Inhibitory effect of dihydromyricetin on the proliferation of JAR cells and its mechanism of action.

Dihydromyricetin (DMY) is a novel natural drug with antitumor activity against some cancer cells without obvious toxicity. Previously, its apoptotic effect on human choriocarcinoma was detected. The present study further investigated the therapeutic potential of DMY as a new drug for the treatment of choriocarcinoma, as well as its anti-proliferative effect and mechanism of action. The short-term proliferation of JAR cells was determined by MTT assay, whereas the effect of DMY on long-term cell proliferation was determined by colony forming assay. Flow cytometry was used to detect changes in the cell cycle. Furthermore, western blotting was used to detect the expression levels of proliferation-associated proteins such as cyclin A1, cyclin D1, SMAD3 and SMAD4. Reverse transcription-quantitative PCR (RT-qPCR) was used to quantify mRNA expression levels. The results indicated that DMY inhibited short and long-term proliferation of JAR cells in a concentration-dependent manner. Flow cytometry demonstrated S/G2/M cell cycle arrest, and western blotting revealed the downregulation of SMAD3, SMAD4, cyclin A1 and cyclin D1 expression levels. The results of RT-qPCR and western blotting were consistent. Overall, the findings of the present study suggest that DMY inhibits the proliferation of human choriocarcinoma JAR cells, potentially through cell cycle arrest via the downregulation of cyclin A1, cyclin D1, SMAD3 and SMAD4 expression levels. Copyright: © Zuo et al.

Introduction

Choriocarcinoma is the most aggressive type of trophoblastic neoplasia (1). It is prone to early distant metastases and has aggressive features, leading to a generally poor prognosis (2,3). With advancements in multidrug combination chemotherapy, the cure rate of choriocarcinoma has significantly improved (4). However, some patients still fail to respond to treatment due to the strong proliferative and invasive abilities of choriocarcinoma, as well as the severe toxic side effects of chemotherapy, and multidrug resistance to chemotherapy (5). Thus, it is important to inhibit the proliferation and invasion of choriocarcinoma cells, and to explore new anticancer drugs with low toxicity and high efficiency.

Dihydromyricetin (DMY) is a natural flavonoid isolated from Ampelopsis grossedentata, which reportedly has anti-inflammatory, antioxidative, anticancer, antimicrobial and immune-enhancing effects (6). DMY exerts a strong antitumor effect with low toxicity to normal cells (7–9). Its mechanism of action primarily involves the inhibition of proliferation, migration and invasion, as well as the induction of apoptosis (10–12). In a previous study, the effect of DMY-induced apoptosis on choriocarcinoma cells was studied (13). The present study aimed to explore the inhibitory effect of DMY on the proliferation of JAR cells and its associated mechanisms of action, in order to provide data to support the development of DMY as a novel therapy for choriocarcinoma.

Cell proliferation is closely associated with the cell cycle, which is an essential mechanism by which all living organisms grow and proliferate. The eukaryotic cell cycle usually comprises four phases. The cells grow continuously during interphase, which comprises three phases: G1, S And G2 (14). The G1 phase is the gap between the M and S phases, and it is an active period of metabolic activity, cell growth and repair. It is an important phase wherein the cells decide whether to enter the S phase. The S phase is flanked by the G1 and G2 phases, during which the cells continue to grow. During the S phase, the cells undergo DNA replication. The G2 phase is the gap between the S and M phases, and DNA synthesis occurs in anaphase, which is the preparatory phase of mitosis. During this period, DNA synthesis is terminated; furthermore, several RNAs and proteins. including tubulin and maturation promoters. are produced. At specific points in the G2 phase, cells encounter different checkpoints at which the decision of proceeding to the next phase or pausing to allow more time for preparation is made. The S, G1 and G2 phases form part of the interphase, which is a markedly active period for proliferating cells. The M phase is the mitotic phase; during this phase, the nucleus divides via mitosis, which is followed by cytoplasmic division via cytokinesis (14). Cyclins are important proteins in the cell cycle control system. Detection of changes in the cell cycle and cyclins can further clarify the mechanisms underlying cell proliferation.

The transforming growth factor-β (TGF-β)/SMAD signaling pathway is also closely associated with cell proliferation. Previous studies have shown that this pathway can regulate the proliferation of choriocarcinoma cells (15,16). However, the inhibitory effect of DMY on proliferation, in association with the TGF-β/SMAD signaling pathway, needs further clarification.

Materials and methods

Reagents

The following commercial reagents were used: DMY (>99% purity; Beijing Century Aoke Biotechnology Co., Ltd.), fetal bovine serum (Gibco; Thermo Fisher Scientific, Inc.), DMEM (Gibco; Thermo Fisher Scientific, Inc.), 0.25% trypsin with 0.02% EDTA (Gibco; Thermo Fisher Scientific, Inc.), MTT reagent (Gibco; Thermo Fisher Scientific, Inc.), antibodies against GAPDH (rabbit anti-human polyclonal; cat. no. E12-052; EnoGene Biotech Co., Ltd.), SMAD3 (rabbit anti-human monoclonal; cat. no. ab40854; Abcam), p-SMAD3 (rabbit anti-human monoclonal; cat. no. ab52903; Abcam), SMAD4 (rabbit anti-human monoclonal; cat. no. ab40759; Abcam), Cyclin A1 (rabbit anti-human polyclonal; cat. no. E1A5313; EnoGene Biotech Co., Ltd.) and Cyclin D1 (rabbit anti-human monoclonal; cat. no. ab134175; Abcam), and a secondary antibody (peroxidase-conjugated AffiniPure goat anti-rabbit IgG; cat. no. ab6721; Abcam).

Cell culture

The human JAR cell line (male fetus choriocarcinoma) was purchased from Guangzhou Jennio Biotech Co., Ltd. JAR cells were cultured in 10% DMEM at 37°C in the presence of 5% CO2. Subsequently, the cells were subcultured with 0.25% trypsin and 0.02% EDTA until confluence reached 70–80%.

MTT assay

MTT assays were performed as described previously (13). Cells were seeded in 96-well plates, cultured overnight until the cell confluence reached 30–40%, and incubated for 48 h with various concentrations of DMY: 0, 20, 40, 80, 100, 120, 160, 240 and 320 mg/l (0, 62, 125, 250, 312, 375, 500, 749 and 999 µM), in 200 µl 10% DMEM per well. There were 6 wells for each group. MTT reagent was used to determine cell proliferation, and absorbance was determined using a microplate reader (Multiskan MK3; Thermo Fisher Scientific, Inc.) at a wavelength of 492 nm. The experiment was repeated three times.

Live cell imaging

According to the MTT results, 0, 40, 60 and 100 mg/l (0, 125, 187 and 312 µM) were selected as the appropriate concentrations to perform further experiments. Cell proliferation was observed under an inverted light microscope at ×100 magnification (Olympus IX73; Olympus Corporation), and cell confluence was analyzed using the PAULA live cell imaging system (Leica Microsystems GmbH).

Colony forming assay

Cells were seeded in culture plates and allowed to adhere overnight. Once the confluence reached 30–40%, the cells were incubated with different concentrations of DMY (0, 40, 60 and 100 mg/l) in 10% DMEM at 37°C of 5% CO2. Following a 48-h incubation, the cells were resuspended and subsequently replated at a density of 400 cells/well in 6-well plates. After a 12-day culture, the cells were stained with crystal violet for 30 min at room temperature, colonies were counted using ImageJ version 1.46 (National Institutes of Health), and images of cell morphology were captured using a camera (Nikon Corporation).

Cell cycle analysis

Cells were seeded in culture plates and allowed to adhere overnight. Once the cell confluence had reached 30–40%, the cells were incubated with different concentrations of DMY (0, 40, 60 and 100 mg/l). After a 48-h incubation period, the cell cycle was analyzed using flow cytometry. The cells were processed with the Cell Cycle kit [cat. no. CCS012; Multi Sciences (Lianke) Biotech Co., Ltd.], according to the manufacturer's instructions. The samples were analyzed using a FACSCalibur flow cytometer (Becton, Dickinson and Company).

Reverse transcription-quantitative PCR (RT-qPCR)

Cells were incubated with different concentrations of DMY (0, 40, 60 and 100 mg/l) for 48 h. Total RNA was isolated using TRIzol® reagent (Sigma-Aldrich; Merck KGaA), according to the manufacturer's instructions. cDNA synthesis was performed using the PrimeScript RT Reagent kit (cat. no. RR036A; Takara Biotechnology Co., Ltd.) according to the manufacturer's instructions. qPCR was performed using the SYBR Green qPCR Master mix (cat. no. RR820A; Takara Biotechnology Co., Ltd.), according to the manufacturer's instructions. PCR was performed under the following conditions: Denaturation at 95°C for 30 sec, followed by 40 cycles at 95°C for 5 sec, 55°C for 30 sec and 72°C for 30 sec, finally 1 cycle at 95°C for 60 sec, 55°C for 30 sec and 95°C for 30 sec. Three repeated wells were set up of each sample. The expression intensity of each gene was normalized against the expression of GAPDH. Differential gene expression was calculated as the ratio of the expression levels of target genes in JAR cells using the 2−ΔΔCq method (17). The PCR primer sequences are presented in Table I.

Table I.

Primer pairs used in quantitative PCR.

Gene	Sequence (5′ to 3′)
SMAD4	F: gacagcagcagaatggat
	R: caggagcaggatgattgg
SMAD3	F: ccagggctttgaggctgtcta
	R: gcaaaggcccattcaggtg
Cyclin A1	F: tgtcaccgttcctccttg
	R: gcatcttcacgctctattt
Cyclin D1	F: gcgaggaacagaagtgcg
	R: tggagttgtcggtgtagatgc
GAPDH	F: gcaccgtcaaggctgagaac
	R: tggtgaagacgccagtgga

F, forward; R, reverse.

Western blotting

Cells were incubated with different concentrations of DMY (0, 40, 60 and 100 mg/l) for 48 h, collected and lysed on ice with RIPA buffer. The cell lysates were centrifuged at 14,000 × g for 10 min at 4°C. Subsequently, the supernatant was extracted, and the protein concentration was quantified using a bicinchoninic acid assay kit (Pierce; Thermo Fisher Scientific, Inc.). For SDS-PAGE, 12% gel was used to separate 30 µg protein per sample, which was then transferred onto a polyvinylidene difluoride membrane. Nonfat milk (5%) was used to block the membranes for 2 h at room temperature. Next, the membranes were incubated with primary antibodies overnight at 4°C. The primary antibodies used were against GAPDH (1:2,000), SMAD3 (1:5,000), SMAD4 (1:5,000), cyclin A1 (1:1,000) and cyclin D1 (1:10,000). Subsequently, the membranes were washed in 0.1% Tween 20-TBST buffer and incubated with an appropriate secondary antibody (1:5,000) for 1 h at room temperature. Immunoreactive bands were detected using an enhanced chemiluminescent substrate (Pierce; Thermo Fisher Scientific, Inc.) and imaged using the Tanon 6100 Chemiluminescent Imaging system (Tanon Science and Technology Co., Ltd.). Band densities were calculated using the Quantity One software version 4.6.2 (Bio-Rad Laboratories, Inc.).

Statistical analysis

All experiments were repeated ≥3 times, unless otherwise indicated. Data were analyzed using SPSS 19.0 (IBM Corp.) and presented as the mean ± standard deviation. Statistical analysis was performed using one-way ANOVA followed by Tukey's post hoc test. P<0.05 was considered to indicate a statistically significant difference.

Results

DMY inhibits the proliferation and colony formation of JAR cells

The MTT assay results showed that the proliferation of JAR cells was significantly inhibited by DMY in a concentration-dependent manner compared with that of the untreated cells (P<0.05; Fig. 1A). Cell confluence decreased significantly with increasing concentrations of DMY, as assessed using an inverted light microscope (P<0.05; Fig. 1B). The colony forming assay demonstrated that the number of JAR cells clones decreased significantly with increasing concentrations of DMY, compared with the number of untreated cell clones (P<0.05; Fig. 1C).

Figure 1.

Proliferative ability of JAR cells incubated with different concentrations of DMY for 48 h. (A) Inhibition of JAR cell proliferation with different concentrations of DMY based on MTT assay. (B) Changes in cell confluence observed using an inverted light microscope at ×100 magnification. Scale bar, 100 µm. (C) Colony forming assay performed in 6-well plates with colonies of JAR cells incubated with 0, 40, 60 and 100 mg/l DMY (0, 125, 187 and 312 µM) for 48 h, after which the cells were resuspended and subsequently replated at a density of 400 cells/well. *P<0.05 vs. 0 mg/l DMY group. DMY, dihydromyricetin. Scale bar, 1 cm.

DMY affects cell cycle distribution in JAR cells

The effects of the different concentrations of DMY were evaluated on the cell cycle distribution in JAR cells using flow cytometry. Compared with the number of untreated cells, increasing concentrations of DMY (60 and 100 mg/l) significantly decreased the number of JAR cells in the G1 phase, whilst increasing the number of cells in the S/G2/M phase (Fig. 2).

Figure 2.

Cell cycle analysis. Different concentrations of DMY (60 and 100 mg/l) significantly decreased the number of JAR cells in the G1 phase and increased the number of cells in the S/G2/M phase. *P<0.05 vs. 0 mg/l group. DMY, dihydromyricetin.

DMY regulates the mRNA expression levels of SMAD3, cyclin A1 and cyclin D1 in JAR cells

After a 48-h incubation with different concentrations of DMY, the mRNA expression levels of SMAD3, SMAD4, cyclin A1 and cyclin D1 were quantified using RT-qPCR (Fig. 3). The results indicated that the mRNA expression levels of cyclin A1 and cyclin D1 decreased with 60 and 100 mg/l DMY (P<0.01). The mRNA expression level of SMAD3 decreased with 40, 60 and 100 mg/l DMY (P<0.05, P<0.01). However, no significant differences were observed in the mRNA expression levels of SMAD4 (P>0.05; Fig. 3).

Figure 3.

Reverse transcription-quantitative PCR detected mRNA expression in JAR cells incubated with different concentrations of DMY (0, 40, 60 and 100 mg/l) for 48 h. The mRNA expression levels of cyclin A1 and D1 decreased with 60 and 100 mg/l of DMY, though particularly with 100 mg/l DMY. The mRNA expression level of SMAD3 decreased with 40, 60 and 100 mg/l DMY. No significant differences were found in the mRNA expression of SMAD4. *P<0.05 and #P<0.01 vs. 0 mg/l DMY. DMY, dihydromyricetin.

DMY regulates the protein expression levels of SMAD3, p-SMAD3, SMAD4, cyclin A1 and cyclin D1 in JAR cells

Western blotting revealed that the protein expression levels of SMAD3, p-SMAD3, SMAD4, cyclin A1 and cyclin D1 were decreased in JAR cells treated with DMY compared with those in the untreated cells particularly in the 100-mg/l group. The ratio of p-SMAD3/SMA3 increased in 100 mg/l group (P<0.05; Fig. 4).

Figure 4.

Western blotting. JAR cells were incubated with different concentrations of DMY (0, 40, 60 and 100 mg/l) for 48 h. The protein expression levels of SMAD3, p-SMAD3, SMAD4, cyclin A1 and D1 were decreased in DMY-treated cells compared with the untreated cells. Of note, the ratio of p-SMAD3/SMA3 increased in 100 mg/l group, we speculate the increase was due to an increase in the bands above p-SMAD 3 instead. *P<0.05 and #P<0.01 vs. 0 mg/l DMY. DMY, dihydromyricetin.

Discussion

In the present study, MTT and colony forming assays were performed to determine the short and long-term inhibitory effects of different concentrations of DMY on the proliferation of JAR cells, respectively. It was found that DMY inhibited the short and long-term proliferation of JAR cells in a concentration-dependent manner. The cell cycle, apoptosis and some signaling pathways are closely associated with cell proliferation. In a previous study, DMY was demonstrated to inhibit cell proliferation by inducing apoptosis in JAR cells (13). In the present study, the inhibitory effect of DMY on the proliferation of JAR cells was explored, based on the cell cycle and TGF-β/SMAD signaling pathway (15). The cell cycle was analyzed using flow cytometry, and the results showed that DMY inhibited cell proliferation in association with the regulation of the cell cycle.

Flow cytometry revealed S/G2/M cell cycle arrest in JAR cells treated with DMY. Cyclins are important in the cell cycle control system; cyclins progressively accumulate throughout interphase and abruptly disappear during mitosis during each cell cycle. Escape from cell cycle arrest or from certain nonproliferating states requires the accumulation of cyclins (14). Since the discovery of cyclins in sea urchins by Evans et al (18) in 1983, our understanding of the functions of cyclins in the cell cycle has improved. In mammalian cells, cyclin A accumulates during the late G1 phase and is degraded before metaphase (19). Cyclin D was first isolated in a study on parathyroid carcinoma conducted by Motokura et al (20), and it is necessary in the G1/S phase of the cell cycle. Cyclin D is generally considered to play a regulatory role by combining with cycle-dependent kinases (CDKs; CDK4 or CDK6) to promote transition from the G1 to the S phase (21). The overexpression of cyclin D1 and CDK4 can cause abnormal proliferation and uncontrolled differentiation of cells; these processes are closely associated with the occurrence and development of tumors (22,23). Numerous studies have shown that cyclins are fundamental to the cell cycle, which is important in cell proliferation and differentiation, and is closely associated with the occurrence and development of tumors. For example, alantolactone suggested to inhibit cell proliferation, inducing G2/M phase arrest by downregulating cyclin A1 in HepG2 cells (24). A naturally occurring sesquiterpene lactone-Santonin causes SKBR-3 cell cycle arrest at the G2/M phase by suppressing the expression of cyclins A and B1 (25). MircoRNA-27a inhibition reportedly triggers G2/M arrest in SKOV-3 and OVACAR-3 cells, accompanied by the downregulation of cyclins A and B1 (26). Fucosterol (a phytosterol in marine algae and many other plant species) also triggers G2/M cell cycle arrest in A549 and SKLU-1 cells, which is associated with the downregulation of CDK1, cyclin A and cyclin B1, as well as the upregulation of negative regulators of the cell cycle (27). WDR5 induces S/G2/M cell cycle arrest via cyclin D1 in a process that is regulated by H3K4me3 (28). In the present study, the results showed that the expression levels of cyclin A1 and cyclin D1 were decreased; thus, DMY may inhibit the proliferation of JAR cells by causing S/G2/M arrest via a decrease in the expression levels of cyclin A1 and D1.

In addition to the association between cyclins and cell proliferation, some studies have shown that the TGF-β/SMAD signaling pathway is also associated with cell proliferation (29,30). TGF-β activates TGF-β receptor (TβR)I and TβRII, resulting in the phosphorylation of receptor-regulated SMAD2/3, which is associated with the common mediator SMAD4. The SMAD2/3/4 complex is translocated to the nucleus, where it binds to DNA and regulates the transcription of several genes (31,32). Increased mRNA and protein expression levels of SMAD3 and SMAD4 promote cell proliferation, migration and invasion (33). Previous studies have shown that the TGF-β/SMAD signaling pathway can regulate the proliferation of choriocarcinoma cells (15,16). In the present study, the mRNA and protein expression levels of SMAD3 and SMAD4 were examined, and the results revealed significantly decreased mRNA and protein expression levels of SMAD3 in cells treated with 100 mg/l DMY. The protein expression levels of p-SMAD3 also decreased in 100 mg/l group. Of note, the ratio of p-SMAD3/SMA3 increased in 100 mg/l group, and this may be due to an increase in the bands above p-SMAD 3 instead. No significant changes were detected in the mRNA expression level of SMAD4; however, its protein expression was significantly reduced in the 100-mg/l group. Therefore, it was hypothesized that the treatment might increase protein degradation. Overall, the results indicate that DMY may inhibit the proliferation of JAR cells by decreasing the expression of SMAD3 and SMAD4.

However, one limitation of the present study is that no rescue experiments were performed to determine whether the overexpression of cyclin A1, cyclin D1, SMAD3 and SMAD4 prevents the inhibitory effect of DMY on the proliferation of JAR cells. Nevertheless, the present results provide a foundation for future investigations into more detailed mechanisms.

In summary, the findings of the present study suggest that DMY inhibits the proliferation of human choriocarcinoma JAR cells, potentially by inducing cell cycle arrest via the downregulation of cyclin A1, cyclin D1, SMAD3 and SMAD4. However, the role of other mechanisms, and whether the overexpression of cyclin A1, cyclin D1, SMAD3 and SMAD4 prevents the inhibitory effect of DMY on the proliferation of JAR cells need further clarification. Future studies on DMY will aid in developing it as a potential novel drug for the treatment of choriocarcinoma.