TNFα promotes glioblastoma A172 cell mitochondrial apoptosis via augmenting mitochondrial fission and repression of MAPK-ERK-YAP signaling pathways.

BACKGROUND AND
OBJECTIVE: The present study was designed to explore the roles of mitochondrial fission and MAPK-ERK-YAP signaling pathways and to determine their mutual relationship in TNFα-mediated glioblastoma mitochondrial apoptosis.
MATERIALS AND METHODS: Cellular viability was measured via TUNEL staining, MTT assays, and Western blot. Immunofluorescence was performed to observe mitochondrial fission. YAP overexpression assays were conducted to observe the regulatory mechanisms of MAPK-ERK-YAP signaling pathways in mitochondrial fission and glioblastoma mitochondrial apoptosis.
RESULTS: The results in our present study indicated that TNFα treatment dose dependently increased the apoptotic rate of glioblastoma cells. Functional studies confirmed that TNFα-induced glioblastoma apoptosis was attributable to increased mitochondrial fission. Excessive mitochondrial fission promoted mitochondrial dysfunction, as evidenced by decreased mitochondrial potential, repressed ATP metabolism, elevated ROS synthesis, and downregulated antioxidant factors. In addition, the fragmented mitochondria liberated cyt-c into the cytoplasm/nucleus where it activated a caspase-9-involved mitochondrial apoptosis pathway. Furthermore, our data identified MAPK-ERK-YAP signaling pathways as the primary molecular mechanisms by which TNFα modulated mitochondrial fission and glioblastoma apoptosis. Reactivation of MAPK-ERK-YAP signaling pathways via overexpression of YAP neutralized the cytotoxicity of TNFα, attenuated mitochondrial fission, and favored glioblastoma cell survival.
CONCLUSION: Overall, our data highlight that TNFα-mediated glioblastoma apoptosis stems from increased mitochondrial fission and inactive MAPK-ERK-YAP signaling pathways, which provide potential targets for new therapies against glioblastoma.

Introduction

Although glioblastoma multiforme (GBM) is a rare tumor whose incidence is less than 3.19/100,000 in the population globally, its poor prognosis with a median survival of 15 months and inevitable recurrence after a median survival time of 32–36 weeks make it a heavy burden on the health care system. Unfortunately, little is known about the etiology of GBM, although several risk factors have been proposed, such as age, exposure to radiation, and family history. Notably, excessive hyperplasia of glial cells is the primary pathogenesis of GBM.1 Accordingly, several approaches have been attempted to induce the death of glial cells, especially TNFα-based therapy.

A gene delivery strategy to induce TNFα overexpression has been attempted to increase the apoptotic index of glioblastoma cells.2 The effectiveness of the TNFα-based therapy is later validated by several clinical studies.3 Ample in vivo and in vitro evidence potentially implies that TNFα considerably augments the apoptosis of glioblastoma cells.4 This information indicates that TNFα-based therapy is a promising tool for the treatment of glioblastoma. However, the molecular mechanisms of TNFα involved in glioblastoma cell death have not been fully described.

Mitochondria control an array of subcellular functions, such as energy metabolism, ROS production, cell proliferation, calcium balance, and cell death.5,6 Previous studies have provided molecular insight into the mitochondrial etiology in GBM and have identified mitochondria as a potentially therapeutic target to modulate the growth of gliomas.7 In addition, TNFα-based therapy has been linked to mitochondrial dysfunction in GBM. For example, TNFα promotes mitochondrial oxidative stress via the JNK–NF–κB pathways.8 Some researchers have demonstrated that TNFα induces mitochondrial apoptosis via increasing tBid stability.9 In addition, other studies suggest that Bnip3-related mitochondrial necrotic death is activated by TNFα.10 This information indicates that TNFα potentially targets mitochondria in glioblastoma cells. Recently, mitochondrial fission has been thought to be the early feature of mitochondrial abnormalities and to promote the death of several kinds of tumors, such as breast cancer,11 ovarian cancer,12 pancreatic cancer,13 and bladder cancer.14 TNFα has been found to be associated with Drp1 activation during the inflammation-mediated cardiomyocyte injury.15 However, no studies have investigated the role of mitochondrial fission in TNFα-treated glioblastoma cells. In the present study, we ask whether mitochondrial fission is required for TNFα-mediated mitochondrial apoptosis in glioblastoma cells.

The MAPK–ERK signaling pathway has been found to be the upstream inhibitor of mitochondrial fission. In liver cancer, defective ERK signaling upregulates FAK expression and the latter promotes mitochondrial fission.16 Moreover, in neuroblastoma N2a cells, increased ERK signaling inhibits mitochondrial fission and sustains cellular viability.17 Furthermore, in-depth studies have indicated that ERK modulates mitochondrial fission via YAP. Increased YAP suppresses mitochondrial fission in human rectal cancer,18 cerebral ischemia-reperfusion injury,19 and dendritic cells.20 These findings uncover the critical role played by ERK–YAP signaling in inhibiting mitochondrial fission. Considering that ERK is also the classical antiapoptotic signal for cancer,21 we ask whether TNFα handles mitochondrial fission via repressing the MAPK–ERK–YAP signaling pathways. Altogether, the aim of our study was to investigate the therapeutic effects of TNFα on glioblastoma cells and determine its influence on mitochondrial fission and the MAPK–ERK–YAP signaling pathways.

Materials and methods

Cell culture and treatment

Human glioblastoma cell line A172 (ATCC® CRL 1620™) was purchased from American Type Culture Collection. These cells were cultured with L-DMEM supplemented with 10% FBS (Biowest, Mexico City, Mexico, USA) and 1% penicillin/streptomycin in a humidified atmosphere with 5% CO2 at 37°C. Different doses of TNFα were added to the medium of A172 cells for 12 hours to induce cell damage (0–20 ng/mL). This concentration of TNFα was chosen according to a previous study.22 Cells were exposed to 10 mM mitochondrial division inhibitor-1 (Mdivi-1; Sigma-Aldrich Co., St Louis, MO, USA; EMD Millipore, Billerica, MA, USA) to inhibit the activity of mitochondrial fission. In contrast, to activate mitochondrial fission, 5 µm FCCP (Selleck Chemicals, Houston, TX, USA) was pretreated for 40 minutes at 37°C in a 5% CO2 atmosphere.23

MTT assay, TUNEL staining, and LDH release assay

The cell viability was determined by MTT assays (Sigma-Aldrich Co.). Briefly, cells were seeded onto 96-well plates, and then 20 µL of MTT at a concentration of 5 mg/mL was added to the medium. The plates were placed for 4 hours in the dark at 37°C and 5% CO2. After that, the medium was removed and 100 µL of dimethyl sulfoxide (DMSO) was added into the medium for 15 minutes in the dark at 37°C and 5% CO2. Then, the samples were observed at a wavelength of 570 nm. The relative cell viability was recorded as a ratio to that of the control group. Apoptotic cells were quantified using a one-step TUNEL kit (Beyotime Institute of Biotechnology, Haimen, China) according to the manufacturer’s instructions.24 Cells were seeded onto the 12-well plates and incubated with fluorescein–dUTP (Beyotime Institute of Biotechnology) for 30 minutes at 37°C in a 5% CO2 atmosphere. After being labeled with DAPI, the cells were observed using a laser confocal microscope (TcS SP5; Leica Microsystems, Inc., Buffalo Grove, IL, USA). LDH was released into the medium when cellular membranes ruptured. To evaluate the levels of LDH in the medium, an LDH Release Detection kit (Beyotime Institute of Biotechnology) was used according to manufacturer’s protocol. Cells treated with PBS were used as the control group for MTT assay, LDH release assay, and TUNEL staining.

Measurement of mitochondrial membrane potential and mitochondrial permeability transition pore (mPTP) opening rate

Mitochondrial potential was evaluated using 5,5′,6,6′-tetrachloro-1,1′,3,3′-tetraethyl-benzimidazolylcarbocyanine chloride (JC-1) staining. Cells were seeded onto 12-well plates. After washing with PBS three times, the cells were treated with the JC-1 probe for 30 minutes in the dark at 37°C and 5% CO2. Then, cells were washed with PBS three times to remove the free JC-1. After replaced with fresh DMEM, the cells were observed using a laser confocal microscope (TcS SP5). At least 30 cells were randomly chosen.25 To measure the mPTP opening, cells were loaded with PBS containing 25 nM tetramethylrhodamine, methyl ester (TMRM, T668; Thermo Fisher Scientific, Waltham, MA, USA). After 30 minutes, cells were washed with PBS again to remove the free TMRM. Then, samples were observed at a wavelength of 480 nm using a microplate reader (Epoch 2; BioTek Instruments, Inc., Tokyo, Japan). Cells treated with PBS were used as the control group for MTT assay and TUNEL staining.

Western blots

Total proteins were extracted using RIPA Lysis Buffer (Cat. No: P0013E; Beyotime, Beijing, China). After that, proteins were rapidly centrifuged (20,000 rpm) for 10 min at 4°C to pellet cell debris. Supernatant was collected and quantified using an Enhanced BCA Protein Assay Kit (Beyotime, Cat. No: P0009). Then, proteins (45–60 µg) were loaded in a 10%–15% SDS-PAGE gel and transferred to PVDF membranes (Bio-Rad, Hercules, CA, USA). Subsequently, membranes were blocked with 5% skim milk for 45 minutes at room temperature. After washing with tris buffered saline with tween 20 (TBST) three times at room temperature, the membranes were incubated with the primary antibodies at 4°C overnight.26 After washing, horseradish peroxidase-conjugated secondary antibodies were incubated with membranes for 50 minutes at room temperature. Then, the bands were observed using an ECL Prime Western Blotting Detection Reagent (GE Healthcare, Buckinghamshire, UK). The primary antibodies used in the present study are described as follows: p-ERK (1:1,000, #ab176660; Abcam, Cambridge, MA, USA), t-ERK (1:1,000, #ab54230; Abcam), Yap (1:1,000; #14074; Cell Signaling Technology, Danvers, MA, USA), complex III subunit core (CIII-core2, 1:1,000, #459220; Invitrogen, Merck KGaA, Darmstadt, Germany), complex II (CII-30, 1:1,000, #ab110410; Abcam), complex IV subunit II (CIV-II, 1:1,000, #ab110268; Abcam), Drp1 (1:1,000, #ab56788; Abcam), Fis1 (1:1,000, #ab71498; Abcam), Opa1 (1:1,000, #ab42364; Abcam), Mfn2 (1:1,000, #ab57602; Abcam), Mff (1:1,000, #86668; Cell Signaling Technology), Tom20 (1:1,000, #ab186735; Abcam), Bcl-2 (1:1,000, #3498; Cell Signaling Technology), Bax (1:1,000, #2772; Cell Signaling Technology), Bcl-2 (1:1,000, #3498; Cell Signaling Technology), Bad (1:1,000, #ab90435; Abcam), and x-IAP (1:1,000, #ab28151; Abcam).

Immunofluorescence

Cells were seeded onto poly-d-lysine-coated coverslips. Then, methanol-free 4% paraformaldehyde was used to fix cells for 15 minutes at room temperature. Subsequently, samples were blocked with 5% goat serum at room temperature for 45 minutes. After washing with TBST, samples were incubated with primary antibody at 4°C overnight. The primary antibodies used in the present study were as follows: p-ERK (1:1,000, #ab176660), Yap (1:1,000, #14074), Tom20 (1:1,000, #ab186735), and cyt-c (1:1,000, #ab90529; Abcam). Subsequently, samples were rinsed three times with TBST for 15 minutes and followed by incubation with secondary antibody for 45 minutes at room temperature; after rinsing three times for 5 minutes using TBST, the samples were labeled with DAPI to tag the nuclei. Cells were observed using a laser confocal microscope (TcS SP5).27

Transfection

The pDC315–YAP vector was designed and purchased from Vigene Biosciences, Inc. (Rockville, MD, USA). Then, the plasmid was transfected in 293 T cells using Lipofectamine 2000®. After 48 hours, the supernatant was collected and amplified to obtain adenovirus-YAP (Ad-YAP). Subsequently, A172 cells were infected with Ad-YAP using Lipofectamine 2000® for 6 hours at 37°C and 5% CO2. Western blot was performed to observe the overexpression efficiency.28

ROS and antioxidant factors quantification

ROS generation was quantified using flow cytometry. Cells were seeded onto the 12-well plates. After washing with PBS, dihydroethidium (DHE) staining was added into the medium and the cells were incubated with the DHE probe for 30 minutes in the dark at 37°C and 5% CO2. Then, PBS was used to wash cells to remove the free DHE probe. Subsequently, 0.25% trypsin was applied to collect the cell. Flow cytometry analysis was performed using the BD FACSCanto II cytometer (BD, San Diego, CA, USA). Analysis of the data was performed using FACSDiva software (BD). Besides, the ROS production was also observed using a laser confocal microscope (TcS SP5). The concentration of cellular antioxidant factors such as GSH (Glutathione Reductase Assay Kit, Cat. No: S0055; Beyotime), SOD (Total Superoxide Dismutase Assay Kit, Cat. No: S0101; Beyotime), and GPX (Cellular Glutathione Peroxidase Assay Kit, Cat. No: S0056; Beyotime) was measured via ELISA according to the manufacture’s guidelines.29 Cells treated with PBS were used as the control group for MTT assay and TUNEL staining.

Caspase-3/9 activities and Trypan Blue staining

Caspase-3 and caspase-9 activities were measured using the Caspase-3 Activity Assay Kit (Cat. No: C1115; Beyotime) and Caspase-9 Activity Assay Kit (Cat. No: C1158; Beyotime) following the manufacturer’s instructions.30 Briefly, cells were seeded onto 96-well plates. Then, 100 µL of caspase-3 and caspase-9 reagents were added to each sample. After incubation for 30 minutes in the dark at 37°C and 5% CO2, the samples were measured at a wavelength of 570 nm using the microplate reader (Epoch 2). The relative caspase activity was recorded as the ratio to that of the control group. Trypan Blue staining was conducted using 0.4% Trypan Blue probe, which was treated with cells for 2 minutes. Then, the number of Trypan Blue-positive cells was calculated by counting at least three random separate fields.

Cellular ATP level detection

Cellular ATP levels were measured using the Enhanced ATP Assay Kit (Cat. No: S0027; Beyotime) following the supplier’s specifications.31 Briefly, cells were seeded onto 96-well plates at a density of 1×104/well. Subsequently, 100 µL of staining solution (Enhanced ATP Assay Kit, Cat. No: S0027) was added to each well and incubated with the cells for 4 hours in the dark at 37°C and 5% CO2. The relative ATP production was recorded using a microplate reader (Epoch 2) at a wavelength of 570 nm.

Statistical data analyses

The results are presented as the mean±standard error (SE) from at least three independent experiments using SPSS 16.0 software (SPSS Inc., Chicago, IL, USA). One-way ANOVAs were carried out for comparisons between control and treated groups. Pairwise comparisons were made by post hoc Tukey’s test. Differences were considered as significant at P<0.05.

Results

TNFα dose-dependently promotes glioblastoma cell apoptosis in vitro

In the present study, glioblastoma cells were incubated with different doses of TNFα for 12 hours. Then, the cellular viability was measured via MTT assay. As shown in Figure 1A, with increasing concentrations of TNFα, the viability of glioblastoma cells decreased progressively. Reduction in cellular viability may result from cell death. To analyze the cellular death rate, the LDH release assay was performed. Compared to the control group, TNFα dose dependently elevated the content of LDH in the medium (Figure 1B), indicating that TNFα promoted glioblastoma cell death. This finding was further supported via Trypan Blue and TUNEL staining, which exhibited an increased number of Trypan-positive (Figure 1C and D) and TUNEL-positive cells (Figure 1E and F) in the presence of TNFα stress. At the molecular levels, cell death is primarily executed via caspase-3 activation, which cleaves DNA into fragments. Accordingly, caspase-3 activity was measured, and the results shown in Figure 1G illustrate that caspase-3 activity was drastically increased with the rise in TNFα. Altogether, our data indicate that TNFα treatment dose dependently promotes glioblastoma cell apoptosis. Notably, no significant difference was observed between the control group and the 1 ng/mL TNFα group. The minimal proapoptotic dose of TNFα is 5 ng/mL; accordingly, 5 ng/mL TNFα was used in the following studies.

Figure 1

TNFα promotes glioblastoma cell apoptosis in a dose-dependent manner.

Notes: (A) Different doses of TNFα were added into the media of glioblastoma cells, and then, the cellular viability was measured via MTT assay. (B) An LDH release assay was performed to detect cell death. (C and D) Trypan Blue staining for cell death. The number of Trypan Blue-positive cell was recorded. (E and F) A TUNEL assay was used to determine the rate of apoptosis. The number of TUNEL-positive cells was measured. (G) Caspase-3 activity was measured to determine the activation level of the caspase-3 protein. *P<0.05 vs the control group. The 0 ng/mL TNFα was used as the control group.

Mitochondrial fission is activated by TNFα treatment

Several thorough studies from many laboratories have reported that mitochondrial fission is an early event leading to cell death.13 In the present study, we explored the functional role of TNFα in mitochondrial fission. The immunofluorescence assay in Figure 2A demonstrated that mitochondria are highly connected networks. However, after TNFα treatment, mitochondria become small, roundish fragments that are characteristic of mitochondrial fission. To quantify mitochondrial fission, we measured the average length of mitochondria with or without TNFα treatment. In the control group, the length of mitochondria was ~8.9 µm. Interestingly, TNFα treatment (5 ng/mL) reduced the mitochondrial length to ~2.3 µm (Figure 2B). This information indicated that mitochondrial fission was activated by TNFα treatment in glioblastoma cells. To provide additional evidence for the role of TNFα in triggering mitochondrial division, Mdivi-1, an antagonist of mitochondrial division was used. Meanwhile, a mitochondrial fission agonist was administered to the normal glioblastoma cells to activate mitochondrial fission, which was used as the positive control group. Then, Western blot was performed to analyze alterations in protein levels related to mitochondrial fission.32,33 When compared to the control group, TNFα treatment increased the levels of Drp1, Mff, and Fis1, the key elements in executing mitochondrial fission (Figure 2C–H). In contrast, inhibitors of mitochondrial fission such as Mfn2 and Opa1 were significantly downregulated in response to TNFα treatment (Figure 2C–H). This effect of TNFα was similar to the action of FCCP, which caused an imbalance between mitochondrial fission factors (Figure 2C–H). Interestingly, Mdivi-1 could abrogate the promotive effects of TNFα on mitochondrial fission-related proteins (Figure 2C–H). Altogether, our data confirm that TNFα promotes mitochondrial fission activation in glioblastoma cells.

Figure 2

TNFα activates mitochondrial fission in glioblastoma cells.

Notes: (A) An immunofluorescence assay for mitochondria using mitochondrial specific antibody Tom20. (B) The average length of mitochondria was measured, which was used to analyze the extent of mitochondrial fission. (C–H) Western blot was performed to analyze protein expression of mitochondrial fission-related factors. To perform the loss- and gain-of-function assays for mitochondrial fission, Mdivi-1, a pharmacological antagonist was used in TNFα-treated cells to inhibit the activation of mitochondrial fission. FCCP, an agonist for mitochondrial fission, was administered to the control group, which was used as the positive control group. Drp1, Fis1, and Mff are mitochondrial fission activators whose levels were upregulated in response to TNFα treatment and downregulated by Mdivi-1. By contrast, Mfn2 and Opa1 are the mitochondrial fission inhibitors whose expression levels were repressed by TNFα stress and were increased by Mdivi-1. *P<0.05.

Abbreviation: Mdivi-1, mitochondrial division inhibitor-1.

TNFα-mediated mitochondrial fission promotes mitochondrial dysfunction

Abnormal mitochondrial fission plays a decisive role in mediating mitochondrial dysfunction. To verify whether TNFα induces mitochondrial damage in glioblastoma cells via mitochondrial fission, mitochondrial function was measured. First, cell ROS production was determined via flow cytometry. When compared to the control group, TNFα treatment significantly increased ROS production in glioblastoma cells (Figure 3A and B), and this effect was similar to the results obtained via administering FCCP (Figure 3A and B). Interestingly, TNFα-mediated ROS production was mostly negated by Mdivi-1 (Figure 3A and B). Because of the cellular ROS outburst, the concentration of cellular antioxidants such as GSH, SOD, and GPX was obviously reduced in response to TNFα treatment (Figure 3C–E). However, Mdivi-1 could reverse the levels of GSH, SOD, and GPX (Figure 3C–E). The abovementioned data suggested that TNFα-mediated mitochondrial oxidative stress via mitochondrial fission.

Figure 3

TNFα-initiated mitochondrial fission contributes to glioblastoma mitochondrial injury.

Notes: (A and B) ROS levels were measured using DHE probe, and flow cytometry was performed to quantify the content of cell ROS. To perform the loss- and gain-of-function assays for mitochondrial fission, Mdivi-1, a pharmacological antagonist was used in TNFα-treated cells to inhibit mitochondrial fission activation. FCCP, an agonist for mitochondrial fission was administered to the control group, which was set as the positive control group. (C–E) ELISAs for cellular antioxidants such as GSH, SOD, and GPX. (F) Cellular ATP production was measured using a commercial kit. (G–J) Western blot was conducted to analyze the protein expression of the mitochondrial respiratory complex. *P<0.05.

Abbreviations: DHE, dihydroethidium; Mdivi-1, mitochondrial division inhibitor-1; CIII-core2, complex III subunit core; CII-30, complex II; CIV-II, complex IV subunit II.

The core function of mitochondria is to produce ATP, which is required for cellular metabolism. Interestingly, the content of ATP was significantly reduced in the presence of TNFα treatment (Figure 3F), similar to the results obtained after administering FCCP. However, Mdivi-1 supplementation abrogated the inhibitory effects of TNFα on ATP production (Figure 3F). At the molecular level, mitochondria produce ATP via the mitochondrial respiratory complex. Notably, the protein expression of mitochondrial respiratory complex was significantly repressed by TNFα (Figure 3G–J), and this effect was negated by Mdivi-1. This information indicated that TNFα-mediated mitochondrial fission reduced the levels of the mitochondrial respiratory complex. Altogether, our data confirm that TNFα treatment causes an obvious mitochondrial malfunction that occurs, at least in part, through mitochondrial fission.

TNFα-mediated mitochondrial fission activates a caspase-9-related mitochondrial apoptotic pathway

Damaged mitochondria initiate cellular apoptosis programs.34 Based on this, we explored whether TNFα-mediated mitochondrial fission accounted for glioblastoma cell apoptosis. An early molecular feature of mitochondrial apoptosis is a drop in the mitochondrial potential. As shown in Figure 4A and B, compared to the control group, TNFα markedly reduced the mitochondrial potential as evidenced by decreased red fluorescence and increased green fluorescence. Interestingly, this alteration could be abrogated by Mdivi-1 (Figure 4A and B), suggesting that inhibition of mitochondrial fission protected the mitochondrial potential in the presence of TNFα treatment. The collapse of the mitochondrial potential indicates hyperpermeability of the mitochondrial outer membrane.35 Accordingly, we evaluated the opening rate of the mPTP. Compared to the control group, TNFα treatment increased the opening rate of mPTP (Figure 4C), similar to the results obtained via administration of FCCP. However, Mdivi-1 supplementation significantly blocked the mPTP opening (Figure 4C). Excessive opening of mPTP could facilitate mitochondrial proapoptotic cyt-c translocation into the cytoplasm where cyt-c interacts with and activates caspase-9.36 The immunofluorescence assay for cyt-c indicated that TNFα treatment promoted cyt-c migration to the nucleus (Figure 4D and E), and this effect was negated by Mdivi-1. In response to the cyc-c liberation, the activity of caspase-9 was increased in TNFα-treated cells, whereas Mdivi-1 treatment prevented caspase-9 activation (Figure 4F).

Figure 4

TNFα-activated caspase-9 apoptosis is regulated by mitochondrial fission.

Notes: (A and B) Mitochondrial potential was observed via JC-1 staining. Red fluorescence of the JC-1 probe indicates the normal mitochondrial potential, whereas green fluorescence of the JC-1 probe means a defective mitochondrial potential. The red-to-green fluorescence intensity was recorded to quantify the mitochondrial potential. To perform the loss- and gain-of-function assays for mitochondrial fission, Mdivi-1, a pharmacological antagonist was used in TNFα-treated cells to inhibit mitochondrial fission activation. FCCP, an agonist for mitochondrial fission, was administered to the control group, which was set as the positive control group. (C) mPTP opening was measured in response to TNFα stress and/or mitochondrial fission inhibition. (D and E) Immunofluorescence assay for mitochondrial cyt-c translocation into nucleus. Nuclei were labeled by DAPI, and the colocalization of cyt-c and DAPI indicates the migration of mitochondrial cyt-c into nucleus. The relative expression of nuclear cyt-c was monitored. (F) Caspase-9 activity was determined via ELISA. TNFα-mediated caspase-9 activation could be abrogated by Mdivi-1. (G–K) Western blot was performed to analyze the alterations in mitochondrial apoptotic proteins. Bax and Bad are proapoptotic proteins, whereas Bcl-2 and x-IAP are antiapoptotic proteins. TNFα regulated the balance of proapoptotic and antiapoptotic proteins via mitochondrial fission. *P<0.05.

Abbreviation: Mdivi-1, mitochondrial division inhibitor-1.

In addition, we also found that the expression levels of mitochondrial proapoptotic proteins such as Bax and Bad were significantly upregulated in TNFα-treated cells (Figure 4G–K), similar to the results obtained via adding FCCP. By comparison, the levels of antiapoptotic proteins such as Bcl-2 and x-IAP were downregulated in response to TNFα stress (Figure 4G–K). Interestingly, Mdivi-1 treatment reversed the levels of antiapoptotic factors. These results indicated that mitochondrial apoptosis was activated by TNFα via mitochondrial fission.

TNFα modulates mitochondrial fission via MAPK–ERK–YAP signaling pathways

Subsequently, we explored the molecular mechanism by which TNFα controlled mitochondrial fission in glioblastoma cells. Previous studies have suggested that mitochondrial fission is negatively regulated by the MAPK–ERK–YAP signaling pathways.37,38 In the present study, we noted abundant p-ERK expression in the control group via Western blot (Figure 5A–C). However, TNFα treatment significantly suppressed p-ERK expression (Figure 5A–C), indicative of ERK inactivation in response to TNFα stimulus. Moreover, the TNFα-mediated decrease in p-ERK expression was closely associated with a drop in YAP expression (Figure 5A–C), suggesting that TNFα inactivated MAPK–ERK–YAP pathways in glioblastoma cells. PD98059 was used to inhibit ERK activity, which was used to mimic the inhibitory effects of TNFα on ERK pathways. This finding was further supported via immunofluorescence (Figure 5D and E). The fluorescence intensities of p-ERK and YAP in the TNFα-treated cells decreased bŷ65% and ~50% of the control levels, respectively. To demonstrate whether MAPK–ERK–YAP signaling pathways were required for TNFα-mediated mitochondrial fission, we overexpressed YAP in TNFα-treated cells. The transfection efficiency was verified via immunofluorescence assay (Figure 5D and E) and Western blot (Figure 5F and G). Then, mitochondrial fission was evaluated again. As shown in Figure 5H and I, TNFα treatment promoted the formation of fragmented mitochondria whose length was shorter when compared to that of the control group. Interestingly, TNFα-mediated mitochondrial division could be inhibited by YAP overexpression (Figure 5H and I). Altogether, our results confirm that MAPK–ERK–YAP signaling pathways are required for TNFα-controlled mitochondrial fission.

Figure 5

TNFα handles mitochondrial fission via MAPK–ERK–YAP pathways.

Notes: (A–C) The expression values of ERK and YAP were determined via Western blot. Phosphorylated ERK and YAP expressions were both downregulated by TNFα. Subsequently, Ad-YAP was transfected into cells to overexpress YAP in TNFα-treated cells. PD98059 was used to inhibit ERK activity, which was used to mimic the inhibitory effects of TNFα on ERK pathways. (D and E) Immunofluorescence of p-ERK and YAP in cells treated with TNFα or transfected with Ad-YAP or Ad-ctrl. (F and G). The overexpression efficiency of Ad-YAP infection. Western blot was performed to analyze the protein expression of YAP in cells treated with Ad-YAP or Ad-ctrl. (H and I) Mitochondrial fission was determined via immunofluorescence using mitochondrial-specific Tom20 antibody. The average length of mitochondria was evaluated to quantify mitochondrial fission. *P<0.05.

Abbreviation: Ad, adenovirus.

MAPK–ERK–YAP signaling pathways are also involved in mitochondrial malfunction and glioblastoma cell death

We explored whether MAPK–ERK–YAP signaling pathways are involved in TNFα-mediated mitochondrial injury and cell death. First, ROS production was measured via immunofluorescence assay. Compared to the control group, TNFα treatment elevated the levels of cell ROS (Figure 6A and B), and this effect was reversed by YAP overexpression. In addition, cyt-c translocation from the mitochondria into the cytoplasm/nucleus was exacerbated by TNFα stress and was repressed by YAP overexpression (Figure 6C and D). In response to cyt-c leakage, caspase-9 activity was augmented in TNFα-treated cells and was reduced to near-normal levels with YAP overexpression (Figure 6E). Altogether, this information indicated that TNFα-mediated mitochondrial injury could be interrupted via activation of the MAPK–ERK–YAP axes.

Figure 6

MAPK–ERK–YAP pathways also participate in the regulation of mitochondrial homeostasis and cell death.

Notes: (A and B) ROS production was measured using immunofluorescence. Ad-YAP was transfected into cells to reactivate MAPK–ERK–YAP pathways. (C and D) Immunofluorescence assays for cyt-c. The cellular location of cyt-c was determined, and DAPI was used to label the nucleus. (E) Caspase-9 activity was examined to determine the role of MAPK–ERK–YAP pathways in caspase-9-mediated mitochondrial apoptosis. (F and G) TUNEL staining for apoptotic cells. The ratio of TUNEL-positive cells was recorded. (H) An LDH release assay was used to analyze cell death. The ratio of relative LDH release was recorded compared to the control group. *P<0.05. Abbreviation: Ad, adenovirus.

With respect to cell apoptosis, TUNEL assays were conducted to observe the apoptotic cells. Compared to the control group, TNFα treatment elevated the number of TUNEL-positive cells (Figure 6F and G), and this effect was abrogated by YAP overexpression. Similarly, the LDH cytotoxic test also indicated that TNFα-mediated LDH release could be suppressed by YAP overexpression (Figure 6H). Collectively, the above data demonstrate that TNFα-mediated mitochondrial damage and cell death are mainly regulated by the MAPK–ERK–YAP axes.

Discussion

The treatment of glioblastomas currently remains difficult due to inevitable recurrence and rapid progression.39 Current treatment options include radiation therapy in addition to surgery or surgery combined with chemotherapy.40 In the present study, we found that TNFα treatment significantly reduced the viability of glioblastoma cells in a dose-dependent manner. Functional investigations revealed that TNFα supplementation activated mitochondrial fission and that mitochondrial fission subsequently mediated mitochondrial injury and initiated caspase-9-involved mitochondrial apoptosis. Inhibition of mitochondrial fission could abrogate the proapoptotic effects of TNFα on glioblastoma cells. Furthermore, we showed that TNFα-induced mitochondrial fission was modified by the MAPK–ERK–YAP signaling pathways. TNFα treatment repressed the activity of MAPK–ERK–YAP signaling pathways, leading to an increase in the content of mitochondrial fission factors such as Drp1. Reactivation of MAPK–ERK–YAP signaling pathways could inhibit TNFα-mediated mitochondrial fission and provide a prosurvival advantage for glioblastomas cells. Collectively, this is the first study to demonstrate that TNFα regulates glioblastoma cell viability and mitochondrial homeostasis by modulating mitochondrial fission through MAPK–ERK–YAP-dependent signaling pathways (Figure 7). Our results lay the foundation to help us understand the molecular mechanisms of TNFα-mediated cancer cytotoxicity.

Figure 7

TNFα treatment elevates the apoptotic rate of glioblastoma in vitro by initiating fatal mitochondrial fission and interrupting MAPK–ERK–YAP signaling pathways.

TNFα, an inflammatory cytokine, is of significant importance in regulating cancer progression in many types of malignant tumors.41,42 This fact led to several animal experiments and clinical studies to explore the detailed role of TNFα in retarding the progression of glioblastomas. Early studies have demonstrated that gene transduction of a human TNFα-vector substantially increased the apoptotic index and reduced the growth rate in human glioblastoma cells.2 Subsequent studies determined that TNFα supplementation enhanced the susceptibility of human glioblastoma cells to natural killer cells.43 In addition, TNFα treatment also reduced the adhesion capacity, evoked cellular oxidant stress,44,45 and suppressed tumor angiogenesis46 in primary or recurrent glioblastomas. In the current study, our results demonstrated that TNFα stress was closely associated with mitochondrial damage in glioblastoma cells. In response to TNFα stimulus, mitochondrial ROS production was increased, which was accompanied by a drop in the levels of antioxidant factors. In addition, mitochondrial ATP production was also impaired, which may result from TNFα-mediated downregulation of the mitochondrial respiratory complex. More importantly, decreased mitochondrial potential, extended mPTP opening time, and more cyt-c liberation into the nucleus were noted in TNFα-treated cells. These alterations worked together to initiate caspase-9-related mitochondrial apoptotic pathways, accounting for glioblastoma cell death. Our findings are similar to previous studies that indicated that TNFα treatment promoted mitochondrial dysfunction in glioblastoma cells.47 This information identifies mitochondria as a primary target for TNFα-based therapy. Based on this, the discovery of other drugs principally acting on mitochondria may provide more clinical benefits for patients with glioblastoma.

The novel finding in our study is that we show that TNFα induces mitochondrial damage via mitochondrial fission. Notably, mitochondrial fission has been suggested as a chief cause of cell death by inducing mitochondrial damage in several diseases. In cardiac ischemia-reperfusion injuries, aberrant mitochondrial fission exacerbates cardiomyocyte death via promoting mPTP opening and cardio-lipin oxidation.5,6,48 Moreover, uncontrolled mitochondrial fission also participates in fatty liver disease by disrupting hepatocyte mitochondrial metabolism.49 In addition, in pancreatic cancer,13 breast cancer,11 ovarian cancer,12 and liver cancer,50 mitochondrial fission exerts negative effects on mitochondrial homeostasis and has been proposed to be a primary apoptotic trigger. In the present study, we show for the first time that mitochondrial fission induces mitochondrial damage, which precedes cell apoptosis in a caspase-9-dependent manner. This is the first study to define the role of mitochondrial fission in glioblastoma. Considering the detrimental effects of mitochondrial fission on cell viability, approaches to activate mitochondrial fission are of utmost importance when designing antitumor therapies. Notably, several studies have also found that TNFα treatment also activated mitochondrial fusion in human kidney-2 cells51 and cardiomyocytes.52 These results establish the various effects of TNFα on mitochondrial fission and mitochondrial fusion. This seems to be dependent on cell types. However, more researches are required to validate our concept.

At the molecular level, we found that TNFα activated mitochondrial fission via repression of MAPK–ERK–YAP signaling pathways. First, more robust data concerning the inhibitory effects of MAPK–ERK pathways on mitochondrial fission have been provided by several in vitro and in vivo studies.53 More importantly, the MAPK–ERK pathway, as the classical antiapoptotic pathway, has been demonstrated to send beneficial signals to cells under various states of stress.54 As a new downstream effector of MAPK–ERK pathways, YAP was originally identified as a proto-oncogene.55 Higher YAP expression is closely correlated with cancer progression and tumor metastasis.18 In addition, increased YAP effectively controls mitochondrial fission and sustains mitochondrial integrity,18 favoring cell metabolism and growth. Based on this finding, several researchers propose that MAPK–ERK–YAP pathways are the upstream inhibitors of mitochondrial fission. This conclusion is supported by our results. We found that reactivation of MAPK–ERK–YAP pathways repressed mitochondrial fission and abrogated TNFα-mediated cell death. Accordingly, our results combined with previous findings highlight the molecular mechanisms by which TNFα regulates mitochondrial fission. At the molecular levels, several researchers have investigated the mechanism by which YAP modulated mitochondrial fission. Increased YAP reduces the transcription and expression of Mff and Drp1, strongly attenuating mitochondrial fission. Moreover, YAP has an ability to modify the phosphorylation of Drp1. In addition, YAP overexpression also reverses mitochondrial fusion via upregulating the expression of mitochondrial fusion factors such as OPA1 and Mfn2. These results explain the inhibitory effect of YAP on mitochondrial fission.

The clinical implication that can be drawn from our study is multifold. Our data provide a piece of evidence for the role of mitochondrial fission in glioblastoma viability. This information indicates that mitochondrial fission would be considered as a potential target to prevent glioblastoma progression via promoting mitochondrial fission-mediated cell apoptosis. On the other hand, our findings identify MAPK–ERK–YAP pathways as novel regulators for handling mitochondrial function and glioblastoma viability. This may highlight a new entry point for treating glioblastoma by targeting the MAPK–ERK–YAP signaling axes.

Limitation

The primary limitation of our study is that only one cell line was used in the present study to explore the roles of TNFα and mitochondrial fission in cell viability. Animal studies and clinical researches are required to further verify our findings.56

Conclusion

Altogether, our results show that TNFα treatment elevates the apoptotic rate of glioblastoma in vitro by initiating fatal mitochondrial fission and interrupting MAPK–ERK–YAP signaling pathways. These findings define mitochondrial fission as a novel tumor suppressor that acts by inducing mitochondrial damage, with potential implications for new approaches to glioblastoma treatment.

Availability of data and materials

The datasets used and/or analyzed during the current study are available from the corresponding author on reasonable request.