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© The Rockefeller University Press,
0021-9525/2000//1247 $5.00
The Journal of Cell Biology, Volume 151, Number 6,
, 2000 1247-1256
Original Article |
Necrotic Death Pathway in FAS Receptor Signaling
nagata{at}genetic.med.osaka-u.ac.jp
A caspase 8–deficient subline (JB6) of human Jurkat cells can be killed by the oligomerization of Fas-associated protein with death domain (FADD). This cell death process is not accompanied by caspase activation, but by necrotic morphological changes. Here, we show that the death effector domain of FADD is responsible for the FADD-mediated necrotic pathway. This process was accompanied by a loss of mitochondrial transmembrane potential (
m), but not by the release of cytochrome c from mitochondria. Pyrrolidine dithiocarbamate, a metal chelator and antioxidant, efficiently inhibited the FADD-induced reduction of m and necrotic cell death. When human Jurkat, or its transformants, expressing mouse Fas were treated with Fas ligand or anti–mouse Fas antibodies, the cells died, showing characteristics of apoptosis. A broad caspase inhibitor (z-VAD–fmk) blocked the apoptotic morphological changes and the release of cytochrome c. However, the cells still died, and this cell death process was accompanied by a strong reduction in m, as well as necrotic morphological changes. The presence of z-VAD–fmk and pyrrolidine dithiocarbamate together blocked cell death, suggesting that both apoptotic and necrotic pathways can be activated through the Fas death receptor.
Key Words: apoptosis • caspase • Fas • mitochondrial membrane potential • necrosis
© 2000 The Rockefeller University Press
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Introduction |
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Apoptosis can be triggered by a variety of stimuli, including genotoxic agents, factor deprivation, and death factors (Nagata 1997; Ashkenazi and Dixit 1998; Green and Reed 1998; Raff 1998). It is involved in the removal of surplus cells generated during mammalian development, and in the removal of virally infected or cancer cells as part of the immune response (Raff et al. 1993; Nagata 1997). Apoptotic signal transduction has been extensively studied biochemically and genetically, and it is now known that a family of cysteine proteases with aspartate specificity are the major effectors of the process (Thornberry and Lazebnik 1998). Caspases exist as inactive precursors in proliferating cells and are activated in a cascade. Upon exposure to apoptotic stimuli, initiator caspases are activated by the formation of a complex with other molecules. Once the initiator caspases are activated, they cause the processing of downstream caspases, which cleave a set of cellular proteins, leading to morphological changes and the degradation of chromosomal DNA. Necrosis is known to occur as a result of complement attack, severe hypoxia, hyperthermia, lytic viral infection, or exposure to a variety of toxins and respiratory poisons. Tumor necrosis factor (TNF) can activate the necrotic death program in some cell lines, such as mouse L929 cells (Vercammen et al. 1998a). An involvement of necrosis in the removal of interdigital cells in the mouse embryo has also been suggested (Chautan et al. 1999). In contrast to apoptotic cell death, the molecular mechanism of necrosis is not well understood, though several mechanisms, including the release of lysosomal enzymes, the generation of toxic oxygen radicals, and the activation of calcium-dependent phospholipases, have been proposed (Fiers et al. 1999).
Fas ligand (FasL) is a member of the TNF family (Nagata and Golstein 1995; Nagata 1997) and was originally identified as a cytokine that triggers apoptosis by binding to Fas (Suda et al. 1993). Studies on FasL-induced apoptosis have revealed the following mechanism. The binding of FasL or agonistic anti-Fas antibody to Fas rapidly recruits procaspase 8 through an adaptor called FADD (Fas-associated protein with death domain) (Boldin et al. 1995; Chinnaiyan et al. 1995). FADD carries a domain called a death domain (DD) at its COOH terminus, which is involved in the binding to Fas. The NH2-terminal part of FADD is called the death effector domain (DED) and is responsible for recruiting procaspase 8 (Boldin et al. 1996; Muzio et al. 1996). Thus, procaspase 8 oligomerized at the plasma membrane is autoactivated to a mature enzyme (Muzio et al. 1998; Yang et al. 1998). Two pathways have been shown for the signal transduction downstream of caspase 8, which are used in different cell types (types I and II) (Scaffidi et al. 1998). In type I cells, caspase 8 directly activates procaspase 3. However, in type II cells, caspase 8 cleaves Bid, a proapoptotic member of the Bcl-2 family (Li et al. 1998; Luo et al. 1998; Gross et al. 1999). The cleaved Bid translocates to the mitochondria and stimulates the release of cytochrome c. Cytochrome c, together with Apaf-1, activates procaspase 9 (Li et al. 1997), which leads to the downstream processing of procaspase 3.
We and others have recently established sublines from human Jurkat cells that do not express caspase 8 (Juo et al. 1998; Kawahara et al. 1998). Engagement of Fas by agonistic anti–human Fas antibodies in these cell lines does not activate caspases, and, thus, cannot induce apoptotic cell death. On the other hand, when FADD was artificially oligomerized, these cells died. This cell death process was not accompanied by either activation of the caspase cascade or apoptotic morphological cell changes. Rather, the cells showed a necrotic morphology, suggesting that FADD can mediate not only caspase-dependent apoptotic signals, but also caspase-independent necrotic death signals.
To examine the necrotic death-signaling pathway, we prepared various mutants of FADD and showed that FADD's DED domain is responsible for transducing the signal. This death process was accompanied by a loss of mitochondrial membrane potential, but not by the release of cytochrome c from mitochondria, and was inhibited by pyrrolidine dithiocarbamate (PDTC). We then showed that Fas engaged by FasL or anti-Fas antibodies could activate both caspase-dependent and -independent cell death signals. The caspase-dependent signal was rapid and was accompanied by the release of cytochrome c from mitochondria, as well as apoptotic morphological changes. In contrast, the caspase-independent cell death occurred slowly and was accompanied by the loss of mitochondrial membrane potential and necrotic morphological changes.
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Materials and Methods |
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Establishment of Stable Transformants
The plasmid pEF-FK-FADD carries the coding sequence (FKBP-FADD) for a myristilation-targeting peptide, two tandem repeats of FKBP12, and human FADD in the pEF-BOS mammalian expression vector (Kawahara et al. 1998). The DNA fragments coding for the DED of FADD (FADD-DED, amino acids 1–117), its truncated form (FADD-DED, amino acids 79–117), and the DD of FADD (FADD-DD, amino acids 105–208) were prepared from human FADD cDNA (Boldin et al. 1995; Chinnaiyan et al. 1995). They were fused to a DNA fragment carrying a myristilation-targeting peptide and two tandem repeats of FKBP12, and inserted into pEF-BOS to generate pEF-FK-DED, pEF-F-FK-DED, pEF-F-FK-DED, and pEF-FK-DD (Fig. 1 A). In pEF-F-FK-DED and pEF-F-FK-DED, a DNA fragment coding for the FLAG epitope (DYKDDDDK) was inserted between the myristilation-targeting peptide and FKBP.
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DED were established, as previously described (Kawahara et al. 1998). In brief, cells were cotransfected with 50 µg of the respective expression vectors with 1 µg of the hygromycin-resistant gene (pMiwhyg) by electroporation. Cells were cultured in the presence of 0.8–1.0 mg/ml of hygromycin, and the transformant clones expressing FADD-DD, FADD-DED, or FADD-DED were identified by Western blotting using anti-FADD or anti-FLAG antibodies. Jurkat cell transformants expressing high levels of mouse Fas were established by introducing the mouse Fas expression plasmid (Watanabe-Fukunaga et al. 1992).
Cell Viability Assay and FACS® Analysis
Cell viability was determined by the WST-1 assay, as described previously (Kawahara et al. 1998). In brief, Jurkat and JB6 cell transformants (5 x 104 cells in 100 µl) expressing the chimeric proteins, which were made from FADD and FKBP, were treated at 37°C with AP1510. Human Jurkat and its transformant expressing mouse Fas (JmF) were treated with 50 ng/ml LZ-FasL and 1 µg/ml of the Jo2 antibody (Ogasawara et al. 1993), respectively. WST-1 reagents, 2-(4-iodophenyl)-3-(4-nitrophenyl)-5-(2,4-disulfophenyl)-2H-tetrazolium monosodium salt (Dojin Laboratories) and 1-methoxy-5-methylphenazinium methylsulfate (Dojin Laboratories), were added to the cells at final concentrations of 5.0 and 0.2 mM, respectively, and incubated for 1 h at 37°C. Using an ELISA autoreader, the cell viability was determined by measuring the difference between the absorbance at 450 and 620 nm. To examine the effect of various metabolic inhibitors on the cell death, cells were preincubated for 1 h with PDTC, z-VAD–fmk, or both.
The m and the formation of oxygen radicals were determined by staining cells with the Mitotracker orange reagents, CMTMRos, and CM-H2TMRos (Molecular Probes), respectively, as described previously (Bossy-Wetzel et al. 1998). In brief, cells (106 cells/ml or otherwise stated) were incubated at 37°C for 30 min with 25 nM CMTMRos or 100 nM CM-H2TMRos in RPMI1640 containing 10% FCS. Cells were washed with PBS and analyzed on a FACS®Calibur flow cytometer (Becton Dickinson) with excitation at 488 nm, using an argon laser. The fluorescence from CMTMRos was detected with a 585/42 nm band pass filter.
Subcellular Fractionation and Western Blotting
The subcellular fractions were prepared essentially according to Gross et al. 1999. In brief, 2 x 107 cells were washed with PBS, suspended in 150 µl ice-cold isotonic homogenizing buffer (buffer A, 200 mM mannitol, 70 mM sucrose, 1 mM EGTA, 10 mM Hepes-KOH, pH 7.4, 0.1 mM pAPMSF), and homogenized in a glass Dounce homogenizer (Wheaton) with a tight pestle (10 strokes). Unlysed cells and nuclei were removed by centrifugation at 120 g for 5 min. The supernatant was spun at 10,000 g for 10 min, and the pellet, which was resuspended in buffer A, was used as the heavy membrane fraction containing mitochondria. The supernatant was further spun at 100,000 g for 30 min, and the resultant supernatant was used as the S100 fraction.
For Western blotting analysis, samples were mixed with an equal volume of 2x Laemmli's sample buffer. After heating at 95°C for 10 min, proteins were separated by electrophoresis in a 15–25% gradient polyacrylamide gel (Dai-ichi Chemical), and then they were transferred to a polyvinylidene difluoride membrane (Hybond P; Amersham-Pharmacia Biotech). The membrane was blocked with PBST (PBS supplemented with 0.05% Tween 20) containing 5% nonfat dry milk, followed by successive incubations with primary and secondary antibodies. Proteins were visualized with the enhanced chemiluminescence system (Renaissance; NEN Life Science Products Inc.).
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Results |
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Inhibitory Effect of PDTC on Caspase-independent Cell Death
To study the molecular mechanism of the caspase-independent necrotic pathway, we first screened various compounds for the ability to inhibit the FKBP-FADD–induced death of JB6 cells. We found that PDTC inhibits the process in a dose-dependent manner. As shown in Fig. 2 A, AP1510 treatment killed 90% of the cells within 4 h. However, when the cells were preincubated with 80 µM PDTC, 90% of the cells survived for at least 4 h. A similar inhibitory effect of PDTC was observed with JB6 cells expressing FKBP-DED. That is, death was substantially delayed by pretreating the cells with PDTC (Fig. 2 B). PDTC is known to work as an antioxidant (Liu et al. 1996). However, other antioxidants, such as butylated hydroxyanisole (BHA, 250 µM) and nordihydroguaiaretic acid (25 µM), showed little inhibitory effect on the FADD-induced death of JB6 cells (data not shown).
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m was measured by staining with the Mitotracker reagent. As shown in Fig. 3 A, m dramatically decreased within 3 h after the addition of AP1510. This reduction in m was greatly inhibited by preincubation of the cells with 80 µM PDTC, which is consistent with the inhibitory effect of PDTC on cell death. The TNF-induced necrotic cell death of mouse L929 cells is accompanied by formation of oxygen radicals (Vercammen et al. 1998a). To examine whether oxygen radicals are involved in the loss of m of FADD-DED–induced death of JB6 cells, the cells were stained with a reduced version of the Mitotracker reagent (CM-H2TMRos) that emits fluorescence only after oxidation. As shown in Fig. 3 B, the staining intensity with this reagent did not increase after the treatment with AP1510. This was not due to the inability of the reagent to enter mitochondria, because the treatment of the cells with t-butyl hydroperoxide significantly increased the fluorescence in the presence or absence of AP1510.
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m (Susin et al. 1997). Therefore, we examined whether cytochrome c was released from mitochondria during the DED-induced death of JB6 cells. The JB6 cells expressing FKBP-DED were treated with AP1510 for various lengths of time and homogenized with a Dounce homogenizer. After removing the unlysed cells, the cell extracts were fractionated into heavy membrane (precipitates of 10,000 g) and cytosolic (supernatant of 100,000 g) fractions, and then they were analyzed by Western blotting with anti–cytochrome c antibodies. As shown in Fig. 3 C, no cytochrome c was detected in the S100 fraction, even 4 h after the addition of AP1510. Rather, the apparent content of cytochrome c in the heavy membrane fractions increased. The amount of cytochrome oxidase subunit II in this fraction increased similarly in a time-dependent manner. These results may indicate that the relative proportion of mitochondria increased in the heavy membrane fraction because the dying cells were more easily disrupted. Cytochrome c that is released into the cytosol can process procaspase 3 into its mature form through the activation of caspase 9 (Li et al. 1997). In agreement with the observed lack of cytochrome c release from mitochondria, the procaspase 3 remained intact 4 h after the addition of AP1510 (Fig. 3 C).
The Death Signals from the Fas Receptor in Fas-overexpressing Cells
When the endogenously expressed Fas of Jurkat cells is engaged by anti–human Fas antibodies, the cells die, and this process can be blocked by caspase inhibitors. On the other hand, the above results indicated that JB6 cells, a Jurkat-derived cell line, can be killed in a caspase-independent manner when FADD or its DED is tightly aggregated. To examine whether the caspase-independent death signal can be transduced from Fas, Jurkat cell transformants (JmF) that overexpress mouse Fas were established. As shown in Fig. 4, when the transformants were treated with the anti–mouse Fas antibody (Jo2), the cells died. This cell death process was quicker than that observed with JmF cells treated with the anti–human Fas antibody, suggesting that overexpression of mouse Fas and its engagement with the anti–mouse Fas antibody can generate a stronger death signal than that elicited through endogenously expressed Fas. Preincubation of JmF cells with 0.5 mM z-VAD–fmk, but not with PDTC, significantly blocked the anti–mouse Fas-induced death process (Fig. 4). However, PDTC in the presence of z-VAD–fmk had an additional inhibitory effect on the death process in a dose-dependent manner. Thus, preincubation of the cells with 0.5 mM z-VAD–fmk and 80 µM PDTC almost completely inhibited the Fas-induced death of JmF cells. These results indicate that when the caspases are functional, the caspase-dependent death pathway dominates the caspase-independent pathway. However, even when the caspases are not functional, cells die by Fas activation via a caspase-independent mechanism.
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m in the Fas-activated cells was then measured in the absence or presence of z-VAD–fmk and PDTC. As shown in Fig. 5, Fas activation caused a gradual reduction in the m of JmF cells. PDTC had little effect on the kinetics of the m loss. Preincubation of the cells with z-VAD–fmk blocked the loss of m at an early stage of cell death. That is, >80% of the cells showed an intact m 6 h after Fas activation, which agrees with the scenario that most cells were alive at this point (Fig. 4). However, at a later time point (9 h after the addition of anti-Fas antibody), the m of half the cell population was completely lost. This loss of m was blocked by adding PDTC together with z-VAD–fmk. These results suggest that the gradual reduction in m during the Fas-induced cell death process is caspase dependent, whereas the Fas-induced caspase-independent death signal causes a sharper reduction in m that occurs later in the cell-death process.
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Discussion |
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m) is observed in the apoptosis induced by a variety of stimuli (Shimizu et al. 1996; Zamzami et al. 1996) and has been proposed to be essential for the release of cytochrome c and/or apoptosis-inducing factor from mitochondria (Susin et al. 1997). However, even when m was substantially lost during the necrosis of JmF cells, no release of cytochrome c was observed (Fig. 6), suggesting that the loss of m is not sufficient to release cytochrome c from mitochondria. A similar conclusion was reported by Bossy-Wetzel et al. 1998 for the staurosporine-induced cell death. A slight and gradual loss of m during the Fas-induced apoptosis of JmF cells may be a secondary effect caused by the release of cytochrome c.
In contrast to apoptosis, the necrotic pathway functioned independent of the caspases. Even caspase 8–null cells died when FADD was oligomerized. This process was accompanied by a sudden loss of m. Such a loss of m would reduce the cellular ATP level, which may lead to necrotic cell death (Eguchi et al. 1997; Leist et al. 1997). In mouse L929 cells, an excess of oxygen radicals is produced upon activation with TNF and Fas (Vercammen et al. 1998a, Vercammen et al. 1998b). Since BHA, a radical scavenger, blocks the TNF- and Fas-induced necrotic pathway in L929 cells, it was postulated that oxygen radicals are responsible for necrotic cell death. Although reactive oxygens would reduce the m, we could not detect a significant production of reactive oxygen intermediates during the FADD-DED–induced necrosis in JB6 cells, and BHA could not inhibit the death process, suggesting that oxygen radicals are not responsible for the loss of m in our Jurkat system. In contrast, we found that PDTC efficiently inhibited Fas- or FADD-DED–induced necrosis. How PDTC blocks the process is not clear. PDTC is a thiol compound, works as an antioxidant or prooxidant, and has cytotoxic effects (Orrenius et al. 1996). It inhibits the induction of some transcription factors such as NF-
B, whereas it activates other transcription factors such as AP-1, p53, and NF-AT. Although the JNK-AP1 pathway has been suggested for Fas-mediated cell death, we found no significant activation of JNK in FADD-induced necrotic cell death (Shimizu, Y., H. Matsumura, A. Kawahara, and S. Nagata, unpublished observations). Moreover, inhibitors of protein synthesis or RNA synthesis (cycloheximide or actinomycin D) did not inhibit the FADD-DED–mediated necrosis (data not shown), indicating that, as for Fas-mediated apoptosis, all components necessary for necrotic cell death are present in proliferating cells. It is likely that a DED-containing molecule(s) binds to FADD-DED and is activated by oligomerization. In any case, this signaling pathway should lead to the loss of m and should be inhibited by PDTC. Identification of molecules involved in necrotic signal transduction and the development of its specific inhibitor will be necessary to address whether or not the inhibition of loss of m blocks eventual cell death.
Our present results indicate that Fas can activate two distinct death pathways that branch at FADD. Since embryonal fibroblasts from caspase 8–null mice or caspase 8–deficient Jurkat cells are resistant to the Fas-activated death (Kawahara et al. 1998; Varfolomeev et al. 1998), we think that the engagement of Fas usually causes apoptosis. However, when Fas is strongly activated for long periods of time without phagocytosis of the apoptotic cells, the cells may undergo necrotic cell death. In this regard, it is noteworthy that the administration of FasL or agonistic anti-Fas antibodies to animals brings about massive hemorrhagic necrosis in the liver (Ogasawara et al. 1993; Walczak et al. 1999), suggesting that Fas can transduce necrotic death signal under some pathological conditions. Recently, Lauzurica et al. 1999 showed that TNF-
–induced septic shock in vivo can be blocked by pretreating mice with PDTC, suggesting that TNF can also transduce the PDTC-inhibited death signal. The demonstration that FADD-DED is responsible for the necrotic death signal in caspase 8–null cells will assist further studies targeted at identifying other molecules involved in necrotic cell death.
Submitted: 31 May 2000
Revised: 11 October 2000
Accepted: 12 October 2000
A. Kawahara's present address is Laboratory of Molecular Genetics, National Institute of Child Health and Human Development, Building 6B, Room 420, NIH, Bethesda, MD 20892.
m, mitochondrial transmembrane potential.
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) or in the presence of 40 µM PDTC (
), 80 µM PDTC (
), 0.5 mM z-VAD–fmk (•), 40 µM PDTC and 0.5 mM z-VAD–fmk (
), and 80 µM PDTC and 0.5 mM z-VAD–fmk (
). After incubation at 37°C for the indicated time periods, cell viability was determined by the WST-1 assay, as described in Materials and Methods. The results are expressed as the percent of the value obtained without the Jo-2 antibody. The experiments were performed in triplicate, and the average values were plotted.
