Skip to main content
PMC home page
Cell Stress & Chaperones logoLink to Cell Stress & Chaperones
. 2018 Sep 12;23(6):1295–1309. doi: 10.1007/s12192-018-0937-7

RETRACTED ARTICLE: Matrine promotes liver cancer cell apoptosis by inhibiting mitophagy and PINK1/Parkin pathways

Runjie Wei 1, Jian Cao 2, Shukun Yao 1,3,✉
PMCID: PMC6237690  PMID: 30209783

Abstract

Matrine is a natural alkaloid isolated from the root and stem of the legume plant Sophora. Its anti-proliferative and pro-apoptotic effects on several types of cancer have been well-documented. However, the role of matrine in regulating mitochondrial homeostasis, particularly mitophagy in liver cancer apoptosis, remains uncertain. The aim of our study was to explore whether matrine promotes liver cancer cell apoptosis by modifying mitophagy. HepG2 cells were used in the study and treated with different doses of matrine. Cell viability and apoptosis were determined by MTT assay, TUNEL staining, western blotting, and LDH release assay. Mitophagy was monitored by immunofluorescence assay and western blotting. Mitochondrial function was assessed by immunofluorescence assay, ELISA, and western blotting. The results of our study indicated that matrine treatment dose-dependently reduced cell viability and increased the apoptotic rate of HepG2 cells. Functional studies demonstrated that matrine treatment induced mitochondrial dysfunction and activated mitochondrial apoptosis by inhibiting protective mitophagy. Re-activation of mitophagy abolished the pro-apoptotic effects of matrine on HepG2 cells. Molecular investigations further confirmed that matrine regulated mitophagy via the PINK1/Parkin pathways. Matrine blocked the PINK1/Parkin pathways and repressed mitophagy, whereas activation of the PINK1/Parkin pathways increased mitophagy activity and promoted HepG2 cell survival in the presence of matrine. Together, our data indicated that matrine promoted HepG2 cell apoptosis through a novel mechanism that acted via inhibiting mitophagy and the PINK1/Parkin pathways. This finding provides new insight into the molecular mechanism of matrine for treating liver cancer and offers a potential target to repress liver cancer progression by modulating mitophagy and the PINK1/Parkin pathways.

Keywords: Matrine, HepG2 cell, Mitochondrial dysfunction, Mitophagy, PINK/Parkin pathways

Introduction

Hepatocellular carcinoma (HCC), the main form of liver cancer, is the sixth most common cancer worldwide. Notably, HCC accounts for ~ 5.7% of the overall cancer-caused mortality of elderly patients, and China alone contributes to ~ 55% of liver cancer deaths worldwide (Wang et al. 2017). Despite great advances in the treatment of HCC, including surgery, embolization, chemotherapy, and targeted therapy (Lee and Back 2017), the 5-year survival rate remains low, ranging from 8% in developing countries to 5% in the USA or Europe (Kalyanaraman 2017). Accordingly, it is necessary to determine effective adjuvant approaches to further reduce the progression of HCC.

Matrine, a type of alkaloid, originates from the root and stem of the legume plant Sophora (Jiang et al. 2018). In a clinical setting, matrine has long been used to treat cardiovascular disorders, inflammation, hepatitis (Guo et al. 2018; Ma et al. 2018; Zhang et al. 2018), and nervous system diseases (Ligeza et al. 2017; Tobisawa et al. 2017). Several biological processes are affected by matrine, including mitochondrial function, cellular oxidative stress, autophagy, metabolic reprogramming, calcium balance, and endoplasmic reticulum stress. Recently, the anti-tumor property of matrine is verified by several in-depth studies in many types of human cancers. Matrine promotes human colon carcinoma cell apoptosis in a caspase-3-dependent manner (Garcia-Nino et al. 2017). In human cervical cancer, matrine represses cancer mobilization and growth (Liu et al. 2017c). Moreover, additional supplementation of matrine reduces the resistance of colorectal cancer to radiation therapy (Van Nostrand et al. 2017). Similarly, matrine treatment enhances the chemotherapeutic response in bladder cancer (Merjaneh et al. 2017). Such evidence indicates that matrine may effectively influence the development and progression of different types of cancer. However, the functional role and exact mechanisms by which matrine modulates the HCC phenotype are incompletely understood.

Mitophagy, the self-repairing system for mitochondria, removes damaged mitochondria and sustains the quantity and quality of the mitochondrial mass (Zhou et al. 2018b, Zhou et al. 2018g). In response to acute and/or chronic stress stimuli, mitophagy is executed by LC3II to engulf the damaged mitochondria (Jin et al. 2018; Shi et al. 2018). Subsequently, LC3II-formed autophagosomes cooperate with lysosomes to degrade the poorly structured mitochondria (Li et al. 2018), maintaining mitochondrial homeostasis. In cardiac ischemia reperfusion, activated mitophagy removes the injured mitochondria and reduces reperfusion-mediated cardiomyocyte death (Zhou et al. 2018g). In chronic metabolic disorders, such as fatty liver disease and type 2 diabetes, upregulated mitophagy is necessary to sustain hepatocyte metabolism and mitochondrial function (Zhou et al. 2018a). In Parkinson’s disease, activated mitophagy reduces inflammation-mediated neuronal apoptosis (Garcia-Ruiz et al. 2017). These data indicate that mitophagy functions as the pro-survival system for cells under acute and chronic stimuli by preserving mitochondrial homeostasis. Because of the protective action of mitophagy on mitochondrial function and cellular viability, mitophagy is a potential target to reduce cancer progression by inducing mitochondrial dysfunction. For example, mitophagy inhibition is linked to increased gastric cancer apoptosis induced by TNFα (Nauta et al. 2017). In colorectal cancer, mitophagy suppression contributes to cancer apoptosis and migration impairment (Schock et al. 2017). Based on the above findings, we determined whether matrine regulates HCC viability by repressing mitophagy activity.

At the molecular level, mitophagy is primarily regulated by three upstream regulators, namely FUNDC1, Mfn2, and Parkin. Notably, FUNDC1-mediated mitophagy is primarily dependent on hypoxia conditions (Zhou et al. 2018e, Zhou et al. 2018,g). In addition, Mfn2-related mitophagy is activated in response to mitochondrial fission (Jovancevic et al. 2017). Interestingly, Parkin-mediated mitophagy is mainly triggered by mitochondrial damage (Nunez-Gomez et al. 2017). Poorly structured mitochondria with lower mitochondrial potential activate PINK1, and PINK1 recruits Parkin to accumulate on the surfaces of mitochondria, finally initiating mitophagy (Zhao et al. 2018). More robust data concerning the causal relationship of Parkin-related mitophagy activation and cancer survival have been provided by several studies (Huang et al. 2018). However, the influence of matrine on Parkin-mediated mitophagy in HCC has not yet been comprehensively studied. Accordingly, the aim of our study was to investigate (1) whether matrine could repress HCC survival and migration, (2) whether mitophagy was inhibited by matrine and promoted HCC mitochondrial apoptosis, and (3) whether the PINK/Parkin pathway was required for matrine-mediated mitophagy inhibition in HCC.

Methods

Cell treatment

HepG2 cells (Cell Bank of the Chinese Academy of Sciences, Shanghai, China) and the Huh7 liver cancer cell line (Cell Bank of the Chinese Academy of Sciences) were used to explore the role of matrine in the liver cancer phenotype in vitro. Analytically pure matrine, purchased from Sigma-Aldrich (Cat.No.M5319, St Louis, MO, USA), was incubated with HepG2 cells for 12 h at different doses (0–20 nM). To activate mitophagy, HepG2 cells were treated with FCCP (5 μm, Selleck Chemicals, Houston, TX, USA) for approximately 40 min at 37 °C in a 5% CO2 atmosphere. To inhibit mitophagy activity, 3-MA (10 mM, Selleck Chemicals, Houston, TX, USA) was added into the medium for approximately 2 h at 37 °C in a 5% CO2 atmosphere (Zhu et al. 2018b).

Cellular proliferation detection

Cellular proliferation was evaluated via EdU assay. Cells were seeded onto a 6-well plate, and the Cell-Light™ EdU Apollo® 567 In Vitro Imaging Kit (Thermo Fisher Scientific Inc., Waltham, MA, USA; Catalog No. A10044) was used to observe the EdU-positive cells according to the manufacturer’s instructions (Ackermann et al. 2017).

Cell viability assays

After treatment with matrine, cell viability was measured via MTT assay. For the MTT assay, cells (3 × 103 cells per well) were cultured on 96-well plates. After 24 h, the medium was replaced with new fresh medium containing MTT solution for an additional 2 h at 37 °C in a 5% CO2 atmosphere. Subsequently, the medium was removed, and DMSO was added into the medium to dissolve the MTT solution at 37 °C in a 5% CO2 atmosphere for 15 min. Finally, the plates were observed at an absorbance of 490 nm according to a previous study (Blackburn et al. 2017). Additionally, cell viability was also evaluated by analyzing the activity of caspase-3 and caspase-9 based on previous studies (Zhou et al. 2018d). The caspase-3 and caspase-9 activity kits were purchased from the Beyotime Institute of Biotechnology. The protein activity was estimated according to the manufacturer’s instructions (Brasacchio et al. 2017).

Transwell migration assay

For Transwell migration assays, the upper chambers of 24-well Transwell assay plates were seeded with 2 × 103 HepG2 cells in 200 μL serum-free medium per well. The lower chambers were filled with 600 μL medium containing 0.5% FBS (Fukumoto et al. 2018). After a 24-h incubation in a humidified incubator at 37 °C, 5% CO2, cells that had migrated to the underside of the membranes were fixed and stained with 0.1% crystal violet. After washing with distilled water, pictures of each chamber were randomly taken using a ×200 microscope field, and these images were used to quantify the total number of migrated cells (Pickard et al. 2017).

Flow cytometric analysis

Cellular ROS measurements were performed using the DHE probe. HepG2 cell was incubated with 5 μM DHE for 30 min at 37 °C in the dark. Then, cells were washed with PBS to remove free ROS probe. Subsequently, cellular ROS was observed under the Olympus IX81 microscope and quantified by fluorescence activated cell sorting (FACS) (Xiao et al. 2017).

ELISA

Cellular proteins were obtained after HepG2 cells were treated with matrine. Then, the concentration of antioxidants (GSH, GPX, and SOD) and mitochondrial respiratory complex were determined by ELISA using commercial ELISA kits, which were purchased from the Beyotime Institute of Biotechnology. The ELISA was performed according to the manufacturer’s instructions (Zhou et al. 2018f). The absorbance of the samples was observed at 450 nm using a microplate reader (Bio-Tek, Winooski, VT, USA) according to a previous study (Du et al. 2017).

Mitochondrial potential and mPTP opening rate measurement

Mitochondrial membrane potential was observed using JC-1 kit. HepG2 cells were incubated with the JC-1 probe for 30 min at 37 °C in the dark (Alghanem et al. 2017). Subsequently, cells were washed with PBS to remove the free JC-1 probe. Then, nuclei were stained with DAPI, and the mitochondrial potential was assessed under an Olympus IX81 microscope using FV10-ASW 1.7 software. The ImageJ software was used to analyze the mitochondrial potential as described previously (Feng et al. 2017). In mPTP opening assay, cells were cultured and then incubated with calcein-AM/CoCl2 staining for 25 min at 37 °C in the dark. Subsequently, the cells were washed with PBS three times to remove the free calcein-AM/CoCl2. The change in fluorescence intensity was measured by a fluorescence microscope according to the previous study (Chang et al. 2017). Then, the mPTP opening was measured.

RNA isolation and qPCR assay

The RNAsimple Total RNA Kit (Beyotime Institute of Biotechnology, China) was used to isolate the total RNA in HepG2 cells. Subsequently, cDNA was synthesized with 1 μg RNA with the Prime Script RT reagent Kit. The qPCR assay was performed using a SYBR Green PCR Kit. The primers used in the present study were as follows: cadherin, F: 5′-CTAGTGTCGAGCTTCGAAATCT-3′, R: 5′-CTGTGGTACTGTTGGACCA-3′; vimentin, F: 5′-TAGTGGTTCTTGGATATTCACT-3′, R: 5′-AGAGTTGTCATTGAATTCGG-3′; EGFR, F: 5′-GCTACCTTTGATGTTAGT-3′, R: 5′-AGAGATACCTGATAGAGTCGT-3′; BRAF: F: 5′-TCAATGACTCCTGGAAGAA-3′, R: 5′-GTGATTGATCTAATGCCTAT-3′; and GAPDH, F: 5′-AAGTTGTGFATTAGTCA-3′, R: 5′-AGAATAGTCCTATAATCA-3′. The targeted mRNA expression was normalized to GAPDH (Boone et al. 2017).

Western blotting

Protein expression was analyzed via western blotting. Primary antibodies against the following proteins were used in the present study: caspase-9 (1:1000, Cell Signaling Technology, #9504), pro-caspase-3 (1:1000, Abcam, #ab13847), cleaved caspase-3 (1:1000, Abcam, #ab49822), PARP (1:1000, Abcam, #ab32064), Bcl2 (1:1000, Cell Signaling Technology, #3498), survivin (1:1000, Cell Signaling Technology, #2808), Bax (1:1000, Cell Signaling Technology, #2772), cyclin D1 (1:1000, Abcam, #ab16663), cyclin E (1:1000, Abcam, #ab171535), cyt-c (1:1000; Abcam; #ab90529), PINK1 (1:1000, Abcam, #ab23707), cadherin (1:1000, Abcam, #ab1416), vimentin (1:1000, Abcam, #ab8978), CDK4 (1:1000, Abcam, #ab137675), ATG5 (1:1000, Cell Signaling Technology, #12994), LC3II (1:1000, Cell Signaling Technology, #3868), LC3I (1:1000, Cell Signaling Technology, #4599), Parkin (1:1000, Cell Signaling Technology, Inc.), and Vps34 (1:1000, Cell Signaling Technology, #4263). GAPDH was used as a loading control, and the targeted protein expression was normalized against GAPDH (Couto et al. 2017).

Parkin overexpression assay

The pDC315-Parkin vector was designed and purchased from Vigene Bioscience. When the cells detached from the plates, the medium supernatant was collected. Then, the viral supernatant was identified and amplified to obtain adenovirus-Parkin. Viral transductions were performed by incubating cells with recombinant Ad-Parkin in Opti-MEM media supplemented with Lipofectamine 2000 (Thermo Fisher Scientific, Inc.) according to the manufacturer’s protocol. Null vector transfection was used as the control group (Ad-ctrl). Infection was performed for 48 h at 37 °C and infection efficiency was confirmed via western blotting.

Immunofluorescence staining

Samples were observed using a Leica DM IL LED inverted fluorescence microscope (magnification ×400; Leica Microsystems, Inc.). Primary antibodies against the following proteins were used in the present study: cyt-c (1:1000; Abcam; #ab90529), PINK1 (1:1000, Abcam, #ab23707), cadherin (1:1000, Abcam, #ab1416), Tom20 (mitochondrial marker, 1:1000, Abcam, #ab186735), and LAMP1 (lysosome marker, 1:1000, Abcam, #ab24170) (Zhou et al. 2018c).

Statistical analysis

Statistical processing was performed using SPSS 16.0 (SPSS, Inc., Chicago, IL, USA). All results in the present study were analyzed by the one-way analysis of variance, followed by Tukey’s test. P < 0.05 was considered a statistically significant difference.

Data availability

All data generated or analyzed during this study are included in this published article.

Results

Matrine promotes HepG2 cell apoptosis

To establish the detailed role of matrine in the liver cancer cell phenotype in vitro, HepG2 cells and Huh7 cells were treated with different doses of matrine for approximately 12 h. Then, an MTT assay was performed to observe cell viability. Compared to the control group, matrine dose-dependently reduced the viability of HepG2 cells and Huh7 cells (Fig. 1A, B). Similarly, the cell death ratio, evaluated by the LDH release assay, was also increased in HepG2 cells and Huh7 cells in the presence of marine (Fig. 1C, D). Notably, matrine at a dose of 1 μM had no cytotoxicity on HepG2 cells, and the minimum lethal concentration of matrine is 5 μM. Accordingly, 5 μM matrine was used in subsequent studies.

Fig. 1.

Fig. 1

Fig. 1

Open in a new tab

Matrine induces cell death in HepG2 cells and Huh7 cells. A, B Cellular viability was evaluated via MTT assay. Different doses of matrine were added into the medium of HepG2 cells and Huh7 cells. C, D Cell death was determined via LDH release. HepG2 cells and Huh7 cells were treated with different concentrations of matrine. E–G The number of TUNEL-positive cells was observed and used to quantify matrine-mediated cell apoptosis in HepG2 cells and Huh7 cells. H–M Western blotting was performed to analyze the proteins caspase-3 and PARP. *P < 0.05 vs. control group. Cont control group, Mat matrine group

To quantify the cellular damage induced by matrine, TUNEL staining was used. Compared to the control group, matrine increased the number of TUNEL-positive cells (Fig. 1E, G). Caspase-3 activation is the molecular hallmark of apoptotic cells. Given this, we measured caspase-3 expression after matrine treatment. As shown in Fig. 1H–M, compared to the control group, 5 μM matrine rather than 1 μM matrine significantly increased caspase-3 expression. Additionally, caspase-3 activation was also associated with an increase in PARP, the substrate of casapse-3 (Fig. 1H–M). Together, these data suggest that matrine has toxicity in liver cancer cells when administered at a high dose. Notably, no phenotypic differences were noted between HepG2 cells and Huh7 cells with respect to cell viability or death. HepG2 cells were used in subsequent studies.

Matrine impairs HepG2 cell migration and growth

We also determined whether matrine regulated liver cancer migration and growth. First, a transwell assay was performed to evaluate the migratory capacity of matrine-treated cells. Compared to the control group, matrine treatment clearly reduced the number of migrated cells (Fig. 2A, B). These data indicated that matrine repressed HepG2 cell mobilization. To provide additional evidence for the regulatory effects of matrine on cancer migration, we measured molecular expression related to cancer metastasis. The transcription of metastatic genes, including EGFR and BRAF, was also reduced in response to matrine treatment (Fig. 2C, D). Similarly, metastatic factors, such as cadherin and vimentin, were also downregulated in the presence of matrine (Fig. 2E–G).

Fig. 2.

Fig. 2

Open in a new tab

Matrine regulates cell migration and proliferation. A, B A transwell assay was used to observe the migratory response to HepG2 cells in the presence of matrine. C, D The transcription of metastatic genes, including EGFR and BRAF, was downregulated in response to matrine treatment. E–G Western blotting was performed to analyze the protein changes of cadherin and vimentin. H, I EdU staining was applied to stain the proliferating cells at S-phase. J–L The protein expression of cyclin D1 and CDK4 was detected via western blotting. *P < 0.05 vs. control group. Cont control group, Mat matrine group

Cancer cell proliferation was also assessed with an EdU assay. As shown in Fig. 2H, I, normal liver cancer cells were ~ 45% EdU positive, and this rate was significantly reduced with matrine treatment. The decrease in the total number of EdU-positive cells indicates a decrease in the cells at S-phase, suggesting cell cycle arrest. To validate this, western blotting was performed to analyze the factors related to the cell cycle. Compared to the control, cyclin D1 and CDK4 were repressed by matrine (Fig. 2J–L). These results may indicate that the anti-proliferative effect of matrine on HepG2 cells is mediated by slowing the cell cycle in a cyclin D/E-dependent manner. Together, our data confirmed the inhibitory actions of matrine on liver cancer proliferation and migration.

Matrine activates caspase-9-dependent mitochondrial apoptosis

To determine the anti-tumor mechanism of matrine on liver cancer, we monitored mitochondrial function because the role of mitochondria goes beyond their capacity to create molecular fuel and includes the generation of reactive oxygen species, the regulation of calcium, and the activation of cell death (Zhu et al. 2018a). First, the mitochondrial membrane potential was observed using JC-1 staining under matrine stimulus. As shown in Fig. 3A, B, normal cells in the control group exhibited high mitochondrial potential as evidenced by more red fluorescence after administering the JC-1 probe. By comparison, the mitochondria in the matrine-treated cells exhibited more green fluorescence, indicating a reduction in mitochondrial potential. Moreover, the mitochondrial potential dissipation was followed by an increase in ROS production. As shown in Fig. 3C, D, compared to the control group, matrine treatment significantly promoted ROS generation, indicating cellular oxidative stress. Furthermore, due to augmented ROS production, the concentrations of antioxidant factors in HepG2 cells, including GPX, GSH, and SOD, were significantly decreased in response to matrine stress compared to the control group (Fig. 3E–G). These data indicated that matrine mediated mitochondrial dysfunction in HepG2 cells.

Fig. 3.

Fig. 3

Fig. 3

Open in a new tab

Matrine treatment causes mitochondrial dysfunction. A, B JC-1 staining was used to observe the mitochondrial membrane potential. The red fluorescence of the JC-1 probe indicates healthy mitochondria, and the green fluorescence of the JC-1 probe indicates damaged mitochondrial potential. C, D ROS production was measured via flow cytometry. E–G The concentration of antioxidants was measured via ELISA. H The ratio of mPTP opening was recorded. Matrine treatment extended the opening of the mPTP. I, J The location of cyt-c using immunofluorescence. DAPI was used to stain the nucleus. Matrine promoted cyt-c leakage from mitochondria into the nucleus. K–Q Western blotting was conducted to analyze the expression of anti- pro-apoptotic factors in matrine-treated cells. *P < 0.05 vs. control group. Cont control group, Mat matrine group

Previous studies confirmed that the redox imbalance, particularly the increase in free radicals, promotes the opening of the mitochondrial permeability transition pore (mPTP) (Daiber et al. 2017). As shown in Fig. 3H, the opening time of the mPTP was significantly extended in the matrine-treated group compared to the control group. Excessive mPTP opening provides a channel for mitochondrial pro-apoptotic factors, such as cyt-c, to enter the nucleus. The immunofluorescence assay for cyt-c showed that matrine facilitated cyt-c leakage into the nucleus compared to the control group (Fig. 3I, J). This finding was further supported by western blotting, which showed a decrease in the levels of mitochondrial cyt-c (mito cyt-c), as well as a parallel increase in cytoplasmic cyt-c (cyto cyt-c) expression (Fig. 3K–Q). In addition, western blotting also demonstrated that several key proteins related to mitochondrial apoptosis, such as Bax and casapse-9 (Fig. 3K–Q), were progressively upregulated in matrine-treated HepG2 cells. However, anti-apoptotic factors including survivin and Bcl-2 were downregulated by matrine (Fig. 3K–Q). Collectively, these data indicated that matrine controlled cell apoptosis by activating the caspase-9-dependent mitochondrial apoptotic pathway.

Matrine induces mitochondrial metabolism disorder

Mitochondria are also central to producing enough ATP to ensure cellular metabolism (Le Cras et al. 2017). Interestingly, cellular ATP production was strongly repressed by matrine treatment (Fig. 4A). At the molecular level, mitochondria generate ATP via the mitochondrial respiratory complex. However, the activity (Fig. 4B–D) and expression (Fig. 4E–H) of the mitochondrial respiratory complex were downregulated in matrine-treated cells compared to the control group. This information indicated that matrine disrupted mitochondrial energy metabolism in HepG2 cells.

Fig. 4.

Fig. 4

Open in a new tab

Matrine modulates mitochondrial energy metabolism. A Cellular ATP production was measured via ELISA. B–D The activity of the mitochondrial respiratory complex via ELISA assay. E–H Western blotting was performed to analyze the protein expression of the mitochondrial respiratory complex in matrine-treated cells. I–J Glucose uptake and lactate production were measured in the medium of matrine-treated cells. *P < 0.05 vs. control group. Cont control group, Mat matrine group

To provide additional evidence for the regulatory effects of matrine on cancer mitochondrial metabolism, we measured the residual glucose in the medium. As shown in Fig. 4I, compared to the control group, matrine treatment increased the concentration of glucose in the medium, indicative of the decline in glucose uptake. Moreover, the increase in glucose in the medium occurred along with a decrease in the levels of lactic acid in the medium supplemented with matrine (Fig. 4J), indicating the termination of cancer cell mitochondrial glucose metabolism. Together, our data demonstrated that mitochondrial energy metabolism was negatively affected by matrine.

Matrine regulates mitochondrial homeostasis by repressing mitophagy

Previous studies have reported that mitophagy is a defensive mechanism to support cancer survival in response to acute and/or chronic stress. Activated mitophagy can remove damaged mitochondria and sustains mitochondrial homeostasis. Given this, we questioned whether mitophagy is involved in matrine-mediated HepG2 cell mitochondrial damage. First, mitophagy was detected using an immunofluorescence assay. As shown in Fig. 5A, B, there was tight cooperation between mitochondria and the lysosome in the control group, suggesting mitophagy activation. However, matrine treatment interrupted the binding of mitochondria to lysosomes, indicating mitophagy inhibition (Fig. 5A, B). Furthermore, western blotting was performed to quantitatively evaluate mitophagy activity. As shown in Fig. 5C–H, compared to the control group, matrine treatment reduced LC3II expression and decreased the LC3II/LC3I ratio. Moreover, mitochondrial LC3II expression was also downregulated in response to matrine administration. Additionally, mitochondrial autophagy markers, such as ATG5 and Vps34, were also reduced in matrine-treated cells compared to the control group. Together, these data indicated that matrine suppressed the activity of mitophagy in HCC.

Fig. 5.

Fig. 5

Open in a new tab

Matrine represses protective mitophagy. A, B Mitophagy activity was monitored via immunofluorescence. Mitochondria were stained with Tom20, and lysosomes were stained with Lamp1. FCCP, an activator of mitophagy, was added into the matrine-treated cells to reactivate mitophagy. 3-MA, an inhibitor of mitophagy, was used in the control group, which was considered the negative control group. Then, the fusion of mitochondria and lysosomes indicates mitophagy, which was recorded. C–H Western blotting was performed to analyze the proteins related to mitophagy. FCCP, an activator of mitophagy, was added into the matrine-treated cells to reactivate mitophagy. 3-MA, an inhibitor of mitophagy, was used in the control group, which was the negative control group. I Caspase-9 activity was measured to reflect the role of mitophagy in mitochondrial apoptosis. J An LDH release assay was used to analyze the contribution of mitophagy to cell death. *P < 0.05 vs. control group. Cont control group, Mat matrine group

To validate whether mitophagy is involved in matrine-mediated mitochondrial damage, loss- and gain-of-function assays for mitophagy were performed. FCCP, a type of mitophagy agonist, was added to matrine-treated cells to induce mitophagy. In contrast, 3-MA, a blocker of mitophagy, was administered to the control group, which was used as the negative control group. Then, mitochondrial damage was assessed with a caspase-9 activity assay. As shown in Fig. 5I, inhibition of mitophagy occurred in the control group, and caspase-9 activity was increased, similar to the results obtained with matrine. However, matrine-increased caspase-9 activity was strongly inhibited by FCCP treatment. These data indicated that reactivation of mitophagy protected mitochondria against matrine-mediated damage in HepG2 cells. These findings were further verified via the LDH release assay (Fig. 5J), which is the early hallmark of cell death. Matrine-mediated LDH release was repressed by FCCP. Taken together, our data confirmed that matrine modulates HepG2 cell mitochondrial damage by repressing mitophagy activity.

Matrine represses mitophagy via the PINK1/Parkin pathway

Previous studies suggested that cancer mitophagy is primarily controlled by the PINK1/Parkin pathway (Yan et al. 2018; Zhao et al. 2018). Accordingly, we asked whether the PINK1/Parkin pathway is involved in matrine-related mitophagy. Western blotting assays demonstrated that the levels of both PINK1 and Parkin expression were mostly downregulated in matrine-treated cells (Fig. 6A–C) compared to the control group. Similarly, the immunofluorescence results of PINK1 and Parkin exhibited a parallel downregulation of PINK and Parkin in response to matrine treatment (Fig. 6D–F). The above data demonstrated the inhibitory action of matrine on the PINK1/Parkin pathway.

Fig. 6.

Fig. 6

Open in a new tab

Matrine regulates mitophagy via PINK1/Parkin pathways. A–C The activity of the PINK1/Parkin pathways was measured using western blotting. D–F The immunofluorescence assay for PINK1 and Parkin. G, H Adenovirus-overexpressed Parkin (Ad-Parkin) was transfected into HepG2 cells, and the overexpression efficiency was confirmed with western blotting. I, J Mitophagy was evaluated using an immunofluorescence assay. Ad-Parkin was transfected into HepG2 cells in the presence of matrine. *P < 0.05 vs. control group. Cont control group, Mat matrine group

Next, to verify whether the PINK1/Parkin pathway is necessary for matrine-modified mitophagy, we overexpressed Parkin in matrine-treated cells. The overexpression efficiency is shown in Fig. 6G, H. Then, mitophagy activity was evaluated again using immunofluorescence. As shown in Fig. 6I, J, the fusion between mitochondria and lysosomes was mostly inhibited by matrine, and this effect was significantly reversed by Parkin overexpression. Together, these data indicated that matrine suppressed mitophagy activity in HepG2 cells by inhibiting the PINK1/Parkin pathway.

PINK1/Parkin pathway is also involved in HepG2 cell migration and growth

Finally, we explored the role of the PINK1/Parkin pathway in HepG2 cell migration and growth. First, a transwell assay demonstrated that matrine-inhibited cell mobilization was reversed by Parkin overexpression (Fig. 7A, B). Additionally, the transcription of migratory factors such as cadherin and vimentin was also downregulated in response to matrine treatment and was returned to near-normal levels after the overexpression of Parkin (Fig. 7C, D). These data indicated that reactivation of the PINK1/Parkin pathway negated the inhibitory effects of matrine on HepG2 cell migration.

Fig. 7.

Fig. 7

Open in a new tab

PINK1/Parkin pathways are also involved in HepG2 cell migration and proliferation. A, B The migratory response of HepG2 cells treated with matrine or transfected with Ad-Parkin was estimated via transwell assay. C, D The transcription of metastatic factors, such as cadherin and vimentin. HepG2 cells were treated with matrine or transfected with Ad-Parkin. E–G Western blotting was used to analyze the protein expression of cyclin D1 and cyclin E. H, I EdU assay for cell proliferation. HepG2 cells were treated with matrine or transfected with Ad-Parkin; then, the number of EdU-positive cells was recorded. *P < 0.05 vs. control group. Cont control group, Mat matrine group

Regarding cellular proliferation, the expression of cyclin D1 and E was downregulated in matrine-treated cells and upregulated in Parkin-overexpressing cells (Fig. 7E–G). Moreover, the EdU assay demonstrated that matrine reduced and Parkin overexpression reversed the number of EdU-positive cells (Fig. 7H, I). Together, these data suggested that matrine regulated liver cancer growth via the PINK1/Parkin pathway.

Discussion

In the present study, our data found that matrine regulated mitochondrial function by repressing mitophagy. At the molecular levels, matrine administration interrupted the cooperation between mitochondria and lysosomes, reducing mitophagy activity. However, reactivation of mitophagy abrogated the pro-apoptotic effects of matrine on HepG2 cells. This experimental evidence substantiated the sufficiency of matrine to repress mitophagy, as well as the necessity of mitophagy for liver cancer survival. To our knowledge, this is the first investigation to identify matrine as an upstream mediator of mitophagy in liver cancer. This finding was similar to previous study. For example, in human acute lymphoblastic leukemia, matrine augments ROS production and causes mitochondrial swelling (Liu et al. 2017b). Additionally, matrine suppresses survivin signaling to promote mitochondrial apoptosis in non-small cell lung cancer (Perdiz et al. 2017). More importantly, in liver cancer, matrine activates Bid-mediated mitochondrial apoptosis in vivo and in vitro (Liu et al. 2017a). These data confirm that mitochondria are the potential target of matrine for regulating cancer viability.

Notably, previous studies found that mitophagy activation provides a pro-survival advantage for several types of cancer. In gastric cancer, although TNFα-based therapy reduces cancer viability and increases the apoptotic index, it also activates mitophagy (Zhang et al. 2016). Augmented mitophagy degrades the damaged mitochondria and prevents the amplification of apoptotic signaling. Additionally, in human cervical cancer, activated mitophagy promotes cancer survival and is involved in a tumor cell’s ability to escape programmed cell death (Sarkar et al. 2017). Consistent with this finding, activated mitophagy also promotes migration and adhesion in endometriosis and liver cancer (Shi et al. 2018; Zhao et al. 2018). Given these observations, previous studies proposed that mitophagy may play a key role in tumor escape and tumor therapy resistance. Our data also provide evidence to further validate this hypothesis. Based on these data, the inhibition of mitophagy is of utmost importance when designing anti-cancer therapies in combination with chemotherapy. More importantly, our data provide an easy and effective method to block mitophagy, namely matrine. Administration of matrine inhibited mitophagy activity and further promoted mitochondrial malfunction as evidenced by increased caspase-9 activity and an augmented apoptotic ratio. Therefore, our results in HepG2 cells lay the foundation for a detailed study of the molecular mechanisms of liver cancer apoptosis management and regulation. Furthermore, we provide new data that show that mitophagy inhibition is responsible for the tumor-suppressive action of matrine.

At the molecular level, mitophagy is primarily regulated by FUNDC1, Mfn2, and Parkin. FUNDC1, the mitophagy receptor expressed on the outer membrane of mitochondria, is preferentially activated by hypoxia stimulus. In ischemia-challenged tissues, such as the heart, FUNDC1 is activated and protects mitochondria against stress-evoked apoptosis (Zhou et al. 2018e, Zhou et al. 2018g). Mfn2-mediated mitophagy is primarily triggered along with mitochondrial fission. In gastric cancer and pancreatic cancer, Mfn2-mediated mitophagy promotes cancer cell survival by sustaining mitochondrial function and structure. Parkin is mainly activated in response to a reduction in mitochondrial potential (Fuhrmann and Brune 2017). The mitochondrial potential collapse causes PINK1 upregulation, and this enhances Parkin expression. Elevated Parkin levels promote the fusion between mitochondria and lysosomes, contributing to the removal of damaged mitochondria (Rossello and Yellon 2017). In this present study, we found that PINK1/Parkin-mediated mitophagy was inhibited by matrine. However, overexpression of PINK1 could reactivate mitophagy in matrine-treated cells and reduce caspase-9 activation and cellular apoptosis. This comprehensively demonstrated that the PINK1/Parkin pathway is required for matrine-related mitophagy. However, more studies are needed to determine whether matrine has a role in regulating mitophagy via FUNDC1 and/or Mfn2.

Together, our data indicated that matrine can reduce HepG2 cell viability, the migratory response, and proliferation in vitro. Mechanistically, matrine suppresses mitophagy activity by downregulating the PINK1/Parkin pathway, evoking mitochondrial dysfunction and activating caspase-9-dependent mitochondrial apoptosis in HepG2 cells. These findings provide new insights into the mechanisms of and new strategies for regulating the progression of liver cancer.

Authors’ contribution

CJ and SKY conceived the research; RJW, CJ, and SKY performed the experiments; all authors participated in discussing and revising the manuscript.

Funding

This study was funded in full by the Leap-forward Development Program for Beijing Biopharmaceutical Industry (G20), grant number Z171100001717008.

References

  1. Ackermann M, Kim YO, Wagner WL, Schuppan D, Valenzuela CD, Mentzer SJ, Kreuz S, Stiller D, Wollin L, Konerding MA. Effects of nintedanib on the microvascular architecture in a lung fibrosis model. Angiogenesis. 2017;20:359–372. doi: 10.1007/s10456-017-9543-z. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Alghanem AF, Wilkinson EL, Emmett MS, Aljasir MA, Holmes K, Rothermel BA, Simms VA, Heath VL, Cross MJ. RCAN1.4 regulates VEGFR-2 internalisation, cell polarity and migration in human microvascular endothelial cells. Angiogenesis. 2017;20:341–358. doi: 10.1007/s10456-017-9542-0. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Blackburn NJR, Vulesevic B, McNeill B, Cimenci CE, Ahmadi A, Gonzalez-Gomez M, Ostojic A, Zhong Z, Brownlee M, Beisswenger PJ, Milne RW, Suuronen EJ. Methylglyoxal-derived advanced glycation end products contribute to negative cardiac remodeling and dysfunction post-myocardial infarction. Basic Res Cardiol. 2017;112:57. doi: 10.1007/s00395-017-0646-x. [DOI] [PubMed] [Google Scholar]
  4. Boone CHT, Grove RA, Adamcova D, Seravalli J, Adamec J. Oxidative stress, metabolomics profiling, and mechanism of local anesthetic induced cell death in yeast. Redox Biol. 2017;12:139–149. doi: 10.1016/j.redox.2017.01.025. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Brasacchio D, Alsop AE, Noori T, Lufti M, Iyer S, Simpson KJ, Bird PI, Kluck RM, Johnstone RW, Trapani JA. Epigenetic control of mitochondrial cell death through PACS1-mediated regulation of BAX/BAK oligomerization. Cell Death Differ. 2017;24:961–970. doi: 10.1038/cdd.2016.119. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Chang SH, Yeh YH, Lee JL, Hsu YJ, Kuo CT, Chen WJ. Transforming growth factor-beta-mediated CD44/STAT3 signaling contributes to the development of atrial fibrosis and fibrillation. Basic Res Cardiol. 2017;112:58. doi: 10.1007/s00395-017-0647-9. [DOI] [PubMed] [Google Scholar]
  7. Couto JA, Ayturk UM, Konczyk DJ, Goss JA, Huang AY, Hann S, Reeve JL, Liang MG, Bischoff J, Warman ML, Greene AK. A somatic GNA11 mutation is associated with extremity capillary malformation and overgrowth. Angiogenesis. 2017;20:303–306. doi: 10.1007/s10456-016-9538-1. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Daiber A, Oelze M, Steven S, Kroller-Schon S, Munzel T. Taking up the cudgels for the traditional reactive oxygen and nitrogen species detection assays and their use in the cardiovascular system. Redox Biol. 2017;12:35–49. doi: 10.1016/j.redox.2017.02.001. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Du GQ, et al. Targeted myocardial delivery of GDF11 gene rejuvenates the aged mouse heart and enhances myocardial regeneration after ischemia-reperfusion injury. Basic Res Cardiol. 2017;112:7. doi: 10.1007/s00395-016-0593-y. [DOI] [PubMed] [Google Scholar]
  10. Feng Dayun, Wang Bao, Wang Lei, Abraham Neeta, Tao Kai, Huang Lu, Shi Wei, Dong Yushu, Qu Yan. Pre-ischemia melatonin treatment alleviated acute neuronal injury after ischemic stroke by inhibiting endoplasmic reticulum stress-dependent autophagy via PERK and IRE1 signalings. Journal of Pineal Research. 2017;62(3):e12395. doi: 10.1111/jpi.12395. [DOI] [PubMed] [Google Scholar]
  11. Fuhrmann DC, Brune B. Mitochondrial composition and function under the control of hypoxia. Redox Biol. 2017;12:208–215. doi: 10.1016/j.redox.2017.02.012. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Fukumoto M, Kondo K, Uni K, Ishiguro T, Hayashi M, Ueda S, Mori I, Niimi K, Tashiro F, Miyazaki S, Miyazaki JI, Inagaki S, Furuyama T. Tip-cell behavior is regulated by transcription factor FoxO1 under hypoxic conditions in developing mouse retinas. Angiogenesis. 2018;21:203–214. doi: 10.1007/s10456-017-9588-z. [DOI] [PubMed] [Google Scholar]
  13. Garcia-Nino WR, et al. Cardioprotective kinase signaling to subsarcolemmal and interfibrillar mitochondria is mediated by caveolar structures. Basic Res Cardiol. 2017;112:15. doi: 10.1007/s00395-017-0607-4. [DOI] [PubMed] [Google Scholar]
  14. Garcia-Ruiz JM, et al. Bloodless reperfusion with the oxygen carrier HBOC-201 in acute myocardial infarction: a novel platform for cardioprotective probes delivery. Basic Res Cardiol. 2017;112:17. doi: 10.1007/s00395-017-0605-6. [DOI] [PubMed] [Google Scholar]
  15. Guo S, Gao C, Xiao W, Zhang J, Qu Y, Li J, Ye F (2018) Matrine protects cardiomyocytes from ischemia/reperfusion injury by regulating HSP70 expression via activation of the JAK2/STAT3 pathway. Shock:1. 10.1097/SHK.0000000000001108 [DOI] [PubMed]
  16. Huang CY, Kuo WW, Ho TJ, Chiang SF, Pai PY, Lin JY, Lin DY, Kuo CH, Huang CY. Rab9-dependent autophagy is required for the IGF-IIR triggering mitophagy to eliminate damaged mitochondria. J Cell Physiol. 2018;233:7080–7091. doi: 10.1002/jcp.26346. [DOI] [PubMed] [Google Scholar]
  17. Jiang JH, Pi J, Jin H, Yang F, Cai JY. Chinese herb medicine matrine induce apoptosis in human esophageal squamous cancer KYSE-150 cells through increasing reactive oxygen species and inhibiting mitochondrial function. Pathol Res Pract. 2018;214:691–699. doi: 10.1016/j.prp.2018.03.015. [DOI] [PubMed] [Google Scholar]
  18. Jin Q, Li R, Hu N, Xin T, Zhu P, Hu S, Ma S, Zhu H, Ren J, Zhou H. DUSP1 alleviates cardiac ischemia/reperfusion injury by suppressing the Mff-required mitochondrial fission and Bnip3-related mitophagy via the JNK pathways. Redox Biol. 2018;14:576–587. doi: 10.1016/j.redox.2017.11.004. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Jovancevic N, Dendorfer A, Matzkies M, Kovarova M, Heckmann JC, Osterloh M, Boehm M, Weber L, Nguemo F, Semmler J, Hescheler J, Milting H, Schleicher E, Gelis L, Hatt H. Medium-chain fatty acids modulate myocardial function via a cardiac odorant receptor. Basic Res Cardiol. 2017;112:13. doi: 10.1007/s00395-017-0600-y. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Kalyanaraman B. Teaching the basics of cancer metabolism: developing antitumor strategies by exploiting the differences between normal and cancer cell metabolism. Redox Biol. 2017;12:833–842. doi: 10.1016/j.redox.2017.04.018. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Le Cras Timothy D., Mobberley-Schuman Paula S., Broering Mary, Fei Lin, Trenor Cameron C., Adams Denise M. Angiopoietins as serum biomarkers for lymphatic anomalies. Angiogenesis. 2016;20(1):163–173. doi: 10.1007/s10456-016-9537-2. [DOI] [PubMed] [Google Scholar]
  22. Lee Hyoung Yool, Back Kyoungwhan. Melatonin is required for H2O2- and NO-mediated defense signaling through MAPKKK3 and OXI1 inArabidopsis thaliana. Journal of Pineal Research. 2016;62(2):e12379. doi: 10.1111/jpi.12379. [DOI] [PubMed] [Google Scholar]
  23. Li R, Xin T, Li D, Wang C, Zhu H, Zhou H. Therapeutic effect of Sirtuin 3 on ameliorating nonalcoholic fatty liver disease: the role of the ERK-CREB pathway and Bnip3-mediated mitophagy. Redox Biol. 2018;18:229–243. doi: 10.1016/j.redox.2018.07.011. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Ligeza J, Marona P, Gach N, Lipert B, Miekus K, Wilk W, Jaszczynski J, Stelmach A, Loboda A, Dulak J, Branicki W, Rys J, Jura J. MCPIP1 contributes to clear cell renal cell carcinomas development. Angiogenesis. 2017;20:325–340. doi: 10.1007/s10456-017-9540-2. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Liu D, Zeng X, Li X, Mehta JL, Wang X. Role of NLRP3 inflammasome in the pathogenesis of cardiovascular diseases. Basic Res Cardiol. 2017;113:5. doi: 10.1007/s00395-017-0663-9. [DOI] [PubMed] [Google Scholar]
  26. Liu Zhenjiang, Gan Lu, Luo Dan, Sun Chao. Melatonin promotes circadian rhythm-induced proliferation through Clock/histone deacetylase 3/c-Myc interaction in mouse adipose tissue. Journal of Pineal Research. 2017;62(4):e12383. doi: 10.1111/jpi.12383. [DOI] [PubMed] [Google Scholar]
  27. Liu Zhenjiang, Gan Lu, Xu Yatao, Luo Dan, Ren Qian, Wu Song, Sun Chao. Melatonin alleviates inflammasome-induced pyroptosis through inhibiting NF-κB/GSDMD signal in mice adipose tissue. Journal of Pineal Research. 2017;63(1):e12414. doi: 10.1111/jpi.12414. [DOI] [PubMed] [Google Scholar]
  28. Ma J, Ma S, Yin C, Wu H. Matrine reduces susceptibility to postinfarct atrial fibrillation in rats due to antifibrotic properties. J Cardiovasc Electrophysiol. 2018;29:616–627. doi: 10.1111/jce.13448. [DOI] [PubMed] [Google Scholar]
  29. Merjaneh M, Langlois A, Larochelle S, Cloutier CB, Ricard-Blum S, Moulin VJ. Pro-angiogenic capacities of microvesicles produced by skin wound myofibroblasts. Angiogenesis. 2017;20:385–398. doi: 10.1007/s10456-017-9554-9. [DOI] [PubMed] [Google Scholar]
  30. Nauta TD, van den Broek M, Gibbs S, van der Pouw-Kraan TC, Oudejans CB, van Hinsbergh VW, Koolwijk P. Identification of HIF-2alpha-regulated genes that play a role in human microvascular endothelial sprouting during prolonged hypoxia in vitro. Angiogenesis. 2017;20:39–54. doi: 10.1007/s10456-016-9527-4. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Nunez-Gomez E, Pericacho M, Ollauri-Ibanez C, Bernabeu C, Lopez-Novoa JM. The role of endoglin in post-ischemic revascularization. Angiogenesis. 2017;20:1–24. doi: 10.1007/s10456-016-9535-4. [DOI] [PubMed] [Google Scholar]
  32. Perdiz D, Lorin S, Leroy-Gori I, Pous C. Stress-induced hyperacetylation of microtubule enhances mitochondrial fission and modulates the phosphorylation of Drp1 at (616)Ser. Cell Signal. 2017;39:32–43. doi: 10.1016/j.cellsig.2017.07.020. [DOI] [PubMed] [Google Scholar]
  33. Pickard JM, Burke N, Davidson SM, Yellon DM. Intrinsic cardiac ganglia and acetylcholine are important in the mechanism of ischaemic preconditioning. Basic Res Cardiol. 2017;112:11. doi: 10.1007/s00395-017-0601-x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Rossello X, Yellon DM. The RISK pathway and beyond. Basic Res Cardiol. 2017;113:2. doi: 10.1007/s00395-017-0662-x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Sarkar C, Ganju RK, Pompili VJ, Chakroborty D. Enhanced peripheral dopamine impairs post-ischemic healing by suppressing angiotensin receptor type 1 expression in endothelial cells and inhibiting angiogenesis. Angiogenesis. 2017;20:97–107. doi: 10.1007/s10456-016-9531-8. [DOI] [PubMed] [Google Scholar]
  36. Schock SN, Chandra NV, Sun Y, Irie T, Kitagawa Y, Gotoh B, Coscoy L, Winoto A. Induction of necroptotic cell death by viral activation of the RIG-I or STING pathway. Cell Death Differ. 2017;24:615–625. doi: 10.1038/cdd.2016.153. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Shi C, Cai Y, Li Y, Li Y, Hu N, Ma S, Hu S, Zhu P, Wang W, Zhou H. Yap promotes hepatocellular carcinoma metastasis and mobilization via governing cofilin/F-actin/lamellipodium axis by regulation of JNK/Bnip3/SERCA/CaMKII pathways. Redox Biol. 2018;14:59–71. doi: 10.1016/j.redox.2017.08.013. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Tobisawa T, Yano T, Tanno M, Miki T, Kuno A, Kimura Y, Ishikawa S, Kouzu H, Nishizawa K, Yoshida H, Miura T. Insufficient activation of Akt upon reperfusion because of its novel modification by reduced PP2A-B55alpha contributes to enlargement of infarct size by chronic kidney disease. Basic Res Cardiol. 2017;112:31. doi: 10.1007/s00395-017-0621-6. [DOI] [PubMed] [Google Scholar]
  39. Van Nostrand JL, Bowen ME, Vogel H, Barna M, Attardi LD. The p53 family members have distinct roles during mammalian embryonic development. Cell Death Differ. 2017;24:575–579. doi: 10.1038/cdd.2016.128. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Wang K, Gan TY, Li N, Liu CY, Zhou LY, Gao JN, Chen C, Yan KW, Ponnusamy M, Zhang YH, Li PF. Circular RNA mediates cardiomyocyte death via miRNA-dependent upregulation of MTP18 expression. Cell Death Differ. 2017;24:1111–1120. doi: 10.1038/cdd.2017.61. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Xiao L, Xu X, Zhang F, Wang M, Xu Y, Tang D, Wang J, Qin Y, Liu Y, Tang C, He L, Greka A, Zhou Z, Liu F, Dong Z, Sun L. The mitochondria-targeted antioxidant MitoQ ameliorated tubular injury mediated by mitophagy in diabetic kidney disease via Nrf2/PINK1. Redox Biol. 2017;11:297–311. doi: 10.1016/j.redox.2016.12.022. [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Yan H, Xiao F, Zou J, Qiu C, Sun W, Gu M, Zhang L. NR4A1-induced increase in the sensitivity of a human gastric cancer line to TNFalpha-mediated apoptosis is associated with the inhibition of JNK/Parkin-dependent mitophagy. Int J Oncol. 2018;52:367–378. doi: 10.3892/ijo.2017.4216. [DOI] [PMC free article] [PubMed] [Google Scholar] [Retracted]
  43. Zhang Y, Zhou H, Wu W, Shi C, Hu S, Yin T, Ma Q, Han T, Zhang Y, Tian F, Chen Y. Liraglutide protects cardiac microvascular endothelial cells against hypoxia/reoxygenation injury through the suppression of the SR-Ca(2+)-XO-ROS axis via activation of the GLP-1R/PI3K/Akt/survivin pathways. Free Radic Biol Med. 2016;95:278–292. doi: 10.1016/j.freeradbiomed.2016.03.035. [DOI] [PubMed] [Google Scholar]
  44. Zhang Y, Yang X, Qiu C, Liu F, Liu P, Liu Z. Matrine suppresses AGE-induced HAEC injury by inhibiting ROS-mediated NRLP3 inflammasome activation. Eur J Pharmacol. 2018;822:207–211. doi: 10.1016/j.ejphar.2018.01.029. [DOI] [PubMed] [Google Scholar]
  45. Zhao Q, Ye M, Yang W, Wang M, Li M, Gu C, Zhao L, Zhang Z, Han W, Fan W, Meng Y. Effect of Mst1 on endometriosis apoptosis and migration: role of Drp1-related mitochondrial fission and Parkin-required mitophagy. Cell Physiol Biochem. 2018;45:1172–1190. doi: 10.1159/000487450. [DOI] [PubMed] [Google Scholar]
  46. Zhou Hao, Du Wenjuan, Li Ye, Shi Chen, Hu Nan, Ma Sai, Wang Weihu, Ren Jun. Effects of melatonin on fatty liver disease: The role of NR4A1/DNA-PKcs/p53 pathway, mitochondrial fission, and mitophagy. Journal of Pineal Research. 2017;64(1):e12450. doi: 10.1111/jpi.12450. [DOI] [PubMed] [Google Scholar]
  47. Zhou Hao, Ma Qiang, Zhu Pingjun, Ren Jun, Reiter Russel J., Chen Yundai. Protective role of melatonin in cardiac ischemia-reperfusion injury: From pathogenesis to targeted therapy. Journal of Pineal Research. 2018;64(3):e12471. doi: 10.1111/jpi.12471. [DOI] [PubMed] [Google Scholar]
  48. Zhou H, Shi C, Hu S, Zhu H, Ren J, Chen Y. BI1 is associated with microvascular protection in cardiac ischemia reperfusion injury via repressing Syk-Nox2-Drp1-mitochondrial fission pathways. Angiogenesis. 2018;21:599–615. doi: 10.1007/s10456-018-9611-z. [DOI] [PubMed] [Google Scholar]
  49. Zhou H, Wang J, Zhu P, Hu S, Ren J. Ripk3 regulates cardiac microvascular reperfusion injury: the role of IP3R-dependent calcium overload, XO-mediated oxidative stress and F-action/filopodia-based cellular migration. Cell Signal. 2018;45:12–22. doi: 10.1016/j.cellsig.2018.01.020. [DOI] [PubMed] [Google Scholar]
  50. Zhou H, Wang J, Zhu P, Zhu H, Toan S, Hu S, Ren J, Chen Y. NR4A1 aggravates the cardiac microvascular ischemia reperfusion injury through suppressing FUNDC1-mediated mitophagy and promoting Mff-required mitochondrial fission by CK2alpha. Basic Res Cardiol. 2018;113:23. doi: 10.1007/s00395-018-0682-1. [DOI] [PubMed] [Google Scholar]
  51. Zhou H, Yue Y, Wang J, Ma Q, Chen Y. Melatonin therapy for diabetic cardiomyopathy: a mechanism involving Syk-mitochondrial complex I-SERCA pathway. Cell Signal. 2018;47:88–100. doi: 10.1016/j.cellsig.2018.03.012. [DOI] [PubMed] [Google Scholar]
  52. Zhou H, Zhu P, Wang J, Zhu H, Ren J, Chen Y. Pathogenesis of cardiac ischemia reperfusion injury is associated with CK2alpha-disturbed mitochondrial homeostasis via suppression of FUNDC1-related mitophagy. Cell Death Differ. 2018;25:1080–1093. doi: 10.1038/s41418-018-0086-7. [DOI] [PMC free article] [PubMed] [Google Scholar]
  53. Zhu H, Jin Q, Li Y, Ma Q, Wang J, Li D, Zhou H, Chen Y. Melatonin protected cardiac microvascular endothelial cells against oxidative stress injury via suppression of IP3R-[Ca(2+)]c/VDAC-[Ca(2+)]m axis by activation of MAPK/ERK signaling pathway. Cell Stress Chaperones. 2018;23:101–113. doi: 10.1007/s12192-017-0827-4. [DOI] [PMC free article] [PubMed] [Google Scholar]
  54. Zhu P, Hu S, Jin Q, Li D, Tian F, Toan S, Li Y, Zhou H, Chen Y. Ripk3 promotes ER stress-induced necroptosis in cardiac IR injury: a mechanism involving calcium overload/XO/ROS/mPTP pathway. Redox Biol. 2018;16:157–168. doi: 10.1016/j.redox.2018.02.019. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Data Availability Statement

All data generated or analyzed during this study are included in this published article.

Articles from Cell Stress & Chaperones are provided here courtesy of Elsevier

RESOURCES