c-FLIP Protects T Lymphocytes from Apoptosis in the Intrinsic Pathway

Cellular caspase 8 (FLICE)-like inhibitory protein (c-FLIP; also named CLARP, FLME, I-FLICE, MRIT, usurpin, Casper, or CASH) is an important regulator in death receptor–mediated cell death pathway, also termed the extrinsic apoptosis pathway (1). Three isoforms of c-FLIP have been identified in humans, 55-kDa c-FLIP_L, 24-kDa c-FLIP_R, and 26-kDa c-FLIP_S, whereas two isoforms, c-FLIP_L and c-FLIP_R, are found in mice (2, 3). The c-FLIP_L isoform has two death effector domains at the N terminus and an inactive caspase-like domain at the C terminus. In contrast, c-FLIP_S and c-FLIP_R contain only two death effector domains without the inactive caspase-like domain. Studies have shown that all three isoforms inhibit death receptor–induced apoptosis (1–3).

Death receptor ligation initiates the extrinsic apoptosis pathway by recruiting procaspase 8 through adaptor protein Fas-associated via death domain or TNFR1-associated via death domain. The dimerization of procaspase 8 upon the recruitment leads to cleavage in their C terminus and release of the active form of caspase 8. c-FLIP protein suppresses apoptosis by forming heterodimers with procaspase 8 to inhibit its activation (1). c-FLIP–deficient T cells have defective survival upon TCR stimulation (4). Interestingly, blocking extrinsic apoptosis pathway by ablating Fas receptor and/or TNF-α only partially rescues the impaired survival of c-FLIP–deficient T cells after TCR activation, suggesting that c-FLIP may promote cell survival in a death receptor–independent pathway (4). Therefore, it is important to examine whether c-FLIP is involved in the intrinsic apoptosis pathway in T lymphocytes, which is triggered by mitochondrial stress and mediated through mitochondrion release of apoptotic factors (5).

The long isoform of c-FLIP, c-FLIP_L, also regulates autophagy and necroptosis, two processes involved in cell survival in T lymphocytes (6). Autophagy is essential for T cell homeostasis (7). Deficiency in any autophagy pathway members, Atg5, Atg3, Atg7, or Beclin-1 causes impaired T cell survival (8–12). However, suppressing autophagy under certain conditions promotes T cell survival (13). c-FLIP overexpression represses autophagy and improves cell survival by binding to Atg3 and preventing Atg3 interaction with LC3, a process essential for autophagosome formation (14). Necroptosis is a type of programmed necrotic cell death, which can be suppressed by caspase 8 activity (15). Loss of the long isoform of c-FLIP leads to necroptosis in T cells upon TCR stimulation (16). Meanwhile, necroptosis is undetected in resting T lymphocytes without apoptosis induction. Whether c-FLIP protects mature T cells from the intrinsic death pathway through regulating autophagy and necroptosis needs to be determined.

Previously we reported that c-FLIP suppresses death receptor–induced apoptosis and TCR activation–induced cell death by inhibiting caspase 8 activation (4). In this study, we found that cell survival was impaired in c-FLIP–deficient naive T lymphocytes when apoptosis is induced through the intrinsic death pathway. Importantly, the defective survival of c-FLIP–deficient T cells upon intrinsic apoptosis induction can be rescued by deletion of proapoptotic Bcl-2 family member Bim. Although autophagy is upregulated in c-FLIP–deficient T cells, inhibition of autophagy does not improve cell survival in c-FLIP^−/− T cells upon intrinsic apoptosis induction, ruling out a possibility that excess autophagy caused cell death of resting c-FLIP–deficient T cells upon induction of the intrinsic apoptosis pathway.

c-FLIP^f/f mice were generated previously (17). c-FLIP conditional knockout mice were generated by crossing c-FLIP^f/f to ER-Cre⁺ mice (18). Mice deficient for Bim, Bax, and Bak were obtained from the Jackson Laboratory (Bar Harbor, ME). Animals were genotyped as described previously (17, 19) and used at 6–8 wk of age. Animal usage was conducted according to protocols approved by the Duke University Institutional Animal Care and Use Committee.

Single-cell suspensions prepared from spleens and peripheral lymph nodes are resuspended in ACK lysis buffer (0.15 M NH₄CL, 10 mM KHCO₃, 0.1 mM EDTA pH 7.4) for up to 3 min for RBC lysis. For induced deletion, total splenocytes were cultured in complete RPMI 1640 medium containing 10% FBS, 200 nM 4-hydroxytamoxifen (4-OHT; Sigma-Aldrich) and 1 ng/ml IL-7 (PeproTech) at 37°C in the presence of 5% CO₂ for 3 d. Live cells were purified after deletion using Ficoll. T lymphocytes were enriched using an EasySep mouse T cell–negative enrichment kit from Stemcell Technologies according to the manufacturer’s instructions. T lymphocytes were cultured in complete RPMI 1640 medium containing 10% FBS at 37°C in the presence of 5% CO₂ for indicated time. A total of 1 ng/ml IL-7 was added in the medium and re-added every 3 d. A total of 10 μM zVAD-fmk (Sigma-Aldrich), 10 μM zIETD-fmk (BD Pharmingen), 10 μM zLEHD-fmk (BD Pharmingen), 100 nM necrostatin-1 (Nec-1; Enzo Life Sciences), and 10 mM acetylcysteine (NAC) were added in lymphocyte cultures as indicated.

T lymphocytes were incubated with an FcR-blocking Ab (2.4G2), stained with FITC-, PE-, PE/Cy5-, allophycocyanin-, allophycocyanin-Cy7–, or Pacific blue–labeled mAbs on ice for 20 min, and washed with FACS buffer (2% FBS, 0.02% NaN₃ in PBS). Then cells were resuspended in Annexin V–binding buffer (10 mM HEPES, pH 7.4, 140 mM NaCl, 2.5 mM CaCl₂) and incubated with Annexin V–PE (BD Biosciences) and 7-aminoactinomycin D (7-AAD; BD Biosciences) at room temperature for 15 min. The cells were then diluted in Annexin V–binding buffer and analyzed by flow cytometry within 1 h. A total of 0.5–20 × 10⁵ events were collected on a FACSCanto II flow cytometer (BD Biosciences) and analyzed using FlowJo software (Tree Star). All fluorescence-labeled Abs, including anti-CD3, anti-CD4, anti-CD8, anti-TCRβ, and anti-CD19 were obtained from Biolegend.

Cytochrome c release was tested based on previously published protocol (20). In brief, T lymphocytes were enriched using an EasySep mouse T cell enrichment kit from StemCell Technologies according to the manufacturer’s instructions. Cell purity was determined by flow cytometry to be >95%. Cytosol was released by 200 mg/ml digitonin in 80 mM KCl buffer. The mitochondria/nuclear fraction was lysed in cell lysis buffer (50 mM Tris-HCl, pH 7.4, 150 mM NaCl, 2 mM EGTA, 2 mM EDTA, 0.2% Triton X-100, 0.3% NP-40, 1× Apotonin), then diluted to sample buffer (50 mM Tris-Cl [pH 6.8], 50 mM 2-ME, 2% SDS, 0.2% bromophenol blue, and 10% glycerol). Cytochrome c release was tested by Western blot.

T cells with a >95% purity were purified using an EasySep mouse T cell enrichment kit (Stemcell Technologies). T cell lysates were prepared in sample buffer (50 mM Tris-Cl [pH 6.8], 50 mM 2-ME, 2% SDS, 0.2% bromophenol blue, and 10% glycerol). Abs used for Western blots were rabbit anti-LC3 (polyclonal P014; MBL International), hamster anti–Bcl-2 (polyclonal; BD Pharmingen), rabbit anti–Bcl-x_L (polyclonal), rabbit anti–Mcl-1 (polyclonal; Rockland Immunochemicals), rabbit anti-Bim (polyclonal; Cell Signaling), rabbit anti-Bax (polyclonal; Cell Signaling), rabbit anti-Bak (polyclonal; Cell Signaling), rabbit anti-Bid (polyclonal; Abcam), rabbit anti–poly (ADP-ribose) polymerase 1 (anti–PARP-1; polyclonal; Cell Signaling), rabbit anti–COX IV (polyclonal; Cell Signaling), mouse anti–cytochrome c (7H8.2C12; Biolegend), mouse anti–α-Tubulin (B-5-1-2; Sigma-Aldrich), and goat anti–β-actin (polyclonal, Santa Cruz Biotechnology). For HRP-labeled Western blot, the secondary Abs were anti-rabbit IgG-HRP, anti-mouse IgG-HRP, anti-hamster IgG-HRP, and anti-goat IgG-HRP (Jackson Immunoresearch). The development of the Western blot was achieved with SuperSignal West Pico Chemiluminescent substrate (Thermo Scientific). For fluorescent Western blot, the secondary Abs were anti-rabbit IgG–Alexa Fluor 680, anti-mouse IgG–Alexa Fluor 680, and anti-goat IgG–Alexa Fluor 790 (Molecular Probes, Invitrogen).

Enriched primary T lymphocytes were first intracellularly stained with anti-LC3 (P015; MBL International), then stained with Cy3-, FITC- and Pacific blue–labeled, anti-rabbit IgG, anti-CD4, and anti-CD8. All images were captured with a custom-built Zeiss Observer D1 using a Zeiss 3100 objective lens and a 1.4 numerical aperture. Images were captured using a Photometrics Cool SNAP HQ2 and analyzed using MetaMorph software. Images were deconvoluted and thresholded. Deconvolution was done blind at 40 iterations. LC3⁺ staining is defined as >180% of background signal. LC3 puncta is defined as any enclosed LC3⁺ staining area no <10 pixels.

Single-cell suspensions were incubated with 2.5 mM dihydroethidium (DHE; Sigma-Aldrich) in RPMI 1640 medium at 37°C for 30 min or with 2.5 mM CM-H₂DCFDA (Invitrogen) in Dulbecco's PBS at 37°C for 15 min. The cells were then washed with RPMI 1640 medium; stained with anti–CD4-allophycocyanin, anti–CD8-allophycocyanin/Cy7, and 7-AAD; and then washed with FACS buffer and analyzed by FACS immediately.

Unpaired two-tailed Student t tests were used to compare the means of different samples.

Conditional deletion of c-FLIP in the thymocytes leads to impaired survival and severe periphery T cell reduction (17). The low cell number of c-FLIP–deficient T cells, along with the accumulative effect of the deletion, makes this model unsuitable for studying the role of c-FLIP in mature T lymphocytes. Therefore, we crossed c-Flip^f/f mice with ER-Cre⁺ mice to generate inducible c-FLIP knockout mice. Because in vivo deletion of c-FLIP leads to lethality of the animals caused by liver failure (21, 22), c-FLIP was deleted in vitro in T lymphocytes by 4-OHT (4). In brief, T cells were cultured in the presence of 0.2 μM 4-OHT and 1 ng/ml IL-7 for 72 h. This culture generally maintained the viability of unstimulated T cells, and no difference in the survival rates was observed between c-FLIP^f/f and c-FLIP^f/f ER-Cre⁺ T cells during the culture (Supplemental Fig. 1). After deletion, live cells were purified and the cell viability was confirmed by flow cytometry–based apoptosis assay (Fig. 1A, 1B). Although short-term IL-7 stimulation downregulated surface IL-7Rα, T cells restored their IL-7Rα expression on the cell membrane at the end of 3-d culture (Supplemental Fig. 2A). After in vitro–induced deletion, IL-7 stimulation effectively induced phosphorylation of Stat5 in purified wild type (WT) and c-FLIP–deficient T cells (Supplemental Fig. 2B). These data demonstrate that in vitro–induced deletion system did not alter T cell viability or IL-7R–mediated cell signaling.

After the 3-d induced deletion, live cells were enriched by Ficoll and cultured in the presence of 1 ng/ml IL-7. More than 95% of T cells were Annexin V⁻ 7-AAD⁻ after the enrichment (Fig. 1B). Apoptosis was induced in the cultured cells by the addition of staurosporine (STS) or etoposide (ETP). STS inhibits Protein kinase C and induces caspase 3–dependent apoptosis (23). ETP causes irreversible double-strand or single-strand breaks in DNA (24). Cell death was analyzed by Annexin V/7-AAD staining in CD4⁺ and CD8⁺ T cells 16 h after the addition of STS or ETP. As previously reported (4), c-FLIP–deficient T lymphocytes showed increased spontaneous death (Fig. 1A). The survival of c-FLIP–deficient T cells was dramatically impaired compared with that of WT T cells when treated with STS (Fig. 1B). In contrast, genomic toxin ETP did not trigger more cell death in c-FLIP–deficient T cells than in WT T cells (Fig. 1B).

c-FLIP_L has been shown to regulate necroptosis in activated T lymphocytes (16). Therefore, we first determined whether STS treatment induces necroptosis in c-FLIP–deficient T cells. The induction of apoptosis involves the activation of a series of caspases, which can be inhibited by pan-caspase inhibitor z-VAD-fmk. Necroptosis induction requires the activity of RIP-1 kinase, which can be repressed by Nec-1 (25). The impaired survival of c-FLIP^−/− T cells was partially rescued by z-VAD-fmk, but not by Nec-1 (Fig. 2A). Addition of Nec-1 to z-VAD-fmk–treated T cells did not further increase cell survival. Another biochemical marker of apoptosis is the caspase-dependent cleavage of PARP-1 (26). Cleaved form of PARP-1 was observed in both WT and c-FLP^−/− T cells after STS treatment (Fig. 2B). Furthermore, c-FLIP^−/− T cells showed spontaneous PARP-1 cleavage and higher ratio of cleaved PARP-1 to full-length PARP-1 3 h after STS addition, indicating more apoptosis in those cells (Fig. 2B). These data suggest that STS-treated T lymphocytes die through apoptosis, not through necroptosis.

Caspase 1 and caspase 3 activation are involved in the apoptosis induction by STS (27, 28). We used inhibitors of different caspases to test which apoptotic pathway is activated. Extrinsic pathway of apoptosis is triggered by death receptor ligation, followed by activation of caspases 8. Intrinsic pathway is characterized by cytochrome c release from mitochondria to cytosol and the activation of caspases 9 by apoptosome (29). The impaired survival of c-FLIP–deficient T lymphocytes was partially rescued by both caspases 9 inhibitor z-LEHD-fmk and pan-caspase inhibitor z-VAD-fmk (Fig. 3A). Caspases 9 inhibitor z-LEHD-fmk showed better rescue, especially in CD4⁺ T cells. Caspase 8 inhibitor z-IETD-fmk, however, failed to rescue the enhanced apoptosis in c-FLIP–deficient T lymphocytes (Fig. 3A). We did not detect active form of caspase 8 in either group of T cells (data not shown). The requirement of caspases 9 in STS-induced apoptosis suggested the involvement of the intrinsic apoptotic pathway in c-FLIP–deficient T cells. We measured cytochrome c release from mitochondria to cytosol in both WT and c-FLIP–deficient T cells, isolating cytosolic fraction by digitonin (20). The kinetic study showed that cytochrome c release was faster in c-FLIP–deficient T cells than in WT controls (Fig. 3B). These data indicate that c-FLIP protects T cells from the intrinsic pathway-induced apoptosis.

To further establish the role of c-FLIP in protecting T cells from the intrinsic apoptotic pathway, we determined whether deletion of Bcl-2 family proapoptotic proteins rescues their impaired survival upon STS treatment. c-Flip^f/f ER-Cre⁺ mice were crossed with Bak^−/−, Bax^−/−, and Bim^−/− mice to generate double knockouts. During apoptosis induction in the intrinsic pathway, Bak and Bax form oligomeric complex in the outer membrane of the mitochondria to mediate the release of cytochrome c (30, 31). Bax and Bak compensate for each other in the process of mitochondrial outer membrane permeabilization. Bak^−/−Bax^−/− cells show resistance to multiple death stimuli that mediates apoptosis through the intrinsic pathway, whereas Bax- and Bak- deficient cells are susceptible to them (32). Deletion of Bak or Bax did not rescue the apoptotic phenotype of c-FLIP^−/− T cells (Fig. 4A, 4B). Proapoptotic protein Bim responds to several stimuli and regulates the homeostasis of hematopoietic cells (33). Loss of Bim increased the survival of c-FLIP^−/− CD4⁺ T cells and fully rescued the impaired survival in c-FLIP^−/− CD8⁺ T cells after STS treatment (Fig. 4C). These results suggest that c-FLIP protects STS-induced apoptosis in T cells in a Bim-dependent pathway.

We next examined the expression level of Bcl-2 family member in c-FLIP–deficient T cells. The expression levels of proapoptotic protein Bcl-2, Mcl-1, and Bcl-x_L were comparable between WT and c-FLIP^−/− T cells (Fig. 4D). After STS treatment, Bcl-2 and Mcl-1 were downregulated in WT cells, whereas the expression of Bcl-x_L was increased (Fig. 4D). c-FLIP–deficient T cells failed to upregulate Bcl-x_L but retained the expression of Mcl-1. Previous data showed that c-FLIP interacts with Bcl-x_L (34). How this interaction regulates the function of Bcl-x_L remains to be studied. Interestingly, the surviving WT and c-FLIP–deficient T cell after apoptosis induction showed decreased levels of Bax and Bim, which may be the consequence that cells expressing a low level of proapoptotic proteins preferentially survived. These data suggest the cross talk between c-FLIP and Bcl-2 family members in the regulation of apoptosis in T lymphocytes.

Because c-FLIP_L modulates autophagy in T cells upon TCR stimulation (16), we tested whether c-FLIP–modulated autophagy in T lymphocytes was involved in the impaired survival in c-FLIP–deficient T cells upon intrinsic apoptosis induction. The conversion of LC3-I to LC3-II isoform and the consequent insertion of LC3-II in the isolation membrane are essential for autophagosome formation. Monitoring LC3 puncta formation inside the T cells is used in autophagy detection (35). We first determined the level of LC3-II in c-FLIP–deficient and WT T cells by Western blot. LC3-II level was very low in both WT and c-FLIP–deficient T cells in the absence of apoptosis stimuli. After STS treatment, LC3-II was increased, especially in c-FLIP–deficient T cells (Fig. 5A). We then intracellularly stained c-FLIP–deficient T cells and WT cells with anti-LC3 to examine the LC3 puncta for autophagosome quantitation. The number of LC3⁺ puncta in c-FLIP^−/− T cells was not significantly higher than that in WT T cells (Fig. 5B). However, c-FLIP–deficient T cells showed larger LC3 puncta and stronger fluorescent intensity, indicating larger autophagosome formed or higher LC3 protein level in the absence of c-FLIP (Fig. 5B). We further investigated whether blocking autophagy altered the cell survival in c-FLIP^−/− T lymphocytes upon STS treatment. Loss of autophagy-related gene Atg3 in T lymphocyte leads to inhibition of autophagy (11). Short-term deletion of Atg3 does not cause increased apoptosis in resting T cells (16). Therefore, c-Flip^f/f ER-Cre⁺ mice were crossed to Atg3^f/f mice to generate Atg3^f/fc-Flip^f/f ER-Cre⁺ mice. Atg3^f/fc-Flip^f/f littermates were used as controls. Inducible deletion of c-FLIP and Atg3 was conducted in vitro by 4-OHT treatment as previously described (16). Deletion of Atg3 in c-FLIP–deficient T cells that were treated with STS had no effect on their cell survival after STS treatment (Fig. 5C). These data suggest that autophagy is not the cause of the enhanced apoptosis of c-FLIP–deficient T lymphocytes after STS treatment.

STS has been shown to rapidly increase intracellular reactive oxygen species (ROS) in HeLa cells, and the high level of ROS contributes to caspase 3 activation and apoptosis induction (23). Notably, the long isoform of c-FLIP has been shown to promote cell survival by controlling ROS in activated T lymphocytes (16). Therefore, it was important to test whether ROS caused the increased apoptosis in STS-treated c-FLIP–deficient T lymphocytes. We examined the intracellular ROS by analyzing DHE and DCFDA staining using flow cytometry and found that loss of c-FLIP did not alter ROS production in T cells, in the presence or absence of STS (Fig. 6A, 6B). Addition of antioxidant NAC did not significantly improve the cell survival after the STS treatment in both WT and c-FLIP–deficient T cells (Fig. 6B and Supplemental Fig. 3). These results suggest that c-FLIP protects T cells against intrinsic apoptosis induction in an ROS-independent pathway.

Although the role of c-FLIP in the extrinsic apoptosis pathway in T cells has been well established, it is not clear whether c-FLIP also plays a critical role in the intrinsic apoptosis pathway in T cells. In this report, we show that c-FLIP–deficient T cells have impaired cell survival upon intrinsic apoptosis induction by STS. The enhanced apoptosis in c-FLIP–deficient T cells upon STS treatment is likely caused by a more rapid cytochrome c release and caspase 9 activation. The autonomous activation of procaspase 8 was not the cause of the apoptotic phenotype in resting c-FLIP–deficient T lymphocytes upon STS treatment. Furthermore, deletion of Bim significantly improved the survival of c-FLIP^−/− T cells in the presence of STS. Although autophagy is enhanced in c-FLIP–deficient T cells, inhibition of autophagy fails to rescue their impaired survival upon intrinsic apoptosis induction.

Interestingly, c-FLIP–deficient T cells were susceptible to STS-induced, but not ETP-induced, apoptosis. Although both drugs are widely used as intrinsic apoptosis inducers, they provoke apoptosis in distinct manners. ETP induces apoptosis through generating double-strand or single-strand break in the genomic DNA (24), whereas STS inhibits Protein kinase C and induces caspase 3–dependent apoptosis (23). The different susceptibility of c-FLIP–deficient T cells to these intrinsic apoptosis stimuli is likely because different apoptotic stimuli are sensed by different proapoptotic molecules in the cell. For example, Noxa targets Mcl-1 for proteasomal degradation in response to DNA damage (36). Puma is shown to be the predominant mediator in γ-radiation–induced apoptosis in T lymphocytes (37). IL-7 deprivation–induced apoptosis is primarily mediated by Bim (38). Our results show that c-FLIP protected T cells from STS-induced apoptosis in a Bim-dependent pathway. Therefore, Bim is likely to be the responding sentinel upon STS treatment. Our results provide important insights in understanding the cellular process of STS-induced apoptosis in T cells.

The enhanced intrinsic pathway-mediated apoptosis in resting c-FLIP–deficient T cells after STS treatment is not due to the secondary effect of spontaneous caspase 8 activation. In the Fas-induced extrinsic apoptosis pathway, active caspase 8 cleaves Bid, a BH3-only proapoptotic protein, to generate the truncate form of Bid (tBid). tBid efficiently inserts into mitochondrial outer membrane and augments cytochrome c release (39). It has been shown that c-FLIP–deficient T cells have higher caspase 8 activity after TCR activation (4). However, the intracellular level of active caspase 8 in resting T cells is too low to be detected (data not shown). Furthermore, caspase 8 inhibitor z-IETD-fmk did not rescue the impaired survival of c-FLIP–deficient T cells upon intrinsic apoptosis induction. Therefore, although caspase 8 augments cytochrome c release upon Fas ligation via tBid, it is unlikely to be the case in resting T cells in the absence of death receptor ligands. These results are consistent with the previous findings that the full activation of initiator caspases requires the assembly of a multicomponent complex (40). Caspase 9–activating apoptosome can be assembled in resting T cells after Bax/Bak-mediated mitochondrial outer membrane permiabilization and cytochrome c release. However, caspase 8–activating death-inducing signaling complex cannot be formed because of the lack of extrinsic apoptotic signals.

Our results demonstrate a critical role for c-FLIP in the intrinsic apoptotic pathway in resting T cells. Caspase 9 inhibitor z-LEHD-fmk and pan-caspase inhibitor z-VAD-fmk exhibit remarkable protection from apoptosis in c-FLIP–deficient T cells upon STS treatment. c-FLIP protects T cells from intrinsic apoptosis by antagonizing proapoptotic BH3-only protein Bim. Interestingly, Bim deficiency did not improve the survival of resting c-FLIP^−/− T cells under spontaneous condition. How c-FLIP regulates Bim function remains to be determined. In HIV gp120 protein–treated human Jurkat T cells, c-FLIP_L suppresses apoptosis by inhibiting Bax activation (41). However, our results show that c-FLIP promotes the survival of resting T cells upon intrinsic apoptosis induction in Bax-independent manner in mice.

STS treatment induces superoxide production and subsequent cell death in hippocampal neurons (27). ROS production may also contribute to the enhanced apoptosis in STS-treated, c-FLIP–deficient T lymphocytes. Antioxidant NAC slightly enhanced the survival of all STS-treated T cells. Thus, the minimum cytoprotective effect of NAC was not specific to c-FLIP–deficient T cells. Furthermore, the loss of c-FLIP (both isoforms) did not lead to an increase of ROS production in resting T cells. In contrast, the loss of c-FLIP_L leads to increased ROS production and ROS-dependent cell death in activated T cells (16). These results suggest that c-FLIP proteins function differently in resting and activated T lymphocytes.

Necroptosis was not detected in resting c-FLIP^−/− T cells. This result was consistent with previous publications showing necroptosis induction only in activated T lymphocytes (13, 15, 42, 43). Why activated T cells are susceptible to necroptosis has yet to be investigated. RIP-1 kinase plays an important role in T lymphocytes for proper TCR signaling. RIP-1–deficient T cells show impaired TCR-induced phosphorylation of p65 NF-κB (44). Activated T cells may upregulate RIP-1 for the benefit of cell signaling, whereas loss of adequate inhibition in RIP-1’s kinase activity ends up killing the cells.

Together, we show that c-FLIP regulates apoptosis signaling in the Bim-dependent intrinsic pathway. The mechanism of the cross talk of c-FLIP and Bim remains to be determined in future investigation. Our results suggest that c-FLIP protects T lymphocyte survival through multiple pathways.

We thank Dr. Nu Zhang for generating the c-FLIP^f/f mice and Dr. Qi-Jing Li for the assistance in fluorescent microscopy and advice.

The authors have no financial conflicts of interest.