Requirement of FADD, NEMO, and BAX/BAK for aberrant mitochondrial function in tumor necrosis factor alpha-induced necrosis

Abstract

Necroptosis represents a form of alternative programmed cell death that is dependent on the kinase RIP1. RIP1-dependent necroptotic death manifests as increased reactive oxygen species (ROS) production in mitochondria and is accompanied by loss of ATP biogenesis and eventual dissipation of mitochondrial membrane potential. Here, we show that tumor necrosis factor alpha (TNF-α)-induced necroptosis requires the adaptor proteins FADD and NEMO. FADD was found to mediate formation of the TNF-α-induced pronecrotic RIP1-RIP3 kinase complex, whereas the IκB Kinase (IKK) subunit NEMO appears to function downstream of RIP1-RIP3. Interestingly, loss of RelA potentiated TNF-α-dependent necroptosis, indicating that NEMO regulates necroptosis independently of NF-κB. Using both pharmacologic and genetic approaches, we demonstrate that the overexpression of antioxidants alleviates ROS elevation and necroptosis. Finally, elimination of BAX and BAK or overexpression of Bcl-x(L) protects cells from necroptosis at a later step. These findings provide evidence that mitochondria play an amplifying role in inflammation-induced necroptosis.

Fig. 1.

TNF-α–CHX–zVAD challenge results in FADD-, RIP1-, NEMO-, and BAX/BAK-dependent reduction of mitochondrial oxygen consumption. (A) Determination of mitochondrial oxygen consumption by a Clark-type electrode in wild-type MEFs challenged with or without TNF-α (25 ng/ml)–cycloheximide (0.25 μg/ml)–zVAD (50 μM) (TCZ) for 6 h. For the following experiments, MEFs were challenged with TCZ for 20 h. (B to G) Wild-type (B), FADD^−/− (C), FADD^−/− FADD (D), RIP1^−/− (E), NEMO^−/− (F), and BAX^−/− BAK^−/− (G) MEFs. These cells were then permeabilized with intracellular medium buffer containing digitonin (40 μg/ml). Mitochondrial complex I (malate/pyruvate [Mal/Pyr]; 5 mM), II (succinate [Succ]; 5 mM), and IV (TMPD at 0.4 mM plus ascorbate [ASC] at 2.5 mM) substrates were subsequently added to the closed chamber, and the rate of oxygen consumption was measured. Black traces, untreated samples; red traces, representative results for samples treated for 20 h with TCZ. MEFs were pretreated with CHX (0.25 μg/ml) plus zVAD-FMK (50 μM) for 1 h prior to TNF-α stimulation. (H) Quantitation of mitochondrial respiratory complex IV oxygen consumption rate following TCZ treatment. Oxygen consumption was measured using TMPD-ascorbate as the electron donor, and the results of the analysis are expressed as complex IV oxygen consumption (nanomoles O₂/2.0 × 10⁶ cells/min). Note that the rate of oxygen consumption by complexes I and II was reduced at 20 h but not 6 h of TCZ challenge in wild-type MEFs. Black traces, untreated samples; red traces, representative results for samples treated with TCZ. The bar graph represents means ± SEMs of at least three independent experiments.

Fig. 2.

Disruption of mitochondrial membrane potential gradient in TNF-α-induced necroptosis requires FADD, RIP1, NEMO, and BAX/BAK. MEFs were treated with dimethyl sulfoxide or TCZ for 20 h. Permeabilized cells were loaded with the ratiometric mitochondrial membrane potential indicator JC-1, and ΔΨ_m was examined for 18 min. At 1,000 s, cells were challenged with a mitochondrial uncoupler (carbonyl cyanide m-chlorophenylhydrazone [CCCP]; 2 μM) to dissipate the total ΔΨ_m. (A) Wild-type cells challenged with TCZ induced a large ΔΨ_m loss. (B) FADD^−/− MEFs were resistant to TCZ-induced ΔΨ_m loss. (C) In contrast, reintroduction of full-length FADD in FADD^−/− MEFs reestablished the ΔΨ_m loss in response to TCZ challenge. (D) Similarly, RIP1^−/− MEFs failed to undergo ΔΨ_m loss after TCZ treatment. (E) TCZ challenge partially affected the caspase 8^−/− MEFs ΔΨ_m gradient. NEMO^−/− MEF (F) and BAX^−/− BAK^−/− MEF (G) ΔΨ_m was unaffected by TCZ challenge. (H) Quantitation of ΔΨ_m maintenance in MEFs before and after TCZ treatment. Data are represented as means ± SEMs from three to six independent experiments.

Fig. 3.

Proximal candidates FADD, RIP1, and NEMO are required for depolarization of mitochondrial membrane during necroptosis. MEFs were challenged for 20 h with TCZ or dimethyl sulfoxide. To perform cytosol exchange experiments, TCZ-treated wild-type (A), FADD^−/− (B), FADD^−/− FADD (C), RIP1^−/− (D), caspase 8^−/− (E), NEMO^−/− (F), and BAX^−/− BAK^−/− (G) cells were permeabilized for 6 min with intracellular medium containing digitonin (40 μg/ml). The cytosol was prepared after centrifugation at 20,000 rpm for 5 min. Similarly, a wild-type mitochondrion-enriched pellet was prepared for cytosol swap experiments. Wild-type ΔΨ_m was unaffected by the cytosol from FADD^−/− (B), RIP1^−/− (D), and NEMO^−/− (F) TCZ-challenged MEFs. (E) Although ΔΨ_m was maintained at the earlier time point, caspase 8^−/− cytosol also elicited a considerable amount of wild-type ΔΨ_m loss. In sharp contrast, cytosol from the FADD-reconstituted FADD^−/− cells induced complete ΔΨ_m loss (C). Interestingly, TCZ-treated BAX^−/− BAK^−/− cytosol induced a substantial amount of wild-type ΔΨ_m loss. (H) Quantitation of wild-type ΔΨ_m preservation after cytosol exchange. The results in panel H are representative of three to six independent experiments.

Fig. 4.

FADD, RIP1, NEMO, and BAX/BAK determine the preservation of ΔΨ_m and cell survival in TNF-α-induced necroptosis. (A) FADD is required for RIP1-RIP3 complex formation following TCZ challenge. WT, FADD^−/−, and RIP1^−/− MEFs were treated as indicated, and equal amounts of cell lysates were used for coimmunoprecipitation (IP). The formation of FADD-RIP1-RIP3 complex is shown by RIP3 coimmunoprecipitation in wild-type, FADD^−/−, and RIP1^−/− MEFs. Pellets from immunoprecipitation using an anti-RIP3 antibody were analyzed by Western blotting (immunoblotting [IB]) using anti-RIP1 and anti-FADD antibodies. Nec-1 (30 μM) pretreatment inhibits the TCZ-induced RIP1-RIP3 complex formation. MEFs were cultured in glass-bottom petri dishes, and cells were challenged with TCZ for 24 h. After the treatment, live cells were simultaneously loaded with mitochondrial membrane potential indicator TMRE (50 nM) and plasma membrane integrity marker TOTO-3 to assess mitochondrial function and cell viability, respectively. (B) Wild-type MEFs are positive for TOTO-3, and complete ΔΨ_m loss was very significant. Conversely, FADD^−/−, RIP1^−/−, NEMO^−/−, and BAX^−/− BAK^−/− MEFs retained the ΔΨ_m and cell viability (C, E, G, H, and K). (D) Reconstitution of full-length FADD in FADD^−/− MEFs rescued the wild-type phenotype. (F) Caspase 8^−/− MEFs experience a significant ΔΨ_m loss but failed to undergo necroptotic cell death. (I) RIP1 inhibitor necrostatin-1 (30 μM) prevented the TCZ-induced ΔΨ_m loss and cell death. (J and K) Quantitation of ΔΨ_m maintenance and cell viability after TCZ treatment. The data in panels J and K are means ± SEMs of three or four independent experiments.

Fig. 5.

Silencing of RIP3 attenuates TCZ-induced ΔΨ_m loss. Human pulmonary endothelial cells transfected with a smart pool of human RIP3 siRNA were cultured for 48 h. (A) RIP3 protein levels after siRNA gene silencing at 48 h posttransfection in human pulmonary microvascular endothelial cells. (B) Cells were challenged with TCZ for 24 h and incubated with the ΔΨ_m indicator TMRE for 20 min. ΔΨ_m was measured using confocal microscopy. Data are presented as means ± SEMs of at least three independent experiments. Ser, scrambled.

Fig. 6.

TCZ-induced dissipation of ATP levels requires FADD, RIP1, NEMO, and BAX/BAK. Wild-type (A), FADD^−/− (B), FADD^−/− FADD (C), RIP1^−/− (D), caspase 8^−/− (E), NEMO^−/− (F), and BAX^−/− BAK^−/− (G) MEFs were treated with TCZ for 20 h. (H) Cell viability was assessed by measuring ATP levels using a CellTiter-Glo luminescent cell viability assay kit. Data are presented as means ± SEMs of at least three independent experiments.

Fig. 7.

NEMO but not RelA is required for the induction of TCZ-mediated necroptosis. (A) Expression of recombinant WT NEMO in NEMO-KO MEFs. NEMO^−/− MEFs were transfected with human NEMO plasmid. Cell lysates were subjected to Western blotting and probed with anti-NEMO antibody. (B) MEFs were challenged for 20 h with TCZ or dimethyl sulfoxide. After the treatment, NEMO^−/− MEFs and MEFs in which NEMO was reexpressed were permeabilized (40 μg/ml digitonin) and loaded with the ratiometric ΔΨ_m fluorophore JC-1 and ΔΨ_m was monitored. Cells were pulsed with the uncoupler carbonyl cyanide m-chlorophenylhydrazone (CCCP; 1 μM) at 1,000 s. (C) Quantitation of ΔΨ_m maintenance in NEMO^−/− MEFs and MEFs in which NEMO was reexpressed before and after TCZ treatment (n = 3; means ± SEMs). (D) Cell viability was determined at 24 h posttreatment. (E) Lack of NEMO does not alter the TCZ-induced FADD-RIP1-RIP3 complex formation. Wild-type and NEMO^−/− MEFs were pretreated with Nec-1 (30 μM). These cells were then treated with TCZ and RIP3, and coimmunoprecipitation (IP) was performed. Immunoprecipitated eluates were subjected to immunoblotting (IB) for detection of the FADD-RIP1-RIP3 complex. (F) Wild-type MEFs were pretreated with selective IKKβ inhibitor IMD-0354 alone (1 μM; Tocris Bioscience, MO) or in combination with TCZ for 20 h. Cells were treated with dimethyl sulfoxide or IMD-0354 for 2 h prior to the TCZ challenge. ATP levels were determined by a CellTiter-Glo luminescent cell viability assay kit. Data are presented as means ± SEMs of at least quadruplicate experiments. (G) RelA^−/− MEFs are susceptible to TCZ-induced cytotoxicity. RelA^+/+ and RelA^−/− MEFs pretreated with or without Nec-1 (30 μM) were challenged with TCZ for 24 h, and cell viability was determined. Data are presented as means ± SEMs of at least three independent experiments.

Fig. 8.

RIP1-RIP3 complex-mediated ΔΨ_m loss requires FADD and NEMO. FADD^−/− and NEMO^−/− MEFs were challenged for 20 h with TCZ or dimethyl sulfoxide. To perform cytosol exchange experiments, TCZ- or dimethyl sulfoxide-treated NEMO^−/− MEFs were permeabilized for 6 min with intracellular medium containing digitonin (40 μg/ml). The cytosol was prepared after centrifugation at 20,000 rpm for 5 min. Similarly, an untreated FADD^−/− MEF mitochondrion-enriched pellet was prepared for cytosol swap experiments. (A) TCZ-treated NEMO^−/− MEF cytosol did not trigger ΔΨ_m loss in FADD^−/− MEF mitochondria. The arrow indicates the time at which FCCP was added. Data are presented as means ± SEMs of at least three independent experiments. TCZ induces association of FADD to mitochondria. CCCP, carbonyl cyanide m-chlorophenylhydrazone. (B) Subcellular fractionation of wild-type MEFs. Control and TCZ-treated wild-type MEFs were fractionated into mitochondrial, microsomal, and cytosolic fractions. The mitochondrial and microsomal fractions were lysed using radioimmunoprecipitation assay buffer, and the soluble proteins were analyzed for their purity. Subcellular fractions were immunoblotted with antibodies against cytochrome c oxidase (Cox), calnexin, and β-actin. (C) Purified mitochondria from both untreated and TCZ-treated MEFs (4 and 6 h) were lysed, and the supernatant was then analyzed by Western blotting with FADD, RIP3, and cytochrome c oxidase antibodies. Data are presented as means ± SEMs of at least three independent experiments.

Fig. 9.

Cytosolic calcium is dispensable for TCZ-induced RIP1-RIP3 complex formation and mitochondrial dysfunction during necroptosis. (A) Wild-type MEFs were challenged with TCZ in the absence or presence of BAPTA pretreatment. Cell lysates were subjected to immunoprecipitation (IP) with anti-RIP3 antibody. Immunoprecipitates were analyzed by Western blotting (immunoblotting [IB]) with anti-RIP1 antibody. (B) Chelation of cytosolic calcium did not prevent TCZ-induced ΔΨ_m loss. Wild-type MEFs were treated with dimethyl sulfoxide or TCZ for 20 h. Permeabilized cells were loaded with the ratiometric mitochondrial membrane potential indicator JC-1, and ΔΨ_m was examined. At 1,000 s, carbonyl cyanide m-chlorophenylhydrazone (CCCP) was added to dissipate the total ΔΨ_m. (C) MEFs were subjected to TCZ exposure following BAPTA pretreatment, and ATP levels were determined. (D) Wild-type MEFs were pretreated with cyclosporine (2.5 μM) before TCZ challenge. Permeabilized cells were loaded with JC-1, and ΔΨ_m was measured. (E) Quantitation of ΔΨ_m maintenance in wild-type MEFs with either TCZ alone or the combination of cyclosporine (CsA) and TCZ treatment. (F) Cell viability was determined at 24 h posttreatment. Data are presented as means ± SEMs of at least three independent experiments.

Fig. 10.

Overexpression of Bcl-x_L attenuates TCZ-induced necroptosis. (A) Wild-type MEFs were transfected with Bcl-x_L plasmid. The overexpression of Bcl-x_L was assessed by Western blotting. (B) MEFs were cultured in glass-bottom petri dishes, and cells were challenged with TCZ for 20 h. After the treatment, live cells were simultaneously loaded with the ΔΨ_m indicator TMRE and the plasma membrane integrity marker TOTO-3 to assess the mitochondrial function and cell viability, respectively. (C) Quantitation of ΔΨ_m maintenance and cell viability after TCZ treatment. (D) Wild-type MEFs were treated with TCZ for 4 and 6 h. Cells were isolated and treated with 10 mM bis-maleimide cross-linker. The BAX oligomers were analyzed by anti-BAX immunoblotting. (E) Wild-type MEFs were treated with TCZ for 6 h, and cells were fixed and immunostained with anti-cytochrome c antibody (green), anti-Smac/Diablo antibody (green), anti-HtrA2/Omi (green), and anti-AIF antibody (green). Hoechst dye (blue) was used as a nuclear stain. (F) Quantitation of mitochondrial intermembrane space proteins (cytochrome c, Smac/Diablo, HtrA2/Omi, and AIF). Data are presented as means ± SEMs of at least three independent experiments.

Fig. 11.

TCZ-induced reactive oxygen species production is FADD and RIP1 dependent. MEFs were treated with dimethyl sulfoxide (DMSO) or TCZ for 12 h. Cells were then loaded with ROS indicator dichlorofluorescein diacetate (H₂DCF-DA; 10 μM), and fluorescence intensity was assessed by flow cytometry. (A) Wild-type MEFs challenged with TCZ exhibited a large rise in ROS levels, whereas FADD^−/− cells failed to increase ROS production. Stable reexpression of full-length FADD in FADD^−/− cells reestablished the ROS elevation after TCZ challenge. RIP1^−/− (B), caspase 8^−/− (C), and BAX^−/− BAK^−/− (E) cells produced nominal ROS. (D) NEMO^−/− MEFs challenged with TCZ displayed ROS elevation. (F) Quantitation of DCF fluorescence change after TCZ treatment. (G) Quantitation of DHE fluorescence after TCZ treatment. Data are presented as means ± SEMs of at least three independent experiments.

Fig. 12.

Overexpression of cytosolic and mitochondrial antioxidants alleviates the TCZ-induced ROS production. Wild-type MEFs were transduced with empty adenoviral vector (Ad5CMV), MnSOD, or GPX for 36 h. Transduced cells were treated with TCZ for 14 h. Wild-type cells were treated with TCZ for 12 h and the flavoprotein inhibitor DPI (20 μM) for 2 h. Cells were loaded with CMH₂DCF-DA, and ROS levels were assessed by flow cytometry. (A) Cells transduced with the empty vector alone challenged with TCZ demonstrated an increase ROS production. MnSOD (B) or glutathione peroxidase (C) overexpression attenuated the TCZ-induced ROS production. (D) Similarly, DPI supplementation prevented the ROS elevation. (E) Quantitation of ROS levels in MnSOD, glutathione peroxidase-overexpressed, or DPI-treated TCZ samples. Data are presented as means ± SEMs of at least triplicate experiments. (F) Wild-type MEFs were overexpressed with adenoviral empty vector, MnSOD, GPX, or the combination of MnSOD and glutathione peroxidase. MEFs were subjected to TCZ exposure, and ATP levels were determined by a CellTiter-Glo luminescent cell viability assay kit. Data are presented as means ± SEMs of at least three independent experiments.