The Journal of Toxicological Sciences
Online ISSN : 1880-3989
Print ISSN : 0388-1350
ISSN-L : 0388-1350
Original Article
Euptox A Induces G0 /GI arrest and apoptosis of hepatocyte via ROS, mitochondrial dysfunction and caspases-dependent pathways in vivo
Samuel Kumi OkyereQuan MoGao PeiZhihua RenJunliang DengYanchun Hu
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2020 Volume 45 Issue 11 Pages 661-671

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Abstract

As a toxin of Ageratina adenophora (A. adenophora), euptox A (9-oxo-10, 11-dehydroageraphorone) is known to cause hepatotoxicity in animals. In this study, we examined the effects of euptox A on mouse liver cells and its underlying mechanisms for the first time. We found that euptox A induced liver cell cycle arrest and apoptosis in a dose-dependent manner mainly by mitochondria -related pathways, with the affected cells characterized by the appearance of DNA fragmentation, membrane blebbing, and chromatin condensation. The results showed that euptox A similarly induced hepatocyte G0 /GI arrest and apoptosis mainly by ROS accumulation and mitochondria-mediated and caspase-dependent pathways, elucidated by the loss of mitochondrial membrane potential, release of cytochrome C and AIF, activation of caspase-3/-9, Bax, as well as suppression of Bcl-2. This paper will provide new insights into the mechanisms involved in liver toxicity caused by euptox A in mice.

INTRODUCTION

A. adenophora has been reported to induce hepatotoxic effects on various organs of animals (Oelrichs et al., 1995; Katoch et al., 2000; Sun et al., 2019). Euptox A (9-oxo-10, 11-dehydroageraphorone) is one major cadenine sesquiterpene toxin of A. adenophora leaves (Fig. 1) (Ouyang et al., 2014; Seawright et al., 1998; He et al., 2008). Bai et al. (2011) reported that euptox A was mainly distributed in leaves with a detected value of 1.34-8.41 mg∙g–1 while the amount of euptox A present in flowers, stems and roots was less than 0.06 mg∙g–1. Euptox A has been reported to induce lesions in the liver of rats and mice (Oelrichs et al., 1995; Bohlmann and Gupta, 1981; Kaushal et al., 2001a). Also, Euptox A induces pathological changes in hepatic lobules and hepatocytes with focal necrosis of the parenchyma and degeneration of the epithelial lining in the small bile ducts in the liver of mice (Ouyang et al., 2014; Kaushal et al., 2001b). However, the mechanism underlying the liver toxicity of euptox A is still infinite.

Fig. 1

Structure of euptox A.

The liver plays a major function in the detoxification of ingested toxins (Xie et al., 2018). Our previous studies reported that A. adenophora caused liver inflammatory injury and induced hepatocyte pyroptosis by activating NLRP3 inflammasome, which was triggered by elevating ROS production levels (Sun et al., 2018). Thus, for the first time, we want to evaluate the hepatotoxicity induced by euptox A in relation to cell arrest and apoptosis and further investigate the underlying mechanisms associated with euptox A toxicity.

Reactive oxygen species (ROS) play important roles in modulating signaling pathways for various cellular events, such as cell proliferation and apoptosis (Jabs, 1999). ROS are highly reactive oxygen free radicals or non-radical molecules which can be produced by multiple mechanisms, and mainly originate from NADPH oxidase (NOX) and the mitochondria (Li et al., 2011). Intracellular ROS generation plays a key role in numerous physiological and pathological processes, and a high level is often closely associated with apoptotic cell death (Laurent et al., 2005).

Apoptosis is a major type of programed cell death (Neuman et al., 1999). Apoptosis, programed cell death, is triggered by activating the intrinsic mitochondrial and/or the extrinsic death receptor pathways. The intrinsic mitochondrial pathway is regulated by multiple factors such as the Bcl-2 family, Cyc c, AIF, and caspase, while the extrinsic death receptor pathway is mediated by a variety of death receptor ligands such as tumor necrosis factor (TNF), Fas, and Fas ligand (FasL) (Zeng et al., 2012). The caspase family plays an important role in the initiation and execution of cell apoptosis pathways. Caspase-8 and -9 both initiate caspases in different pathways, with the former in the death receptor pathway while the latter in the mitochondrial pathway. Caspase-3, which is a downstream of caspase-8 and -9, plays a key role in the apoptosis pathway as momentous executioner caspase (Kantari and Walczak, 2011; Li and Yuan, 2008).

In this report, we studied the cytotoxic effects of euptox A on hepatocytes and the ability of euptox A to induce apoptosis in both liver cells and tissues. We also investigated the ability of euptox A to modulate the cell cycle and ROS elevation for understanding the mechanisms related to the hepatotoxicity of euptox A in animals.

MATERIALS AND METHODS

Extraction and purification of euptox A

A. adenophora was collected from Xichang city in Sichuan province, southwest China. Fifty grams of leaves were milled and mixed with 100 mL water. A mixture that contained euptox A, coumarin, gallotannic acid and volatile oil was ultrasonically extracted by carbinol and hexyl acetate for 30 min at 40°C. In order to separate euptox A from the extraction, samples were purified using the Silica Column Chromatography method. Silica Gel Thin-layer Chromatography was then used to analyze the existence of euptox A in the final extraction. HPLC was used to determination purity results of the toxin we extracted, and the result showed 96% purity.

Animal model

The animal model was established in accordance with several previous reports. Adult mice (8 weeks, 25-30 g) were obtained from the animal experimental center of Sichuan Agricultural University. The animals were maintained in an air-conditioned room (22 ± 3°C) on a 12-hr light/dark cycle with free access to water and food. The animals were randomly divided into four groups with eighteen animals in each group: control group (n = 18) and treatment groups (n = 54). The control group received 0.9% normal saline, while treatment groups received 200, 400 and 800 mg/kg euptox A via intragastric gavage daily for 30 days, respectively referring to the findings of Ouyang et al. (2014). After the experiment, mice were sacrificed via cervical dislocation to obtain the liver tissues. All methods were carried out in accordance with the approved guidelines and experimental protocols by the Animal Care and Use Committee of Sichuan Agricultural University, China.

Measurement of intracellular ROS

The formation of intracellular ROS was measured using the DCFH-DA method with slight modifications in this study (Sohn et al., 2005). The livers were taken from each mouse immediately and then minced by scissors in order to make a cell suspension. Then the cell suspension was filtered through a 300-mesh nylon screen, washed twice with cold PBS and incubated in darkness with DCFH-DA at a final concentration of 5 µm for an additional 40 min at 37°C. Intracellular ROS generation was measured by the flow cytometer with excitation and emission wavelengths set as 488 and 528 nm, respectively. Each experiment was conducted three times, and the results were reported as the mean of the three experiments.

Apoptosis assessment

This was performed by using 4’, 6-diamidino-2-phenylindole (DAPI) and acridine orange/ethidium bromide (AO/EB) staining. After removing the livers, we minced them to make a cell suspension which was filtered through a 300-mesh nylon screen subsequently. For DAPI staining, the hepatic cells were first fixed with 80% ethanol at room temperature for 30 min. Then, we immediately removed the fixative and washed the hepatic cells with PBS for 3 times. After that, we incubated the hepatic cells with DAPI (1 μg/mL) in the dark at room temperature for 45 min. For AO/EB staining, 100 μL fresh-prepared AO/EB staining solution (100 μg/mL) were loaded into the liver cells that had not been fixed before, and then the liver cells were observed with a Nikon fluores-cence microscope (Nikon Inc., Tokyo, Japan) within 20 min.

Mitochondrial membrane potential assay

The transmembrane potential, ΔΨm was analyzed using a JC-1 Mitochondrial Potential Detection Kit (Biotium Inc., Hayward, CA, USA). The cell suspension was filtered through a 300-mesh nylon mesh, washed twice with cold PBS and stained by 5,5′,6,6′-tetrachloro-1,1′,3,3′ tetraethylbenzimidazolcarbocyanine iodide (JC-1; Molecular Probes, Eugene, OR, USA) in PBS for 15 min at room temperature in the dark, followed by flow cytometric analysis.

Transmission electron microscopy (TEM)

The ultrastructure of hepatic cells was observed by TEM. After the end of intragastric administration, the cell pellet was fixed with 40 g/L glutaraldehyde in 0.1 M sucrose with 0.2 M sodium cacodylate buffer (pH 7.4) overnight at 4°C. Washed with sodium cacodylate buffer, the cells were re-fixed with 10 g/L osmium tetroxide for 1.5 hr. Following routine dehydration, epoxy resin embedding and ultrathin section, the specimens were stained with 20 g/L uranyl acetate-lead citrate and observed with an H700 transmission electron microscope (Hitachi, Tokyo, Japan).

Flow cytometry (FCM)

The cell cycle and phosphatidylserine (PS) externalization were detected by FCM. In short, the cell suspension was first filtered through 300-mesh nylon, and washed twice with cold PBS. Subsequently, the cells were suspended in PBS at a concentration of 1 × 106 cells/mL. For cell cycle assay, the cells were washed and resuspended in 500 mL propidium iodide (PI) (BD Biosciences, San Jose, CA, USA), and incubated in the dark for 30 min. For PS externalization assay, 5 mL FITC-labeled Annexin-V and 5 mL PI (BD Biosciences) were added into 500 mL of cell suspension, and incubated in the dark for 15 min. The stained hepatic cells were assayed with a FC500 MPL flow cytometer (Beckman Coulter, Fullerton, CA, USA).

Caspase activity assay

According to the manufacturer’s instructions, colorimetric assay kits (BioVision, Inc., Mountain View, CA, USA) were used to measure the activity of caspases-8, -9 and -3. Briefly, we harvested and incubated hepatic cells in ice-cold cell lysis buffer on ice. After 30 min, we collected the supernatants. BCA Protein Assay Reagent (Pierce, Rockford, IL, USA) was used for the determination of the protein concentrations. For each sample, an equivalent amount of protein was incubated with interested caspase substrate. After incubation for 4 hr at 37°C, we examined the protease activity at 405 nm with microplate spectrophotometer (Bio-Tek Instruments, Inc, Winooski, VT, USA).

TUNEL assay

The liver tissues were fixed in 4% paraformaldehyde. After embedding them in paraffin, we cut them into 6 um sections. A TUNEL assay was then conducted to examine DNA fragmentation using an in situ cell death detection kit (Vazyme, Piscataway, NJ, USA) according to the manufacturer’s instructions. After mounting the TUNEL-positive cells, DAPI was used to counterstain the nuclei and a Nikon microscope (Nikon Inc.) was used to observe the sections at × 1000 magnification.

Western blotting

After a formal trial of 30 days, the liver was carefully removed and stored at 80°C until use. To prepare lysates, the tissues samples were weighed and minced with eye scissors on ice. The livers were dissected and mechanically homogenized in 500 mL of lysis buffer (125 mM Tris, pH 6.8, 40 mM EDTA, 4% SDS), and centrifuged at 14,000 g and 4°C for 15 min. The supernatant was collected. After, protein concentration was determined with BCA protein assay kit (Thermo Scientific, Rockford, IL, USA), the protein samples were boiled for 15 min, and subjected to SDS-polyacrylamide gel electrophoresis (SDS-PAGE). The resolved protein samples were transfered to a polyvinylidene diflouride filter (PVDF) membrane (Immobilon; Millipore, Bedford, MA, USA) by a transfer apparatus at 300 mA for 2 hr. The membranes were blocked by 5% non-fat milk in TBST for 2 hr at room temperature, followed by incubation with rabbit ployclonal antibodies against caspase-8/9 /3, Cyt c, Bcl-2, Bax, Bid, AIF, PARP, Fas, FasL, p53, β-actin and COX 4 (1:1000; Santa Cruz, CA, USA) at 4°C overnight. Secondary antibody incubation was performed using HRP-conjugated goat anti-rabbit IgG (1:2000) for 2 hr, and then protein bands were visualized by using an enhanced chemiluminescence system (ECL; Pierce).

Quantitative real-time PCR

Total RNA was extracted from 50 mg liver powder using TRIzol (Aidlab, Beijing, China). The synthesis of single-stranded cDNA from 5 μg of RNA was performed using TUREscript 1st-strand cDNA Synthesis Kit (Aidlab), according to the manufacturer’s instructions. The mRNA was then reverse transcribed into cDNA which was used as a template for quantitative reverse transcription-polymerase chain reaction (qRT-PCR) analysis. The definition of relative gene expression was the ratio of target gene versus reference gene (β-actin and COX 4) expression. For the calibration of gene expression, its values in the control group were used. The results were analyzed using the 2−ΔΔCt method (Livak and Schmittgen, 2001). And the primers as follows (Table 1) used in this study were designed by NCBI BLAST tool (Boratyn et al., 2019).

Table 1. Gene-specific primers used for (qRT-PCR) analysis.
Genes Forward and reverse Primers(5- 3)
forward reverse
Fas GAACUCCGUGAGUUCACCAACCAAA UUGUCAUGUCUUCAGCAAUUCUCGG
Fasl GAACUCCGUGAGUUCACCAACCAAA UUUGGUUGGUGAACUCACGGAGUUC
Caspase 8 CAAGAAGCAGGAGACCATCGA GGTCCCACCGACTGATGT
Caspase 9 GGCCAGCCACCTCCAGTT TTCTCAATGGACACGGAGCAT
Caspase 3 CTGGACTGCGGTATTGAGAC CCGGGTGCGGTAGAGTAAGC
Bax TGGCTGGGGAGACACCTGAGC TCAGCCCATCTTCTTCCAGATG
Bcl-2 TGACTTCTCTCGTCGCTACCGT CCTGAAGAGTTCCTCCACCACC
β-actin ATATCGCTGCGCTGGTCGTC AGGATGGCGTGAGGGAGAGC

Statistical analysis

All data are expressed as the means ± SD of three independent experiments. Software SPSS18.0 (SPSS Inc., Chicago, IL, USA) was utilized for statistical analyses which were conducted to compare the control group with the experimental groups through one-way analysis of variance (ANOVA), followed by the Tukey-Kramer multiple comparison test with an equal sample size. Values with P < 0.05 and P < 0.01 were considered as statistically significant.

RESULTS

Induction of apoptosis and cell cycle arrest by euptox A

To determine the pathological effects of euptox A on the liver after 30 days treatment with 200, 400, and 800 mg/kg euptox A, livers were surgically removed during necropsy. DAPI and AO/EB staining were used for apoptosis and necrosis bodies’ identification while transmission electron microscopy (TEM) analysis was used to examine membrane blebbing of the hepatocytes. Also, Terminal deoxynucleotidyl transferase (TdT) deoxyuridine 5’-triphosphate (dUTP) nick-end labeling (TUNEL) staining, agarose gel electrophoresis and flow cytometry (FCM) assay were used to detect hepatocyte apoptosis, nuclear fragmentation and chromatin condensation in situ.

Compared with the control group, the results showed that euptox A treatment caused remarkable morphological changes including membrane blebbing, granular apoptotic bodies, chromatin condensation, and nuclear fragmentation. According to the flow cytometry analysis, the normal hepatocytes percentages in euptox A -treated groups were significantly decreased compared to the control group with the maximum effects observed in the high-dose group (Fig. 2A and 2B). A characteristic DNA ladder was apparent in the gel results of the euptox A-treated hepatocytes, and the 800 mg/kg dose group showed more intense DNA laddering than the 400 and 200 mg/kg doses group (Fig. 2B1). The flow cytometry (FCM) assay showed that, the ratio of early apoptotic cells increased from 1.20 ± 0.75% in untreated cells to 3.70 ± 0.58%, 14.54 ± 2.51%, and 19.41 ± 1.79% in the 200, 400, and 800 mg/kg euptox A-treated cells, respectively, indicating that the apoptotic rate of hepatocytes increased in a dose-dependent manner (Fig. 2C and 2C1). The DNA distribution of hepatocytes in the different cell cycle phases was analyzed using FCM. As shown in (Fig. 2D and 2D1), euptox A concentrations of 200, 400, and 800 mg/kg significantly increased the G0/G1 phase cell number from 47.2% (control) to 55.3%, 79.2%, and 88.9%, respectively. Furthermore, the percentage of hepatocytes in the S, G2 + M, and PI phases cells were significantly reduced in the 400 and 800 mg/kg groups and were markedly decreased in 200 mg/kg group compared with the control, indicating the occurrence of G0/G1 phase arrest. These results indicate that orally administration of euptox A to mouse can inhibited cell proliferation and caused cell death in hepatocytes.

Fig. 2

Euptox A induces cell cycle arrest and apoptosis in hepatocytes. Mice were treated with different doses of euptox A for 30 days. (A) Apoptotic hepatocytes were detected using DAPI and AO/EB staining as well as TEM and TUNEL assays. Representative liver sections from mice were analyzed using TUNEL assays to detect apoptotic cell death. TUNEL-positive cells (black arrows) in the liver were counted from five random microscopic fields at 600× magnification. Nuclear morphological changes in hepatocytes were observed using fluorescence microscopy after DAPI (200×) staining with changes represented with white arrows and AO/EB (400×) staining with early apoptosis, late apoptosis and necrosis represented with white, yellow, and red arrows, respectively. Ultrastructural morphological changes such as membrane blebbing of hepatocytes were also observed using TEM at 10000× magnification. (B) DNA fragmentation was determined using DNA ladder extraction kit. (B1) DNA laddering histogram of hepatocytes shows that, the 800 mgkg-1 dose group showed more intense DNA laddering than the 400 and 200 mgkg-1 doses group. (C) Annexin V-PI double staining shows significant euptox A-induced apoptosis of hepatocytes. (C1) Histogram of apoptotic rate of hepatocytes was measured with the flow cytometry (FCM) assay which showed that, the ratio of early apoptotic cells increased from 1.20 ± 0.75% in untreated cells to 3.70 ± 0.58%, 14.54 ± 2.51%, and 19.41 ± 1.79% in the 200, 400, and 800 mgkg-1 euptox A-treated cells, respectively, indicating that the apoptotic rate of hepatocytes increased in a dose-dependent manner. (D) The DNA distribution of hepatocytes in the different cell cycle phases was analyzed using FCM. (D1) DNA histogram of hepatocytes cell cycle was analyzed using flow cytometry (FCM) with PI staining and percentage G0/G1, S, and G2 + M phases of hepatocytes was analyzed. Percentage (%) proliferating index (pi) value = [S + (G2 + M)]/[(G0/G1) + S + (G2 + M)] × 100. Data are presented as means ± standard deviation (SD) of three independent experiments. *p < 0.05 and **p < 0.01 compared with control group. DAPI, 4’,6-diamidino-2-phenylindole; AO/EB, acridine orange/ethidium bromide; TEM, transmission electron microscopy. TUNEL, terminal deoxynucleotidyl transferase (TdT) deoxyuridine 5’-triphosphate (dUTP) nick-end labeling; PI, propidium iodide.

Euptox A increased intracellular ROS generation in hepatocyte apoptosis

In determining intracellular ROS levels, FCM with dichlorodihydro-fluorescein diacetate (DCFH-DA) staining and fluorescence assays were used for analysis. As shown in Fig. 3A, liver cells treated with 200, 400, and 800 mg/kg euptox A for 30 days showed a concentration-dependent increase in ROS levels. This effect was also observed in the fluorescence assays (Fig. 3B), which suggests that ROS may play a significant role in the euptox A-induced apoptosis and cell arrest.

Fig. 3

Intracellular accumulation of reactive oxygen species (ROS). (A) Euptox A induced ROS production in liver cells. Mice were treated with different doses of euptox A for 30 days, cells were harvested and incubated with 5 μm DCFDA for 40 min and then analyzed using flow cytometry (FCM). Ratio of green fluorescence intensity was presented as means ± standard deviation (SD) of three experiments in triplicate. *p < 0.05 and **p < 0.01 compared with control group. (B) Intracellular ROS level was evaluated using fluorescent microscopy (400× magnification).

Euptox A -induced hepatocyte apoptosis by caspase-9 and -3 dependent pathways

To determine the caspases involved in the euptox A-induced apoptosis, the activities of caspase-8, -9, and -3 were measured in hepatocytes using colorimetric assay kits, Western blot and qRT-PCR. In comparison with the control groups, euptox A treatment groups significantly induced the activation of caspases-9 and -3, but not caspase-8 during the treatment period (Fig. 4A). Western blot analysis revealed that as the dose of euptox A increased, full-length procaspase-9 and -3 declined while their cleavage increased (Fig. 4B). However, the cleaved form of procaspase-8 was not found in this study (Fig. 4B). PARP could sever as an indicator of caspase-3 activation during apoptosis (Thornberry and Lazebnik, 1998). PARP appeared to be obviously cleaved after 400 and 800 mg/kg euptox A treatment for 30 days (Fig. 4B). Moreover, exposure of hepatocytes to euptox A 200, 400, and 800 mg/kg resulted in a 1.72-, 3.03-, and 3.76-fold increase in the mRNA levels of caspase-9, respectively. The mRNA levels of caspase-3 increased by 1.80-, 3.12-, and 4.19-fold when hepatocytes were treated with euptox A 200, 400, and 800 mg/kg/day respectively. However, the mRNA levels of caspase-8 were not significantly changed except in the high-dose group (Fig. 4C) indicating that orally administration of euptox A to mouse can induce caspase-9 and caspase-3 in hepatocyte apoptosis.

Fig. 4

Effects of euptox A treatment on caspases activation and poly ADP-ribose polymerase (PARP) cleavage in hepatocytes. (A) Caspase activities in euptox A-treated hepatocytes. BCA assay was used to verify equal amounts of protein and enzymatic activities of caspase-8, -9, and -3 were measured using colorimetric assay kits. (B) Western blot analysis and quantification of caspase-8/9/3 and cleaved-PARP expression with β-actin as loading control. (C) Relative mRNA levels of caspase-8, -9, and -3. Mice were treated with different doses of euptox A for 30 days, and mRNA was extracted from hepatocytes and analyzed using qRT-PCR analysis. Dates are presented with the means ± SD and mean values of three independent experiments. *P < 0.05 and **P < 0.01, compared with the control group. BCA, bicinchoninic acid; qRT-PCR, quantitative reverse transcription-polymerase reaction.

Euptox A induces hepatocyte apoptosis by mitochondria dysfunction pathways

In this study, western blot and qRT-PCR were used to show the protein and mRNA levels of Fas, FasL, and Bid. Also, in accessing the changes in mitochondrial potential ΔΨm, a JC-1 fluorescence probe was used. The results showed that, Fas/FasL/Bid pathways did not play a significant role in euptox A-induced hepatocyte cell arrest and apoptosis (Fig. 5A). Also, western blot showed that euptox A treatment increased Bax expression and decreased Bcl-2 expression in a dose-dependent manner in hepatocytes. In the absence of caspase-8 activation, the levels of Bid changed inconspicuously in this study, suggesting that the activation of the mitochondrial pathway was not dependent on the activation of caspase-8 and Bid (Fig. 5B). The mRNA levels of Bax increased by 1.52-, 2.71-, and 2.89-fold in hepatocytes from mice treated with euptox A at doses of 200, 400, and 800 mg/kg, respectively, when compared to the control cells. The mRNA levels of Bcl-2 in the medium- and high-dose groups were reduced by 25.14% and 39.40%, respectively. These results revealed that euptox A administrated orally can increased the activation of Bax and decreased the activation of Bcl-2, respectively, resulting in a Bax/Bcl-2 complex that tends to activate the mitochondrial pathway (Fig. 5C).

Fig. 5

Euptox A-induced hepatocytes apoptosis was mediated by mitochondrial pathway. (A) Protein levels of Fas, FasL, and Bid measured using western blot. Euptox A treatment did not induce any changes. (B) Euptox A treatment increased protein levels Bax and decreased that of Bcl-2. β-Actin was the internal control. (C) qRT-PCR showed that euptox A increased Bax expression, but decreased that of expression Bcl-2, resulting in change of Bax/Bcl-2 ratio. (D) Euptox A treatment induced Bax translocation and cytochrome c (Cyt c) release dose-dependently. Cytosolic and mitochondrial protein fractions were collected and detected using western blot. COX 4 and β-actin were internal controls for mitochondrial and cytosolic fractions, respectively. (E) Western blot analysis revealed that euptox A treatment induced cytoplasmic release of AIF from mitochondria with subsequent dose-dependent nuclear translocation. COX 4 was internal control for mitochondrial fraction while β-actin was internal control for cytosolic and nuclear fraction. (F) ΔΨm collapse was analyzed using FCM with JC-1 staining. Data are means ± standard deviation (SD) of three independent experiments. *P < 0.05 and **P < 0.01 compared with control group. Bcl-2, B-cell lymphoma 2; Bax, Bcl-2-associated X protein; qRT-PCR, quantitative reverse transcription-polymerase reaction; COX, cyclooxygenase; ΔΨm, mitochondrial potential.

Next, the translocation of Bax and cyc c in euptox A-treated hepatocytes was analyzed. We found a dose-dependent decrease in mitochondrial Cyt c and an attendant increase in the cytosolic fraction. Accordingly, we also observed the translocation of Bax from the cytosol to the mitochondria (Fig. 5D). In addition, our results showed that euptox A treatment induced the cytoplasmic release of AIF from the mitochondria (Fig. 5E). The balance of pro- and anti-apoptotic Bcl-2 family members, such as Bax and Bcl-2, determines the change in mitochondrial potential (ΔΨm). The result also showed a significantly collapse of ΔΨm among euptox A treatments (Fig. 5F). Therefore, our results demonstrated that oral administration of euptox A induced hepatocytes apoptosis via mitochondria dysfunction pathway.

DISCUSSION

In this study, we found that orally administered euptox A directly induced cell cycle arrest and apoptosis in mouse hepatocytes via ROS accumulation, Cyt c and AIF release into the cytosol, caspase-9 and caspase-3 activation as well as PARP cleavage with observable cell morphological changes. Euptox A also led to collapse of mitochondria potential, decreased the mRNA level of Bcl-2 and increased the mRNA levels of Bax. These suggest that the liver is one of the target organs of euptox A.

At the cellular level, oral administration of euptox A-induced hepatocyte apoptosis with typical morphological characteristics such as membrane blebbing, chromatin condensation, DNA fragmentation, and phosphatidylserine externalization, which were partly in agreement with a study which also observed similar morphological changes in goats fed with A. adenophora (He et al., 2016). Other animal studies have also reported that euptox A caused lesions and impaired the histological structure of the hepatocytes (Kaushal et al., 2001b; Bhardwaj et al., 2001). These results indicated that administering euptox A orally could induce hepatocyte cell arrest and death.

Euptox A inhibited cell growth and apoptosis by ROS generation. ROS act as important multi-faceted signaling molecules that regulate numerous cellular pathways and are involved in cell fate determination (Younce and Kolattukudy, 2012; Pierre et al., 2013). The accumulation of ROS is known to lead to oxidative stress, impairment of cell function, and necrosis or apoptosis. Numerous experiments have demonstrated that ROS-mediated pathways play an important role in drug-induced cell apoptosis (Lu et al., 2012; Jang et al., 2012). In this study, the euptox A-induced ROS generation was concentration-dependent when compared with that in the control group, which is similar to the study of Sun et al. (2018), who also reported that A. adenophora treatment increases ROS levels in mouse hepatocytes. Therefore, euptox A apoptosis was modulated by the ROS-mediated pathway.

Euptox A activated the mitochondria-mediated apoptotic pathway. In our study, we observed that the euptox A-induced apoptosis involved the release of Cyt c from the mitochondria into the cytosol to form apoptosomes together with Apaf-1 and procaspase- 9. This was followed by the activation of caspase-9, -3, and the cleavage of PARP. Two central pathways are known to induce cell apoptosis: the extrinsic death receptor pathway and intrinsic mitochondria-mediated pathway (Long et al., 2004). The former is typically triggered by the ligation of death receptors such as Fas and FasL. Fas-associated proteins were recruited with a death domain, procaspase-8, and a death-inducing signaling complex were formed to activate the caspase-8. The activation of caspase-8 can cleave BH3 interacting protein (Bid, the only pro-apoptotic protein) to truncated Bid (tBid), which subsequently translocate to the mitochondria and make the mitochondrial pathway activated (Li and Yuan, 2008). The intrinsic mitochondria-mediated pathway is activated when Cytc is translocated from the mitochondria into cytosol, to form a large multiprotein complex comprising Cytc, Apaf-1 and procaspase-9. Furthermore, euptox A did not activate Fas, FasL, Bid, and the death receptor-mediated caspase-8 pathway, which indicated that the mitochondria-mediated apoptosis pathway was activated in hepatocytes treated with euptox A, and the subsequent apoptosis was induced via the Cyt c-mediated and caspase-dependent pathways. Moreover, euptox A-induced apoptosis also involved the cytoplasmic release of flavoprotein (AIF) from the mitochondria with its subsequent dose-dependent translocation to the nucleus. The mitochondria-localized AIF is involved in apoptosis induction through a caspase-independent pathway where its nuclear translocation, from the mitochondria is important for nuclear condensation and large-scale DNA fragmentation (Candé et al., 2002).

Euptox A also induced the collapse of the mitochondrial membrane potential ΔΨm in this study. Chemical agents-induced apoptosis is associated with the mitochondrion (Xu et al., 2009). The pro- and anti-apoptotic members of the Bcl-2 family are capable of regulating the mitochondrial membrane integrity. Bcl-2 protects cells against apoptosis by interacting with Bax, which blocks the release of Cyt c and AIF from the mitochondria to the cytosol. The final destiny of the cell is determined by the balance between pro- and anti-apoptotic members of the Bcl-2 family (Cory et al., 2003). In our study, deregulation of the mitochondrial integrity was associated with Bcl-2 and Bax in the euptox A -treated hepatocytes, which further confirms that activation of the mitochondria-mediated apoptosis pathway is the main underlying apoptotic mechanism in mouse hepatocyte. Moreover, disruption of the mitochondrial membrane potential is a prerequisite for triggering apoptosis by the release of apoptotic proteins from mitochondria such as Cyt c and AIF. Some studies have shown that the ΔΨm could be collapsed by Bax translocation to the mitochondria and its subsequent insertion into the outer mitochondrial membrane (Uchime et al., 2016). Euptox A induced the collapse of the ΔΨm in our study. Therefore, the translocation of Bax may be related to the ΔΨm collapse and release of mitochondrial pro-apoptotic proteins. He et al. (2015a, 2015b, 2016) reported similar results, that, mitochondria-mediated pathways played a major role in A. adenophora-induced apoptosis and death of cells. Therefore, euptox A may also cause cell death and apoptosis by the activation of mitochondria-mediated pathways.

In conclusion, our study showed that euptox A significantly caused the accumulation of ROS, downregulated Bcl-2, promoted Bax translocation into the mitochondria, and activated the mitochondria-dependent apoptotic pathway. Collectively, these actions resulted in Cyt c and AIF release into the cytosol, followed by caspase-9 and caspase-3 activation as well as PARP cleavage. Therefore, euptox A causes mouse liver cell arrest and apoptosis mainly by ROS accumulation, mitochondrial dysfunction and capases-dependent pathways (Fig. 6). This study provides new insights into the mechanisms underlying hepatocytes apoptosis caused by euptox A in mice.

Fig. 6

Schematic representation of the mechanisms by which Euptox A induces hepatocytes cell arrest and apoptosis in mice. Euptox A induces the hepatocytes cell arrest and apoptosis via three major pathways: ROS accumulation pathway, Mitochondria dysfunction pathway and Caspases 3- and 9-dependent pathways.

Malfunction of the liver may pose both economical and medical challenge to the animal rearing industry by reducing yield, quality of meat and even death, hence it is suggested that effective monitoring over grazing animals and regular weed control husbandry practices should be undertaken regularly to curb the harm this poisonous plants causes in livestock production.

ACKNOWLEDGMENTS

This research was supported by Science and Technology Support Program and Key Research & development Project of Sichuan Province (Grant No. 2015SZ0201 and 2020YFS0337).

Conflict of interest

The authors declare that there is no conflict of interest.

REFERENCES
 
© 2020 The Japanese Society of Toxicology
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