TAZ inhibition promotes IL-2-induced apoptosis of hepatocellular carcinoma cells by activating the JNK/F-actin/mitochondrial fission pathway

The HepG2 cell line (ATCC, Cat# HB-8065™) and Hep3B cell line (ATCC^® HB-8064™) were purchased from ATCC. The Huh7 cell line was purchased from National Infrastructure of Cell Line Resource (Beijing, China). These cells were maintained in L-DMEM with 10% serum and 1% streptomycin/penicillin. To induce cell damage, IL-2 (0–20 ng/mL) was used for 12-h. To activate mitochondrial fission in HepG2 cells, cells were pre-treated with FCCP (5 μM) for approximately 30 min. To inhibit mitochondrial fission, cells were treated with mitochondrial division inhibitor-1 (Mdivi-1; 10 mM; Sigma-Aldrich; Merck KGaA) for 2 h. To suppress the activity of the c-Jun N-terminal kinase (JNK) pathway, HepG2 cell was treated with SP600125 (SP, 10 µM) (Selleck Chemicals, Houston, TX, USA) for 2 h [30]. Conversely, to activate the JNK pathway, Anisomycin (Ani, 10 µM) (Selleck Chemicals, Houston, TX, USA) was added to the medium for 2 h at room temperature [31].

Mitochondrial function was evaluated via detecting the mitochondrial membrane potential and measuring the ROS levels. HepG2 cells were incubated with the JC-1 probe for 30 min at 37 °C in the dark [32]. 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 [33]. 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) [34].

Cellular glutathione (GSH), glutathione peroxidase (GPx) and SOD were measured by ELISA according to the manufacturer’s instructions [35]. Cellular lactate production was measured using a lactate assay kit (#K607-100; BioVision, Milpitas, CA, USA) as described in a previous study. The glucose uptake rate was detected using a glucose absorption assay kit (#K606-100; BioVision) [36].

Caspase-3 and caspase-9 activity were detected according to previous study [37]. The caspase-3/9 activity kits (cat. nos. C1158 and C1115; Beyotime Institute of Biotechnology) were used based on the manufacture’s instruction [38]. To measure ATP production, a luciferase-based ATP assay kit (Celltiter-Glo Luminescent Cell Viability assay; Promega, Madison, WI, USA; Catalog No. A22066) was used according to the manufacturer’s instructions [39]. ATP production was measured using a microplate reader (Epoch 2; BioTek Instruments, Inc.).

An siRNA targeting the TAZ gene (5′AAAGACUCCUUUCUUGCUUGU-3′) and a control siRNA (5′UUCUCCGAACGUGUCACGUTT-3′) were obtained from Sangon Biotech (Shanghai, China) [40]. HepG2 cells were seeded in 6-well plates in 2 mL growth medium per well without antibiotics and were cultured to 70% confluence. Prior to transfection, 5 μL Lipofectamine 2000 (Invitrogen, Ltd., USA) and 1 nmol siRNA were added to separate 250 μL aliquots of Opti-MEM (Invitrogen, Ltd., USA). These solutions were mixed and incubated for 15 min at room temperature [41]. Then, the two solutions were combined and mixed for another 15 min to allow complexes to form. Complexes were then added to each well of cells, and plates were gently agitated. After incubation for 16 h at 37 °C in a humidified incubator, total protein was extracted from cells, and gene knockdown was evaluated by western blotting [42].

HepG2 cells were detached by scraping and were then washed and lysed on ice. Samples (60 μg protein each) were resolved by electrophoresis and transferred to polyvinylidene fluoride membranes. After blocking, membranes were incubated with primary antibodies at 4 °C overnight. This was followed by a 1-h incubation with a horseradish peroxidase-conjugated secondary antibody [43]. Membranes were visualized using Chemiluminescence HRP Substrate (Millipore, Billerica, USA) and a Luminescent Imaging Workstation (Tanon, Shanghai, China) [44]. β-Actin and GAPDH were used as an internal control. The primary antibodies used in the present study were as follows: caspase-9 (1:1000, Cell Signaling Technology, #9504), TAZ (1:1000, Abcam, #ab224239), pro-caspase-3 (1:1000, Abcam, #ab13847), cleaved caspase-3 (1:1000, Abcam, #ab49822), Cyclin E (1:1000, Abcam, #ab171535), Bcl-2 (1:1000, Cell Signaling Technology, #3498), CXCR7 (1:1000, Abcam, #ab38089), Bax (1:1000, Cell Signaling Technology, #2772), c-IAP (1:1000, Cell Signaling Technology, #4952), complex III subunit core (CIII-core2, 1:1000, Invitrogen, #459220), survivin (1:1000, Cell Signaling Technology, #2808), Cyt-c (1:1000; Abcam; #ab90529), Drp1 (1:1000, Abcam, #ab56788), complex II (CII-30, 1:1000, Abcam, #ab110410), Fis1 (1:1000, Abcam, #ab71498), F-actin (1:1000, Abcam, #ab205), complex IV subunit II (CIV-II, 1:1000, Abcam, #ab110268), complex I subunit NDUFB8 (CI-20, 1:1000, Abcam, #ab110242), Mff (1:1000, Cell Signaling Technology, #86668), Tom20 (1:1000, Abcam, #ab186735), CXCR4 (1:1000, Abcam, #ab1670), Cyclin D (1:1000, Abcam, #ab134175), t-JNK (1:1000; Cell Signaling Technology, #4672), p-JNK (1:1000; Cell Signaling Technology, #9251).

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 [45]. 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 [46].

Cell death was measured by TUNEL staining, PI staining and MTT assays. In the TUNEL assays, cells (HepG2 cells, Huh7 cells and Hep3B cells) were fixed in paraformaldehyde at room temperature for 15 min. Then, cells were washed three times in cold PBS and treated with 0.05% Triton X-100 for 15 min on ice [47]. Subsequently, a TUNEL kit (Roche Apoptosis Detection Kit, Roche, Mannheim, Germany) was used to stain cells, as described in a previous study [48]. For the PI staining assays, HepG2 cells were washed three times with cold PBS, and PI (1 mM) was added to the medium. The cells were cultured in the dark at 37 °C, 5% CO₂, for 15 min. Finally, at least 10 fields were randomly chosen under a digital microscope, and the apoptotic cells in these fields were quantified [49]. MTT assays were carried out according to a previous report [50]. Briefly, HepG2 cells were seeded into 96-well plates. Cells were treated with MTT (2 mg/mL; Sigma-Aldrich) for 4 h at 37 °C. Then, cells were washed with PBS to remove free MTT, and DMSO was added to the wells. Finally, the OD of each well was recorded, and cellular viability was calculated as described in a previous study [51].

HepG2 cells were fixed in 4% paraformaldehyde at room temperature for 30 min and then permeabilized using 0.05% Triton X-100 at 4 °C for 4 h [52]. Then, cells were washed with cold PBS to remove free Triton X-100. Subsequently, samples were incubated with primary antibodies at 4 °C overnight. The primary antibodies used in the present study were as follows: Cyt-c (1:500; Abcam; #ab90529), F-actin (1:500, Abcam, #ab205), p-JNK (1:500; Cell Signaling Technology, #9251) and Tom20 (1:500, Abcam, #ab186735). Tom20 was used to observe mitochondrial fission according to previous study [53]. Immunofluorescence was assessed under an Olympus IX81 microscope using FV10-ASW 1.7 software [54].

All experiments were performed in triplicate. Normal distribution of data was firstly analyzed and then statistical analysis was performed using one-way analysis of variance (ANOVA) followed by Bonferroni’s multiple comparison test. P values < 0.05 were considered statistically significant.

In in vitro experiments, different concentrations of IL-2 were administered to HepG2 cell to induce cell damage. Compared to the control group, IL-2 treatment reduced HepG2 cell viability in a dose-dependent manner, as assessed by MTT assays (Figs. 1a, 2). Notably, cell viability was lowest in the 10 ng/mL IL-2 group (Fig. 1a). Beyond this concentration, increases in the IL-2 dose could not further reduce HepG2 cell viability. Based on this result, 10 ng/mL IL-2 was used for subsequent experiments.

Next, the expression of TAZ was evaluated to verify the functional role of TAZ in IL-2-mediated HepG2 cell death. As shown in Fig. 1b, c, TAZ protein expression significantly increased in response to IL-2 treatment. Because TAZ has been recognized as an essential cancer promoter, we asked whether TAZ deletion would further promote IL-2-mediated HepG2 cell death. To answer this question, a TAZ siRNA was transfected into HepG2 cell in the presence of IL-2. Analysis of the transfection efficiency was assessed via western blotting (Fig. 1d, e). To more specifically test the effects of TAZ deletion on IL-2-mediated HepG2 cell death, TUNEL assays were performed. As shown in Fig. 1f, g, IL-2 supplementation augmented the number of TUNEL-positive cells; this effect was further enhanced by TAZ knockdown. This finding was further supported using other liver cancer cell lines such as Huh7 cells (Fig. 1h, i) and Hep3B cells (Fig. 2j, k).

Subsequently, to identify the signaling pathways responsible for HepG2 cell death, we examined the caspase-9-related mitochondrial apoptotic pathways. Compared to the control group, IL-2 treatment significantly upregulated the expression of pro-apoptotic proteins, such as Bax, Bad, caspase-9 and caspase-3 (Fig. 2l–s). In comparison, the levels of anti-apoptotic proteins were markedly decreased in response to IL-2 treatment (Fig. 2l–s). Interestingly, this tendency was further enhanced by TAZ deletion. Taken together, these data indicate that IL-2 treatment promotes HepG2 cell death by activating caspase-9-dependent mitochondrial apoptosis and that this effect is further enhanced by TAZ inhibition.

Next, experiments were performed to investigate the influence of TAZ on HepG2 cell migration and proliferation in the presence of IL-2. Through Transwell assays, we found that IL-2 treatment reduced HepG2 cell migration compared to the control group (Fig. 3a, b), suggesting that IL-2 impairs HepG2 cell motility. Interestingly, TAZ siRNA transfection further attenuated cell migration (Fig. 3a, b). At the molecular level, cancer cell migration is dependent on the expression of chemotactic molecules such as CXCR4 and CXCR7. The protein levels of CXCR4 and CXCR7 were drastically downregulated in response to IL-2 stress, and this effect was enhanced by TAZ deletion (Fig. 3c–e).

Subsequently, cellular proliferation was examined via EdU staining. The results shown in Fig. 3f, g demonstrated that the number of EdU⁺ cells was significantly reduced following IL-2 treatment, indicating that IL-2 can inhibit HepG2 cell growth to some extent. Interestingly, the anti-proliferative effect of IL-2 on HepG2 cell was further enhanced by TAZ inhibition (Fig. 3f–h), suggesting that the loss of TAZ could improve the anti-proliferative effects of IL-2 on HepG2 cell. This observation was further verified by detecting the expression of Cyclin D1 and Cyclin E, regulatory molecules that promote the G1/S transition in cancer cells. IL-2 treatment decreased the protein expression of Cyclin D1 and Cyclin E; this effect was augmented by TAZ deletion (Fig. 3h–j). Taken together, these results indicate that TAZ deficiency could sensitize HepG2 cell to IL-2-mediated proliferation arrest and migration inhibition.

Cellular migration and proliferation are highly reliant on the cell’s supply of ATP, which is produced by the mitochondria [44, 55]. Additionally, mitochondrial energy deficiency is closely associated with mitochondrial apoptosis [56, 57]. Therefore, we questioned whether IL-2 and TAZ deletion were also involved in cellular energy metabolism disorders. We first measured cellular ATP production by ELISA and found that IL-2 treatment repressed ATP generation (Additional file 1: Fig. S1A). Interestingly, this effect was further enhanced by TAZ knockdown via siRNA transfection (Additional file 1: Fig. S1A). Considering that ATP is primarily produced by mitochondrial respiration, we hypothesized that the drop in ATP production was due to decreased expression of the mitochondrial respiratory complex. Through western blot analysis, we demonstrated that the expression of mitochondrial respiratory complex components was noticeably downregulated in response to IL-2 treatment; this trend was augmented by siRNA-mediated TAZ deletion (Additional file 1: Fig. S1B–F).

Subsequently, more solid evidence was obtained by measuring glucose and lactate concentrations in the medium of HepG2 cell transfected with TAZ siRNA in the presence of IL-2. As illustrated in Additional file 1: Fig. S1G, H, when compared to the control group, IL-2 treatment increased residual glucose and decreased lactate in the medium. These data suggested that IL-2 administration represses HepG2 cell glucose intake and lactate production. Interestingly, TAZ deletion further increased the residual glucose concentration and thus limited lactate generation (Additional file 1: Fig. S1G, H). Taken together, our results demonstrate the functional importance of IL-2 treatment and TAZ inhibition in suppressing mitochondrial energy metabolism in HepG2 cell.

Mitochondrial fission has been reported to be an early sign of mitochondrial dysfunction [58, 59]. Based on the data described above, we asked whether mitochondrial fission was required for IL-2-induced mitochondrial damage. As shown in Fig. 4a, IL-2 treatment caused most strip-shaped mitochondria divisions to several round fragmentations; this effect was augmented by TAZ siRNA transfection. These data hinted that mitochondrial fission could be activated by IL-2 and exacerbated by TAZ inhibition. We next measured the average length of mitochondrial fissions and found that mitochondrial length decreased to ~ 40% and ~ 80% of control levels in the IL-2-treated and TAZ-deleted cells, respectively, suggesting that TAZ deletion further aggravates IL-2-induced mitochondrial fission (Fig. 4b). Western blotting was then performed to analyze the expression of proteins related to mitochondrial fission. Compared to the control group, IL-2-treated HepG2 cell exhibited elevated expression of proteins related to mitochondrial fission, including Drp1, Fis1 and Mff (Fig. 4c–f). These effects were strongly augmented by TAZ deletion. These data confirmed that TAZ deficiency exacerbated mitochondrial fission in HepG2 cell in the presence of IL-2. Finally, to explore whether mitochondrial fission is responsible for the apoptosis of HepG2 cell, we blocked mitochondrial fission in TAZ-depleted cells using Mdivi-1. We then measured caspase-3 activity by ELISA. As shown in Fig. 4g, caspase-3 activity was increased by IL-2 administration or TAZ siRNA transfection compared to the control group. Conversely, inhibition of mitochondrial fission via Mdivi-1 abolished IL-2 and TAZ silencing-induced caspase-3 activation (Fig. 4g). Cell death was further assessed via PI staining. As shown in Fig. 4h, i, IL-2 increased the number of PI-positive cells, and this effect was further enhanced by TAZ deletion. However, blockade of mitochondrial fission using Mdivi-1 inhibited the pro-apoptotic effects of TAZ deletion. Collectively, through loss- and gain-of-function assays, we confirm that mitochondrial fission is activated by IL-2 and TAZ deficiency and is an upstream trigger of HepG2 cell apoptosis.

Next, we asked how mitochondrial fission promotes the apoptosis of HepG2 cell. Previous findings in a myocardial reperfusion injury model indicated that mitochondrial fission is able to initiate caspase-9-dependent mitochondrial apoptosis by inducing oxidative stress and promoting pro-apoptotic factor leakage. Based on these findings, we first examined cellular oxidative stress by flow cytometry. As shown in Fig. 5a, b, IL-2 treatment elevated ROS generation compared to the control group; this effect was augmented by TAZ inhibition. Interestingly, inhibition of mitochondrial fission repressed ROS production (Fig. 5a, b). We also found that TAZ deletion-induced ROS overproduction was closely accompanied by a drop in cellular antioxidant factors such as SOD, GSH and GPx (Fig. 5c–e). Interestingly, inhibition of mitochondrial fission preserved the expression of SOD, GSH and GPx in TAZ-deleted cells, suggesting that cellular redox imbalance is triggered by TAZ deletion and can be abrogated by inhibition of mitochondrial fission.

In response to IL-2-induced mitochondrial oxidative damage, the mitochondria released cyt-c into the nucleus. This effect was exacerbated by TAZ deletion in a mitochondrial fission-dependent manner (Fig. 5f, g). Cyt-c, a pro-apoptotic factor primarily located in mitochondria, can activate caspase-9 and subsequently elevate caspase-3 activity [60], causing cells to undergo mitochondrial apoptosis. Therefore, we measured caspase-9 activity. Our results demonstrated that TAZ deficiency increased (Fig. 5h) and Mdivi-1 reduced caspase-9 activity in the presence of IL-2. Taken together, these findings indicate that IL-2-induced mitochondrial apoptosis in HepG2 cell is attributable to activated mitochondrial fission, which is enhanced by TAZ deletion.

Next, experiments were conducted to explore the upstream mediators responsible for TAZ-mediated mitochondrial fission in the presence of IL-2. In light of the contributions of the JNK/F-actin pathway to mitochondrial fission in gastric cancer and cervical cancer [1, 24], we investigated whether TAZ could regulate mitochondrial fission via the JNK/F-actin pathway. Western blotting for JNK and F-actin demonstrated that the JNK/F-actin pathway was activated by IL-2 treatment (Fig. 6a–c), as evidenced by increased p-JNK and F-actin expression. Notably, loss of TAZ further upregulated p-JNK and F-actin expression in the presence of IL-2. This finding was further verified via immunofluorescence staining for p-JNK (Fig. 6d, e) and F-actin (Fig. 6f, g). These findings indicated that the JNK/F-actin pathway is activated by IL-2 and TAZ deletion. To establish the causal role of the JNK/F-actin pathway in TAZ-induced mitochondrial fission, a JNK inhibitor (SP600125) and an agonist (Anisomycin; Ani) were used to inhibit and re-activate, respectively, the JNK pathway in the presence of IL-2. Then, mitochondrial fission was measured again. Similar to the above results, IL-2-activated mitochondrial fission was further enhanced by TAZ deletion (Fig. 6h, i). However, SP600125 significantly repressed TAZ-mediated mitochondrial fission in the presence of IL-2. By contrast, Ani application activated mitochondrial fission in the control group (Fig. 6h, i). Overall, these data confirmed that TAZ deletion initiates mitochondrial fission by activating the JNK/F-actin pathway.

Finally, we wanted to investigate whether the JNK/F-actin pathway is also implicated in mitochondrial dysfunction and HepG2 cell apoptosis. To address this question, we measured mitochondrial potential. Consistent with the data described above, our results demonstrated that IL-2 treatment reduced mitochondrial potential and that this effect was enhanced by TAZ silencing (Fig. 7a, b). Interestingly, JNK inhibition (via SP600125) prevented the reduction in mitochondrial potential induced by IL-2 treatment and TAZ deletion (Fig. 7a, b). Moreover, cellular ROS production was elevated by IL-2 treatment and/or TAZ deletion and was reduced by SP600125 (Fig. 7c, d). Finally, caspase-9 activity, the key hallmark of mitochondrial apoptosis, was increased in response to IL-2 treatment and/or TAZ deletion and was reduced by SP600125 (Fig. 7e). Taken together, these data illustrate that TAZ-mediated mitochondrial apoptosis in HepG2 cell is highly reliant on the activity of the JNK/F-actin pathway.