barker, Texas, USA) and a ChemiDoc detection system (Bio-Rad, Hercules, California, USA).

Statistical analysis

All data were analyzed with one-way analysis of variance using the Microsoft Excel software. The value of p < 0.05 was considered to indicate statistical significance.

Results

Tomentosin inhibits the growth of HepG2 and Huh7 cells

To evaluate the antiproliferative effects of tomentosin, HepG2 and Huh7 cells were treated with 0, 10, 20, and 40 µM of tomentosin for 24 and 48 h, and WST-1 assay was conducted. Compared with the DMSO (vehicle)-treated control, HepG2 cells treated for 24 h with tomentosin (10, 20, and 40 µM) showed decreased viability (89.1%, 70.3%, and 55.4%, respectively), and Huh7 cells treated for 24 h with tomentosin (10, 20, and 40 µM) showed decreased viability (89.5%, 74.6%, and 59.8%, respectively) (Figure 1(a)). Compared with the DMSO-treated control, HepG2 cells treated for 48 h with tomentosin (10, 20, and 40 µM) showed decreased viability (87.1%, 59.6%, and 34.7%, respectively), and Huh7 cells treated for 48 h with tomentosin (10, 20, and 40 µM) showed decreased viability (76.7%, 54.0%, and 25.3%, respectively) (Figure 1(a)). Furthermore, cell counting assay was conducted to investigate the antiproliferative effects of tomentosin. The results indicated that compared with that in the DMSO control group, the number of cells decreased in a dose-dependent manner in HepG2 and Huh7 cells treated with tomentosin (Figure 1(b)). Colony formation assay was also conducted to visualize the antiproliferative effects of tomentosin in HepG2 and Huh7 cells. The images showed that tomentosin decreased the number of HepG2 and Huh7 cell colonies. Thus, our results suggested that tomentosin inhibited cell growth in HepG2 and Huh7 cells.

Figure 1.

Cell viability and proliferation of tomentosin-treated HepG2 and Huh7 cells. (a) Cells were treated with different concentrations (0, 10, 20, and 40 µM) of tomentosin for 24 and 48 h. Cell viability was determined by WST-1 assay. Error bar represents the mean ± SEM. *p < 0.05; **p < 0.01. These results are representative data from three biological replicates. (b) Cells were treated with different concentrations (0, 10, and 20 µM) of tomentosin for 24, 48, and 72 h. Cell survival was determined by cell counting assay. Error bar represents the mean ± SEM. *p < 0.05; **p < 0.01. These results are representative data from three biological replicates. (c) Effect of tomentosin on colony formation of cells. Cells were treated with different concentrations (0, 10, and 20 µM) of tomentosin for 24 h and cultured for 14 days. The number of colonies was measured by using a number counter. Error bar represents the mean ± SEM. *p < 0.05; **p < 0.01. These results are representative data from three biological replicates. SEM: standard error of the mean.

Tomentosin induces cell cycle arrest and apoptosis in HepG2 and Huh7 cells

We investigated the effects of 24- and 48-h treatment with tomentosin (0, 10, 20, and 40 µM) on cell cycle arrest in HepG2 and Huh7 cells. DNA in each cell cycle stage was analyzed by PI staining and analyzed by FACS. The results indicated that tomentosin treatment for 24 h induced the G2/M phase cell cycle arrest in HepG2 and Huh7 cells (Figure 2(a) and (b)). The G2/M population increased from 34.7% in the control to 43.9% in HepG2 cells treated with 40 μM tomentosin for 24 h, and from 27.2% in the control to 51.8% in Huh7 cells treated with 40 μM tomentosin for 24 h (Figure 2(c) and (d)). The G2/M population increased from 23.5% in the control to 33.9% in HepG2 cells treated with 40 μM tomentosin for 48 h, and from 31.2% in the control to 33.4% in Huh7 cells treated with 40 μM tomentosin for 48 h. Interestingly, the SubG1 population also increased in HepG2 and Huh7 cells after treatment with tomentosin for 48 h (Figure 2(a) and (b)). The SubG1 population increased from 2.8% in the control to 11.6% in HepG2 cells treated with 40 μM tomentosin for 48 h, and from 3.9% in the control to 15.4% in Huh7 cells treated with 40 μM tomentosin for 48 h (Figure 2(c) and (d)). Thus, these results implied that tomentosin treatment (40 µM) for 24 and 48 h induced cell cycle arrest and apoptosis, respectively, in HepG2 and Huh7 cells.

Figure 2.

Cell cycle population in tomentosin-treated HepG2 and Huh7 cells. Cells were treated with different concentrations (0, 10, 20, and 40 µM) of tomentosin for 24 and 48 h. Cells were stained with PI and analyzed by flow cytometry (SubG1, G0/1, S, and G2/M). The percentage of cell cycle population of HepG2 (a) and Huh7 (b) cells was indicated in the cytogram. The cell cycle population of HepG2 (c) and Huh7 (d) was graphically analyzed. PI: propidium iodide.

Tomentosin increases apoptosis in HepG2 and Huh7 cells

Annexin V/PI double staining and TUNEL assay were conducted to evaluate the effects of 24- and 48-h treatment with tomentosin (0, 10, 20, and 40 μM) on apoptosis in HepG2 and Huh7 cells. As shown in Figure 3(a) and (b), Annexin V/PI double staining assay results showed that the proportion of early and late apoptotic cells increased in HepG2 and Huh7 cells. After treatment for 24 h, the proportion of apoptotic cells (early + late) increased from 12.2% in the control to 10.7%, 13.3%, and 25.5% in HepG2 cells treated with 10, 20, and 40 μM tomentosin, and from 5.2% in the control to 7.8%, 11.7%, and 23.0% in Huh7 cells treated with 10, 20, and 40 μM tomentosin, respectively. After 48 h of incubation, the population of apoptotic cells (early + late) increased from 4.9% in the control to 27.6%, 47.8%, and 93.4% in HepG2 cells treated with 10, 20, and 40 μM tomentosin, and from 4.3% in the control to 7.5%, 21.0%, and 29.0% in Huh7 cells treated with 10, 20, and 40 μM tomentosin, respectively (Figure 3(c) and (d)). As shown in Figure 4(a) and (b), TUNEL assay results showed that tomentosin treatment increased the proportion of apoptotic cells with fragmented DNA in a concentration-dependent manner. Thus, consistent with cell cycle analysis results, these findings suggested that tomentosin promoted apoptosis in HepG2 and Huh7 cells.

Figure 3.

Apoptotic cell population among HepG2 and Huh7 cells treated with tomentosin (0, 10, 20, and 40 µM) for 24 and 48 h. Apoptotic cell rates were analyzed by Annexin V/PI double staining assay. (B1) Necrotic cells; (B2) late apoptotic cells; (B3) viable cells; (B4) early apoptotic cells. The percentage of cell cycle population of HepG2 (a) and Huh7 (b) cells was indicated in each box. The percentages of apoptotic cells of HepG2 (c) and Huh7 (d) were graphically analyzed. PI: propidium iodide.

Figure 4.

DNA fragmentation in tomentosin-treated HepG2 and Huh7 cells. Cells were treated with different concentrations (0, 10, 20, and 40 µM) of tomentosin for 24 and 48 h. Cell apoptosis was detected by TUNEL assay. Blue and green indicate nuclei and fragmented DNA, respectively, in HepG2 (a) and Huh7 (b) cells. The number of cells with fragmented DNA among HepG2 (c) and Huh7 (d) cells was graphically analyzed.

Tomentosin activates cell cycle arrest and apoptotic signaling pathway in HepG2 and Huh7 cells

Western blotting was performed to elucidate the intracellular signaling pathways associated with the cell cycle arrest and apoptosis induced by tomentosin in HepG2 and Huh7 cells. First, we measured the expression level of cell cycle-associated proteins, such as p21, p27, CDK2, CDK4, CDK6, cyclinB1, cyclinD1, cyclinD2, cyclinD3, and cyclinE in HepG2 and Huh7 cells after treatment with tomentosin (0, 10, 20, and 40 μM) for 48 h (Figure 5). We observed that the expression level of p21 and p27 increased, whereas that of CDK2, CDK4, CDK6, cyclinB1, cyclinD1, cyclinD2, cyclinD3, and cyclinE decreased in a dose-dependent manner, which suggested that tomentosin induced cell cycle arrest in HepG2 and Huh7 cells through regulation of CDKs, cyclins, and CDK inhibitors. Next, we also measured the expression levels of apoptosis-associated proteins, such as caspase3, caspase7, caspase9, PARP1, cleaved PARP1, Bcl2, Bim (short form), Bax, FOXO3, p53, pSer15p53, pSer20p53, and pSer46p53 in the cells under the same condition (Figure 5). We observed that the expression levels of caspase3, caspase7, caspase9, PAPR1, and Bcl2 decreased, whereas those of cleaved PARP, Bax, Bim (short form), FOXO3, p53, pSer15p53, pSer20p53, and pSer46p53 increased after tomentosin treatment. These results implied that tomentosin promoted apoptosis in HepG2 and Huh7 cells through activation of the caspase-mediated intrinsic apoptosis pathway and tumor suppressive transcriptional factors. Furthermore, we analyzed the level of Akt, Erk, p38, Jnk, phosphorylated (p) Akt, pErk, pp38, and pJnk. We observed that the expression level of pAkt decreased, whereas that of pErk increased in a dose-dependent manner. Overall, these results indicated that tomentosin activated the intracellular signaling pathways of cell cycle arrest and apoptosis in HepG2 and Huh7 cells.

Figure 5.

Cell cycle arrest and expression of apoptosis-related proteins in tomentosin-treated HepG2 and Huh7 cells. Cells were treated with different concentrations (0, 10, 20, and 40 µM) of tomentosin for 48 h. Protein expression in HepG2 (a) and Huh7 (b) cells was analyzed by Western blotting using the indicated antibodies. β-actin was used as a gel-loading control. These results are representative data from three biological replicates.

Discussion

Liver cancer is the most common type of cancer, and it represents approximately 15% of all deaths in Asia and Africa.²⁵ In addition, HCC is the top six major cancer type worldwide and the second leading cause of death, especially in males.¹ Although surgical and conventional treatments, such as chemo- and radiation therapy, are available for HCC patients, they still have unfavorable outcomes.³ Therefore, many researchers have tried to find alternatives to conventional therapies against HCC. Thus, we aimed to identify a novel therapy for HCC using natural compounds derived from medicinal plants.

Tomentosin is a phytochemical sesquiterpene lactone mainly found in medicinal herbs, such as Arnica longifolia, Inula japonica, Inula viscosa, and Xanthium strumarium.^26

–29 Tomentosin is known to exert anticancer activity in several types of cancer cells.¹⁶ According to a recent report, tomentosin induces apoptosis and increases the SubG0 and G2/M population in human melanoma cell lines.¹³ Furthermore, our laboratory reported that tomentosin induces apoptosis in osteosarcoma cancer cells by generating reactive oxygen species.³⁰ However, the anticarcinogenic effects of tomentosin in HCC have not been studied well.

In this study, we showed that tomentosin treatment promoted programmed cell death and inhibited cell cycle progression in HepG2 and Huh7 cells. The viability of HepG2 and Huh7 treated with 10 and 20 µM treatment of tomentosin was almost same at 24 and 48 h incubation (Figure 1(a)). We thought that 10 and 20 µM treatment of tomentosin had the similar effect in suppressing cell viability and seemed to be under IC50 value. However, 40 µM treatment of tomentosin caused the reduction of cell viability in both cells with time-dependent manner and could reduce the cell viability under 50% at 48 h incubation in both cells. We observed that relatively low doses of tomentosin (10 and 20 μM) induced cell cycle arrest and apoptosis at limited rates in HepG2 and Huh7 cells. On the contrary, compared with the other concentrations, 40 μM tomentosin significantly increased the ratio of TUNEL-positive cells and population of apoptotic cells. Thus, our results suggested that different concentrations of tomentosin exerted different cytotoxic effects on HepG2 and Huh7 cells.

Caspases initiate apoptosis processes, including DNA fragmentation and membrane blebbing.^31,32 Caspases contribute in the extrinsic and intrinsic apoptosis pathways³³. The extrinsic apoptotic pathway is promoted by Fas ligand, TNF receptor, and TRAIL receptors to induce the activation of caspase8.³⁴ Caspase9 is an important mediator that initiates the intrinsic apoptotic pathway, which is mediated by mitochondrial outer membrane permeabilization.³⁵ Activated (cleaved) caspase9 is known to mediate the release of cytochrome c from the mitochondria into the cytoplasm and formation of apoptosomes, leading to the activation of caspase3 and caspase7.^36
–38 As shown in Figure 5, we confirmed the decreased expression of caspase3, caspase7, and caspase9, suggesting that tomentosin modulated the intrinsic apoptosis pathway in HepG2 and Huh7 cells.

PI3K/Akt is a well-known signaling pathway in cancer development. For example, the PI3K/Akt pathway promotes cancer cell survival by phosphorylating FOXO3 transcription factors, thereby accelerating the protein degradation of FOXO3 in cancer cells.³⁹ As shown in Figure 5, Western blotting results indicated that tomentosin downregulated the expression level of pAkt, whereas the expression level of FOXO3 was significantly upregulated by tomentosin. Thus, we concluded that the half-life of FOXO3 protein was increased through inhibition of Akt activity by tomentosin treatment in HepG2 and Huh7 cells.

FOXO proteins are known to play an important role in the physiological and pathological conditions of cancer.⁴⁰ FOXO3, a member of the FOXO family, is responsible for transcriptional regulation in cell proliferation and death.⁴¹ In addition, FOXO3 controls cell cycle arrest through transcriptional upregulation of p27 endogenous CDK2 inhibitor⁴² and induces the G2/M phase cell cycle arrest in breast cancer cells.⁴³ As shown in Figure 2, tomentosin induced the G2/M phase cell cycle arrest in HepG2 and Huh7 cells. Thus, consistent with the results of previous studies, our results suggested that tomentosin enhanced FOXO3 and p27 expression, thus inducing the G2/M phase cell cycle arrest in HepG2 and Huh7 cells. Furthermore, cell cycle analysis showed increased SubG1 cell population in HepG2 and Huh7 cells after tomentosin treatment. TUNEL and Annexin V/PI double staining assay results indicated that apoptotic HepG2 and Huh7 cells increased following tomentosin treatment. Thus, these results suggested that the increased apoptotic cell population was associated with the enhanced expression of FOXO3 and Bim in HepG2 and Huh7 cells treated with tomentosin.

p53 is a hallmark tumor suppressive transcriptional factor.⁴⁴ For example, the lack of the p53 gene in mice leads to higher ratio of tumor development than that in normal mice.⁴⁵ p53 promotes cell cycle arrest through phosphorylation of the serine 15 and 20 sites, and induces apoptosis via phosphorylation of the serine 46 site.^46

–49 p53 controls cell cycle arrest via activation of p21.⁵⁰ p53 is involved in mitochondrial apoptosis through induction of Bax and PUMA.^51,52 Indeed, we observed that tomentosin increased the expression of p53, pSer15p53, pSer20p53, pSer46p53, p21, and Bax in HepG2 and Huh7 cells (Figure 5). Therefore, specific phosphorylation of p53 might be involved in the cell cycle arrest and apoptosis of HepG2 and Huh7 cells after tomentosin treatment. Although Huh7 cells are known to have the mutant form of p53, p53 and its phosphorylated forms in Huh7 cells were upregulated by tomentosin treatment. Thus, the p53 mutant in Huh7 cells might not be related to the specific phosphorylation site (Ser15, Ser20, and Ser46).

Both FOXO3 and p53 are tumor suppressive transcriptional factors responsible for the regulation of p21 and p27 endogenous cell cycle dependent kinases inhibitors to induce cell cycle arrest. p21 is involved in the G1 cell cycle arrest, and it interferes with the cyclinE/CDK2 and cyclinD/CDK4 complex.⁵³ Similarly, p27 is involved in the G1 cell cycle arrest, and it inhibits the cyclinA/CDK2 complex.^54,55 As shown in Figure 5, Western blotting results showed that tomentosin upregulated the protein expression of p21 and p27 in HepG2 and Huh7 cells, which could be transcriptionally regulated via activation of FOXO3 and p53. Furthermore, tomentosin downregulated the protein expression of CDK2, CDK4, CDK6, cyclinB1, cyclinD1, cyclinD2, cyclinD3, and cyclinE in HepG2 and Huh7 cells. We thought that tomentosin might inhibit the cyclins/CDKs complex by inducing the protein degradation of cyclins and CDKs. Our next research will examine the half-life of cyclins and CDKs, and investigate whether their protein stability is regulated by proteasomal degradation after tomentosin treatment in HepG2 and Huh cells.

We investigated the expression level of mitogen-activated protein kinases, which modulate a wide range of physiological processes in HepG2 and Huh7 cells treated by tomentosin. Jnk and p38 are the fundamental mediators of cell proliferation, differentiation, migration, and survival.⁵⁶ Jnk is an important modulator of mitochondrial apoptotic proteins.⁵⁷ As shown in Figure 5, the phosphorylation of both Jnk and p38 was not changed, which indicated that the tomentosin-induced cell cycle regulation and programmed cell death processes in HepG2 and Huh7 cells were independent of Jnk and p38. Erk is known to participate in cell differentiation⁵⁸ and activate an extrinsic or an intrinsic apoptotic pathway.⁵⁹ As shown in Figure 5, pErk expression was dramatically increased in HepG2 and Huh7 cells treated with tomentosin. These results suggested that the Erk signaling pathway may be involved in the effect of tomentosin on cell cycle and cell survival in HepG2 and Huh7 cells.

In this study, we tested whether tomentosin exerts cytotoxic activity against HCC, HepG2, and Huh7 cells, and investigated its effects on cell division cycle and apoptosis in HepG2 and Huh7 cells. We showed that the cytotoxic activity of tomentosin in HepG2 and Huh7 cells was related to inhibition of Akt and activation of FOXO3, p53, and Erk. Further studies are still required to elucidate the underlying mechanism of tomentosin in animal experiments. We hope that this study advances the development of a novel alternative therapeutic option against HCC.

Footnotes

Acknowledgements

The authors deeply appreciate the technical assistance from their laboratory members.

Author contributions

SHY, CML, SHH, JL, KYJ, and SHP conceived the presented idea, carried out the experiments, and wrote the manuscript. SHY, CML, SHH, JL, KYJ, and SHP contributed to data analysis and interpretation. SHY, CML, SHH, JL, KYJ, and SHP provided critical feedback, discussed the results, and contributed to writing the first and final drafts of the manuscript. SHP sponsored the article processing charge. SHY, CML, and SHH contributed equally to this work.

Declaration of conflicting interests

The author(s) declared no potential conflicts of interest with respect to the research, authorship, and/or publication of this article.

Funding

The author(s) disclosed receipt of the following financial support for the research, authorship, and/or publication of this article: This research was supported by the Basic Science Research Program (Grant No. NRF-2014R1A6A3A04054307) and the Medical Research Center Program (Grant No. NRF-2017R1A5A2015061) through the National Research Foundation of Korea (NRF) funded by the Ministry of Science and ICT (MSIP).

ORCID iD

SH Park

References

Torre

Bray

Siegel

, et al. Global cancer statistics, 2012. CA Cancer J Clin 2015; 65: 87–108.

Torre

Siegel

Ward

, et al. Global cancer incidence and mortality rates and trends—an update. Cancer Epidemiol Biomarkers Prev 2016; 25: 16–27.

Kudo

. Surveillance, diagnosis, treatment, and outcome of liver cancer in Japan. Liver Cancer 2015; 4: 39–50.

Abate

Patel

Rauscher

, et al. Redox regulation of fos and jun DNA-binding activity in vitro. Science 1990; 249: 1157–1161.

Nahata

. Anticancer agents: a review of relevant information on important herbal drugs. Int J Clin Pharmacol Toxicol 2017; 6: 250–255.

Lagiou

Samoli

Lagiou

, et al. Intake of specific flavonoid classes and coronary heart disease—a case-control study in Greece. Eur J Clin Nutr 2004; 58: 1643–1648.

Mursu

Voutilainen

Nurmi

, et al. Flavonoid intake and the risk of ischaemic stroke and CVD mortality in middle-aged Finnish men: the Kuopio Ischaemic Heart Disease Risk Factor Study. Br J Nutr 2008; 100: 890–895.

Nahata

Saxena

Suri

, et al. Sphaeranthus indicus induces apoptosis through mitochondrial-dependent pathway in HL-60 cells and exerts cytotoxic potential on several human cancer cell lines. Integr Cancer Ther 2013; 12: 236–247.

Hernandez

Recio

Manez

, et al. Effects of naturally occurring dihydroflavonols from Inula viscosa on inflammation and enzymes involved in the arachidonic acid metabolism. Life Sci 2007; 81: 480–488.

10.

Park

Kim

, et al. Suppressive effect of tomentosin on the production of inflammatory mediators in RAW264.7 cells. Biol Pharm Bull 2014; 37: 1177–1183.

11.

Weng

Gao

, et al. Antiproliferative and apoptotic sesquiterpene lactones from Carpesium faberi. Bioorg Med Chem Lett 2011; 21: 366–372.

12.

Talib

Mahasneh

. Antiproliferative activity of plant extracts used against cancer in traditional medicine. Sci Pharm 2010; 78: 33–45.

13.

Rozenblat

Grossman

Bergman

, et al. Induction of G2/M arrest and apoptosis by sesquiterpene lactones in human melanoma cell lines. Bio Pharm 2008; 75: 369–382.

14.

Zeggwagh

Ouahidi

Lemhadri

, et al. Study of hypoglycaemic and hypolipidemic effects of Inula viscosa L. aqueous extract in normal and diabetic rats. J Ethnopharmacol 2006; 108: 223–227.

15.

Maoz

Neeman

. Antimicrobial effects of aqueous plant extracts on the fungi Microsporum canis and Trichophyton rubrum and on three bacterial species. Lett Appl Microbiol 1998; 26: 61–63.

16.

Merghoub

El Btaouri

Benbacer

, et al. Tomentosin induces telomere shortening and caspase-dependant apoptosis in cervical cancer cells. J Cell Biochem 2017; 118: 1689–1698.

17.

Maity

McKenna

Muschel

. The molecular basis for cell cycle delays following ionizing radiation: a review. Radiother Oncol 1994; 31: 1–13.

18.

Deng

Zhang

Harper

, et al. Mice lacking p21^CIP1/WAF1 undergo normal development, but are defective in G1 checkpoint control. Cell 1995; 82: 675–684.

19.

Brugarolas

Chandrasekaran

Gordon

, et al. Radiation-induced cell cycle arrest compromised by p21 deficiency. Nature 1995; 377: 552–557.

20.

Cox

Lane

. Tumour suppressors, kinases and clamps: how p53 regulates the cell cycle in response to DNA damage. Bioessays 1995; 17: 501–508.

21.

Elmore

. Apoptosis: a review of programmed cell death. Toxicol Pathol 2007; 35: 495–516.

22.

Igney

Krammer

. Death and anti-death: tumour resistance to apoptosis. Nat Rev Cancer 2002; 2: 277–288.

23.

Zhang

. DNA fragmentation in apoptosis. Cell Res 2000; 10: 205–211.

24.

Pistritto

Trisciuoglio

Ceci

, et al. Apoptosis as anticancer mechanism: function and dysfunction of its modulators and targeted therapeutic strategies. Aging (Albany NY) 2016; 8: 603–619.

25.

Kim

Jeon

, et al. Fisetin induces apoptosis in Huh-7 cells via downregulation of BIRC8 and Bcl2L2. Food Chem Toxicol 2010; 48: 2259–2264.

26.

Passreiter

De Carlo

Steigel

. Sesquiterpene lactone glycosides from Arnica longifolia. Planta Med 1999; 65: 153–156.

27.

Piao

Kim

Zhang

, et al. DNA topoisomerase inhibitory activity of constituents from the flowers of Inula japonica. Chem Pharm Bull (Tokyo) 2016; 64: 276–281.

28.

Fontana

La Rocca

Passannanti

, et al. Sesquiterpene compounds from Inula viscosa. Nat Prod Res 2007; 21: 824–827.

29.

Karmakar

Ishikawa

Toume

, et al. Sesquiterpenes with TRAIL-resistance overcoming activity from Xanthium strumarium. Bioorg Med Chem 2015; 23: 4746–4754.

30.

Lee

Nam

, et al. Tomentosin displays anti-carcinogenic effect in human osteosarcoma MG-63 cells via the induction of intracellular reactive oxygen species. Int J Mol Sci 2019; 20: 1508.

31.

Woo

Hakem

Soengas

, et al. Essential contribution of caspase 3/CPP32 to apoptosis and its associated nuclear changes. Genes Dev 1998; 12: 806–819.

32.

Shi

. Mechanisms of caspase activation and inhibition during apoptosis. Mol Cell 2002; 9: 459–470.

33.

Zhu

Yong

, et al. Regulation of intrinsic and extrinsic apoptotic pathways in osteosarcoma cells following oleandrin treatment. Int J Mol Sci 2016; 17: 1950.

34.

Ashkenazi

. Targeting the extrinsic apoptosis pathway in cancer. Cytokine Growth Factor Rev 2008; 19: 325–331.

35.

Green

Llambi

. Cell death signaling. Cold Spring Harb Perspect Biol 2015; 7: a006080.

36.

Fulda

Galluzzi

Kroemer

. Targeting mitochondria for cancer therapy. Nat Rev Drug Discov 2010; 9: 447–464.

37.

Kroemer

Galluzzi

Brenner

. Mitochondrial membrane permeabilization in cell death. Physiol Rev 2007; 87: 99–163.

38.

Brentnall

Rodriguez-Menocal

De Guevara

, et al. Caspase-9, caspase-3 and caspase-7 have distinct roles during intrinsic apoptosis. BMC Cell Biol 2013; 14: 32.

39.

Brunet

Bonni

Zigmond

, et al. Akt promotes cell survival by phosphorylating and inhibiting a Forkhead transcription factor. Cell 1999; 96: 857–868.

40.

Zhang

Tang

Hadden

, et al. Akt, FoxO and regulation of apoptosis. Biochim Biophys Acta 2011; 1813: 1978–1986.

41.

Nicolas

Peros

Lacombe

, et al. Genetic diversity, linkage disequilibrium and power of a large grapevine (Vitis vinifera L) diversity panel newly designed for association studies. BMC Plant Biol 2016; 16: 74.

42.

Sang

Cao

Wang

, et al. Overexpression or silencing of FOXO3a affects proliferation of endothelial progenitor cells and expression of cell cycle regulatory proteins. PLoS One 2014; 9: e101703.

43.

Sunters

Fernandez de Mattos

Stahl

, et al. FoxO3a transcriptional regulation of Bim controls apoptosis in paclitaxel-treated breast cancer cell lines. J Biol Chem 2003; 278: 49795–49805.

44.

Reinhardt

Schumacher

. The p53 network: cellular and systemic DNA damage responses in aging and cancer. Trends Genet 2012; 28: 128–136.

45.

Jacks

Remington

Williams

, et al. Tumor spectrum analysis in p53-mutant mice. Curr Biol 1994; 4: 1–7.

46.

Oda

Arakawa

Tanaka

, et al. p53AIP1, a potential mediator of p53-dependent apoptosis, and its regulation by Ser-46-phosphorylated p53. Cell 2000; 102: 849–862.

47.

Lambert

Kashanchi

Radonovich

, et al. Phosphorylation of p53 serine 15 increases interaction with CBP. J Biol Chem 1998; 273: 33048–33053.

48.

Stewart

Pietenpol

.p53 Signaling and cell cycle checkpoints. Chem Res Toxicol 2001; 14: 243–263.

49.

Castrogiovanni

Vandaudenard

Waterschoot

, et al. Decrease of mitochondrial p53 during late apoptosis is linked to its dephosphorylation on serine 20. Cancer Biol Ther 2015; 16: 1296–12307.

50.

Vogelstein

Lane

Levine

. Surfing the p53 network. Nature 2000; 408: 307–310.

51.

Wang

Kinzler

, et al. PUMA mediates the apoptotic response to p53 in colorectal cancer cells. Proc Natl Acad Sci U S A 2003; 100: 1931–1936.

52.

Zhang

Hwang

, et al. PUMA induces the rapid apoptosis of colorectal cancer cells. Mol Cell 2001; 7: 673–682.

53.

Harper

Adami

Wei

, et al. The p21 Cdk-interacting protein Cip1 is a potent inhibitor of G1 cyclin-dependent kinases. Cell 1993; 75: 805–816.

54.

Galea

Nourse

Wang

, et al. Role of intrinsic flexibility in signal transduction mediated by the cell cycle regulator, p27^Kip1 . J Mol Biol 2008; 376: 827–838.

55.

Tsytlonok

Sanabria

Wang

, et al. Dynamic anticipation by Cdk2/Cyclin A-bound p27 mediates signal integration in cell cycle regulation. Nat Commun 2019; 10: 1676.

56.

Wagner

Nebreda

. Signal integration by JNK and p38 MAPK pathways in cancer development. Nat Rev Cancer 2009; 9: 537–549.

57.

Dhanasekaran

Reddy

. JNK signaling in apoptosis. Oncogene 2008; 27: 6245–6251.

58.

Johnson

Lapadat

. Mitogen-activated protein kinase pathways mediated by ERK, JNK, and p38 protein kinases. Science 2002; 298: 1911–1912.

59.

Cagnol

Chambard

. ERK and cell death: mechanisms of ERK-induced cell death—apoptosis, autophagy and senescence. FEBS J 2010; 277: 2–21.

Induction of cell cycle arrest and apoptosis by tomentosin in hepatocellular carcinoma HepG2 and Huh7 cells

Abstract

Keywords

Introduction

Materials and methods

Reagent

Cell culture

WST-1 assay

Colony formation assay

Cell counting assay

TUNEL assay

Annexin V/propidium iodide double staining assay

Cell cycle analysis

Western blotting analysis

Statistical analysis

Results

Tomentosin inhibits the growth of HepG2 and Huh7 cells

Tomentosin induces cell cycle arrest and apoptosis in HepG2 and Huh7 cells

Tomentosin increases apoptosis in HepG2 and Huh7 cells

Tomentosin activates cell cycle arrest and apoptotic signaling pathway in HepG2 and Huh7 cells

Discussion

Footnotes

Acknowledgements

Author contributions

Declaration of conflicting interests

Funding

ORCID iD

References