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The Ben May Institute for Cancer Research,1 Department of Medicine,2 Department of Pathology University of Chicago, Chicago, Illinois 606373
Received 11 June 2001/ Returned for modification 17 July 2001/ Accepted 9 October 2001
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ABSTRACT |
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INTRODUCTION |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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We have previously described two different CD95 apoptosis pathways (32). In type I cells, caspase 8 is recruited to the DISC, resulting in release of active caspase 8 in quantities sufficient to directly activate caspase 3 (40). However, in type II cells, despite similar expression levels of surface CD95 and signaling molecules, formation of the DISC is so inefficient that only very small quantities of caspase 8 are generated at the cell surface. This amount of caspase 8 is insufficient to process caspase 3, but sufficient to cleave the BH3-only protein Bid (13, 16, 19), resulting in the apoptogenic activation of mitochondria. Therefore, the execution of apoptosis can be inhibited by overexpression of Bcl-2 or Bcl-xL only in type II cells (32). Recently, a number of transgenic and knockout studies have provided evidence for the existence of the two pathways in vivo (14, 17, 30, 41, 48, 49). In all cases, CD95 apoptosis execution of thymocytes and peripheral T cells was independent of mitochondria, identifying them as type I cells, whereas the liver was found to be a type II tissue that requires the mitochondrial branch of the CD95 pathway to apoptose.
We now show that the efficient formation of the DISC found in type I cells precedes formation of CD95 surface clusters and that clustering is dependent on DISC-generated active caspase 8. We also demonstrate for the first time that during CD95-mediated apoptosis, the CD95-CD95L complex is internalized through an endosomal pathway. The data allow us to distinguish four sequential early signaling events of CD95: (i) ligand-induced formation of receptor microaggregates, (ii) actin-dependent assembly of the DISC, (iii) caspase 8-dependent formation of higher-order surface receptor clusters, and (iv) actin-driven receptor internalization. The data indicate that expression of the surface CD95 receptor and of DISC signaling molecules enables cells to produce small quantities of active caspase 8 at their ligated receptor. However, formation of a DISC that generates large quantities of caspase 8, characteristic of type I cells, and internalization of the activated receptor-ligand complex require actin filaments.
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MATERIALS AND METHODS |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Antibodies and reagents.
The monoclonal antibody against FADD was purchased from Transduction Laboratories (San Diego, Calif.). The rabbit polyclonal anti-CD95 (C20) and horseradish peroxidase (HRPO)-conjugated goat anti-rabbit immunoglobulin G (IgG) were purchased from Santa Cruz Biotechnologies (Santa Cruz, Calif.). The C15 monoclonal antibody (MAb) recognizes the p18 subunit of caspase 8 (31), and anti-APO-1 (anti-CD95) is an agonistic MAb (IgG3, 
) recognizing an epitope on the extracellular part of CD95 (44). The HRPO-conjugated goat anti-mouse IgG1 and IgG2b were from Southern Biotechnology Associates (Birmingham, Ala.). The transferrin receptor (TfR) antibody (clone B3/25) was obtained from Roche (Indianapolis, Ind.). The fluorescein isothiocyanate (FITC)-conjugated anti-human CD19 MAb (mouse IgG1) was obtained from Ancell Corp. (Bayport, Minn.). All other chemicals used were of analytical grade and were purchased from Sigma (St. Louis, Mo.), Molecular Probes (Eugene, Oreg.), or Calbiochem (San Diego, Calif.).
DISC analysis by Western blotting. A total of 107 SKW6.4 or H9 cells were treated with 1 µg of anti-CD95 per ml for 5 min or for the times indicated at 37°C and then lysed in lysis buffer (30 mM Tris-HCl [pH 7.5], 150 mM NaCl, 2 mM EDTA, 1 mM phenylmethylsulfonyl flouride [PMSF], protease inhibitor cocktail [Sigma], 1% Triton X-100, 10% glycerol) (stimulated condition). The CD95 DISC was then immunoprecipitated for 2 h at 4°C with protein A-Sepharose (Sigma). After immunoprecipitation, the beads were washed five times with 1 ml of lysis buffer. For Western blotting, immunoprecipitates were separated by SDS-PAGE (12% polyacrylamide) and transferred to Hybond nitrocellulose membrane (Amersham, Piscataway, N.J.), blocked with 2% bovine serum albumin (BSA) in phosphate-buffered saline (PBS)-Tween (0.05% Tween 20) for 1 h, washed with PBS-Tween, and incubated with the primary antibody in PBS-Tween for 16 h at 4°C. Blots were then incubated with HRPO-conjugated secondary antibody diluted 1/20,000 in PBS-Tween. After being washed with PBS-Tween, the blots were developed by the enhanced chemiluminescence (ECL) method according to the manufacturer’s protocol (Amersham).
Induction of apoptosis and cytotoxicity assay. A total of 105 cells in 200 µl of medium were incubated in 96-well plates (Corning, Inc., Corning, N.Y.) in the absence or presence of increasing concentrations of lantrunculin A (Ltn A) for 1 h prior to anti-CD95 stimulation. Cells were stimulated with 1 µg of anti-CD95 per ml (unless otherwise stated) or 1 µg of LZ-CD95L per ml at 37°C by incubation for 16 h. Quantification of DNA fragmentation, as a measure of apoptosis, was carried out by nuclear staining with propidium iodide as previously described (32).
Flow cytometric analysis of mitochondrial membrane potential (
m).
To measure m, anti-CD95 (2 µg/ml)-treated or untreated SKW6-vec or SKW6-Bcl-2 cells (5 x 105/ml) were incubated with 5 µg of JC-1 (5, 5', 6, 6'-tetrachloro-1, 1', 3, 3'-tetraethylbenzimidazolylcarbocyanine iodide) per ml (Molecular Probes). The analysis was performed as previously described (33).
Caspase activity assay. Caspase 8 activity was determined from cell lysates as follows. Cells were stimulated for the indicated times with 1 µg of anti-CD95 per ml. Lysates from 106 cells were incubated in cleavage buffer containing 40 µM amino trifluoromethyl coumarin (AFC)-labeled caspase 8-specific peptide IETD for 1 h at 37°C. Caspase activity was determined with a microplate fluorescence reader with a 400-nm excitation filter and 508-nm emission filter. Values of unstimulated cells were taken as background and subtracted from those obtained with stimulated cells. We have recently established that under the conditions used, caspase 8 activity is measured without any appreciable cross-reactivity by effector caspases such as caspase 3 (39).
Detection of CD95 aggregates. To analyze ligand-dependent SDS-stable CD95 microaggregates, 106 cells were stimulated with 1 µg of anti-CD95 per ml for 20 min at 37°C. Cells were washed once with PBS and lysed in lysis buffer (50 mM HEPES [pH 7.4], 150 mM NaCl, 10 mM NaF, 10 mM iodoacetamine, 200 µM Na3VO4, 1% NP-40). Postnuclear supernatants were analyzed by SDS-PAGE (8% polyacrylamide), and CD95 was detected with the antibody C20.
Internalization assay and immunofluorescence. To monitor the fate of CD95 after stimulation, 106 cells/ml were treated with 1 µg of anti-CD95 per ml for 45 min on ice. Unbound antibody was removed by washing with medium at 4°C. Cells were then stained with 1 µg of FITC-conjugated goat anti-mouse IgG per ml for 45 min on ice. Next, cells were washed, warmed, and kept at 37°C for the indicated time points to trigger CD95 stimulation or were kept on ice (time [t] = 0). In some experiments, FITC-conjugated anti-CD95 antibody was used as indicated in the figure legends. After stimulation, cells were adhered to immunofluorescence slides (Polysciences, Warrington, Pa.) precoated with poly-L-lysine (Sigma) and fixed in acetone-methanol (1:1) for 5 min at room temperature. Slides were washed with PBS, rinsed with tap water, mounted with Vectashield (Vector Laboratories, Burlingame, Calif.), and visualized with a Zeiss Axiovert S100 fluorescence microscope equipped with an Axiocam digital camera or with a confocal microscope (Zeiss, LSM 510).
For ligand stimulation, cells were treated with 1 µg of LZ-CD95L per ml for 45 min on ice and stimulated at 37°C as described above. After stimulation, cells were adhered to immunofluorescence slides (Polysciences) precoated with poly-L-lysine (Sigma) and fixed in acetone-methanol (1:1) for 5 min at room temperature. Fixed cells were stained with anti-CD95L antibody (clone G247-4) (Pharmingen, San Diego, Calif.) followed by goat anti-mouse FITC (1:200 dilution in PBS). After staining, slides were washed with PBS and mounted with Vectashield (Vector Laboratories). For costaining with the endocytic marker transferrin receptor (TfR), cells were stained for CD95 as described above, fixed, and then incubated with the anti-TfR antibody (IgG1) (1:200 dilution in PBS) for 45 min. After washing, a Texas red-labeled goat anti-mouse IgG1 antibody (which did not cross-react with the anti-CD95 antibody [IgG3]) was applied and incubated for 45 min.
To distinguish between surface and internalized CD95, 106 cells/ml were incubated with 1 µg of FITC-conjugated anti-CD95 antibody per ml for 45 min on ice. After being washed, the cells were resuspended in 37°C medium and incubated at 37°C for the indicated times or kept on ice (time 0). Cells were then resuspended in 300 µl of cold 2% BSA-PBS, stained with Texas red-conjugated goat anti-mouse IgG for 30 min on ice, washed, and resuspended in serum-free medium. The cells were then plated for 5 min at 37°C on poly-L-lysine (Sigma)-coated slides and fixed in methanol-acetone (1:1). Slides were washed and rinsed with water, and coverslips were mounted with Vectashield mounting medium. For quantification of internalization, 100 cells for each condition per cell line were counted by fluorescence microscopy. Cells with 50% or more of CD95 structures stained only green (FITC-anti-CD95) were scored as having internalized the receptor.
Cholesterol depletion studies. SKW6.4 or H9 cells were pretreated with the following cholesterol-depleting agents: 1 µg of filipin per ml, 10 µg of nystatin per ml, or 2 mM methyl-ß-cyclodextrin in serum-free medium for 1 h at 37°C. Thereafter, DISC formation and CD95 clustering were analyzed as described above. For induction of apoptosis, cells were pretreated with the cholesterol-depleting agents as described above and incubated with increasing concentrations of anti-CD95 for 9 h (cells in serum-free medium) or for 16 h (cells in serum-containing medium). Apoptosis was quantified by DNA fragmentation by nuclear staining with propidium iodide as previously described (32).
Electron microscopy. For electron microscopy analysis, 3 x 106 cells were resuspended in 300 µl of medium and stained with 6 µg of anti-CD95 antibody for 45 min at 4°C. The cells were then washed and stained with 3 µg of 20-nm-diameter gold particle-conjugated goat anti-mouse IgG (Ted Pella, Inc., Redding, Calif.) for 1 h at 4°C. Cells were washed and kept on ice (t = 0) or resuspended in warm medium and incubated at 37°C for 20 min. After incubation, cells were washed with serum-free medium followed by PBS and fixed with 2% glutaraldehyde buffered with 0.1 M Na cacodylate (pH 7.35) for 10 to 15 min at room temperature. Fixed cells were gently centrifuged, and excess fixative was removed and replaced. Fresh fixative was added to the tube without disturbing the pellet or cell layer, and fixation was continued for 10 min at room temperature. The tube was then transferred to 4°C and centrifuged at 10,000 x g for 10 min to create a tight pellet. This pellet was then stored overnight at 4°C until further processing. Portions of the pellet were washed several times with 0.1 M Na cacodylate (pH 7.35) and postfixed for 1 h on ice in freshly prepared 1% osmium tetroxide-1% potassium ferrocyanide-0.1 M Na cacodylate (pH 7.4). After being washed, cells were stained en bloc in 2% uranyl acetate dehydrated through a graded series of ethanol and embedded in Epon resin. Sixty- to 70-nm-thick sections were cut with a Reichert Ultracut-E microtome and stained for 5 min with 1.5% uranyl acetate, followed by 10 min in Reynolds lead citrate. Samples were visualized with a JEOL Philips CM-120 electron microscope operated at 60 kV.
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RESULTS |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Stimulation-induced clustering is specific for CD95 and can be detected in a variety of different cells. Although clustering of CD95 was a late event when compared to formation of the DISC, most cells displaying CD95 clustering were viable, since they excluded the viability dye propidium iodide (Fig. 2A). This experiment also demonstrated that the clustering of CD95 that we detected was not a fixation artifact, since it was observed in living cells after addition of a directly FITC-conjugated anti-CD95 antibody to SKW6.4 cells in culture medium (Fig. 2A).
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Since our data are in contradiction to two recent reports that detected clustering of CD95 as early as 30 s after receptor triggering (6, 9), we tested other cell lines to exclude the possibility that the effect was limited to SKW6.4 cells. We found similar kinetics of CD95 clustering in the lymphoid cell lines K50 and H9 (Fig. 2C) and in two cell lines of nonlymphoid origin, L929 and MCF7, both transfected with human CD95 (Fig. 2D). In most cases, maximal CD95 clustering was detected 60 min after triggering. CD95 clustering peaked at 20 min only in detached L929-Fas cells. In none of the cell lines could clustering be observed earlier than 5 min of stimulation with either anti-CD95 or LZ-CD95L (data not shown). CD95 did not cluster at all on one cell line, BoeR, which expresses large quantities of CD95 on the cell surface and is resistant to CD95-mediated apoptosis (11, 33) (Fig. 2C, last column), again confirming that clustering of CD95 was not artificially induced by adding a cross-linking antibody. This result also suggested that clustering of CD95 was an active process that does not occur in a cell line that is resistant to CD95-mediated apoptosis.
A redistribution of CD95 was also observed in adherent L929-Fas cells (Fig. 2D, first column). However, we found detached L929-Fas cells to be much more sensitive to CD95-mediated apoptosis, and under these conditions, the effect of receptor clustering was more pronounced. The same was true for MCF7 cells stably expressing CD95 (data not shown) (Fig. 2D, right column). Again, the clustering effect of CD95 was not caused by the fixation method, since it was also observed in living L929-Fas cells stimulated with directly FITC-labeled anti-CD95 (Fig. 2D). In summary, stimulation of all tested cell lines with either anti-CD95 or LZ-CD95L resulted in intense clustering of CD95 that in all cases followed formation of the DISC, revealing an unexplained delay between formation of the DISC and clustering of CD95.
Ligand-induced clustering of CD95 is followed by receptor internalization. In many of the cell lines tested, the punctate pattern of CD95 labeling after ligation appeared to be cytoplasmic rather than clustered on the cell surface. In adherent L929-Fas cells, the observed juxtanuclear staining was particularly indicative of receptor internalization (Fig. 2D). Internalization of CD95 has not yet been reported. A fluorescence-activated cell sorting-based assay as well as many of the typical methods to quantify of receptor internalization (20) proved unreliable in the case of CD95, due to its intense aggregation properties (data not shown). We therefore developed a method to qualitatively determine whether CD95 was on the cell surface or intracellular and to quantify this effect by counting single cells. To this end, living cells were incubated with directly FITC-conjugated anti-CD95 and incubated for different times to induce receptor clustering. After washing, cells were stained with Texas red-conjugated goat anti-mouse IgG. Surface expression was then evaluated by fluorescence microscopy after fixing the cells (Fig. 3A). At the zero time point, CD95 was evenly distributed on the surface of the cells. Both the green (all CD95) and the red (only surface CD95) completely colocalized, indicating that CD95 was at the cell surface. After 20 min, CD95 formed clusters, and at the applied resolution, all clusters were stained in red with the goat anti-mouse antibody, indicating that most CD95 was at the cell surface. At 60 min, however, the majority of the cells showed only a green staining, indicating that CD95 on these cells was inaccessible from the cell surface and suggesting that it had been internalized (arrowheads in Fig. 3A). To further test for the intracellular localization of activated CD95, we used a confocal microscope to generate three-dimensional (3D) images of these cells. In Fig. 3B, 2D projections of such 3D images of representative cells are shown at different stages of CD95 internalization. It is apparent that during the course of the incubation, CD95 moved from the cell surface (red and green = yellow staining) to the interior of the cell (green staining). We then used this assay to quantify the extent of CD95 internalization in different cell lines (Fig. 3C). After 60 min, approximately 70% of the SKW6.4 and K50 had internalized CD95.
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The finding that the internalization of CD95 could be prevented by inhibiting actin filaments prompted us to test whether Ltn A would also have an effect on the formation of the DISC and subsequent generation of active caspase 8. Ltn A inhibited recruitment of FADD (Fig. 5D) and caspase 8 (data not shown) to the DISC of SKW6.4 and H9 cells, resulting in severely reduced activation of caspase 8 at the two concentrations of Ltn A used in the experiments (Fig. 5E). To test whether inhibition of receptor clustering or internalization would affect signaling through CD95, we induced apoptosis in SKW6.4 and H9 cells by stimulation with either anti-CD95 or CD95L in the presence of increasing concentrations of Ltn A and quantified DNA fragmentation (Fig. 5F). Ltn A inhibited CD95-mediated apoptosis, resulting in up to 50% inhibition of DNA fragmentation (Fig. 5F). These data suggest that an intact actin cytoskeleton is required for formation of the DISC as well as for internalization of CD95 in these cells, but do not exclude the possibility that actin is also involved in processes further downstream in the pathway.
Clustering of CD95 is an active process that depends on activation of caspase 8 at the DISC.
The results observed with the apoptosis-resistant BoeR cells and the kinetics of CD95 clustering in many cells suggested that binding of anti-CD95 antibodies or CD95L to CD95 alone is not sufficient to induce receptor clustering. To test whether the initiation of the caspase cascade at the DISC was required to induce clustering of CD95, we tested BJAB cells stably expressing a dominant-negative version of FADD (FADD-DN) that is recruited to the CD95 DISC, but is unable to bind to procaspase 8 (5). Triggering CD95 in the BJAB-vec cells resulted in clustering of CD95 (Fig. 6A). In contrast, FADD-DN-transfected BJAB cells did not show any signs of CD95 clustering. These data suggest that caspase activation is required to induce receptor clustering. In the CD95 pathway, caspases are activated at two steps, upstream or downstream of mitochondria. We have previously shown that the type I SKW6 cells expressing Bcl-2 activate caspases in a way that is absolutely independent of mitochondrial functions (32). Like SKW6.4 cells, these transfectants responded to triggering through CD95 with receptor clustering (Fig. 6B) and internalization (data not shown). The intensity and kinetics of this process were identical to those of the vector-transfected control (SKW6-vec). In contrast to the vector control, we could not detect a drop in the mitochondrial transmembrane potential (m) in the SKW6-Bcl-2 cells (Fig. 6C), demonstrating that Bcl-2 fully prevented mitochondrial dysfunction in these cells. We can therefore exclude a mitochondrial contribution to the clustering and internalization of CD95 in these cells, suggesting that a DISC-generated caspase was regulating receptor clustering. To test whether caspase 8 regulates CD95 clustering, we used MCF7-Fas cells, which do not express caspase 3 and activate only caspase 8 within the first 4 h of receptor stimulation (39). In MCF7-Fas cells, CD95 efficiently clustered following stimulation (and was internalized [Fig. 4A and B]) (32). We again chose to analyze detached MCF7 cells, since we found that they activate caspase 8 much more efficiently than adherent cells (data not shown). Addition of the poly-caspase inhibitor zVAD-fmk prevented receptor internalization in these cells. To test whether an IETD (caspase 8)- and not a DEVD (caspase 3)-cleaving activity was involved in receptor internalization, we compared the effects of zIETD-fmk with those of zDEVD-fmk (Fig. 6D). The caspase 8-selective inhibitor blocked receptor internalization at 20 µM, whereas the caspase 3-selective inhibitor did not inhibit this process at concentrations as high as 50 µM. Inhibition of caspase 8 also efficiently inhibited receptor clustering in other cells, such as SKW6.4, K50, and H9 (data not shown). Taken together, all data suggest that early activation of caspase 8 at the DISC regulates clustering of CD95, which is a prerequisite for receptor internalization in these cells.
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DISCUSSION |
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Involvement of the actin filament system in the ligand-induced internalization of many receptors is well documented (28). Ltn A, a toxin isolated from a Red Sea sponge, disrupts microfilament organization in cultured cells by binding to monomeric G-actin in a 1:1 complex at submicromolar concentrations (38) and is a widely used inhibitor of receptor-mediated endocytosis (15, 18). Ltn A has been shown to inhibit CD95-mediated apoptosis in certain cells (42), suggesting that actin is involved in signaling through CD95. Recently, we reported a role for actin in regulating apoptosis sensitivity of restimulated Th2 cells (46). However, under these conditions, polymerized actin impaired CD95-mediated apoptosis by inhibiting lateral diffusion of the receptor and consequently cytochalasin D-sensitized apoptosis-resistant Th2 cells. In contrast to this role of the actin cytoskeleton in maintaining the CD95 apoptosis resistance of restimulated Th2 cells, we now report a general role for actin in regulating formation of the DISC and internalization of CD95 in all tested apoptosis-sensitive type I cells.
Formation of the DISC in type I cells is detectable seconds after receptor triggering (22, 31). Since type II cells form almost no DISC, inhibition of actin polymerization does not affect generation of caspase 8 in these cells (data not shown). Consistent with this finding, Ltn A could not completely block generation of active caspase 8 or apoptosis in type I cells. The cells still died with mitochondrial signal amplification, reminiscent of the situation found in type II cells (32).
CD95 has previously been shown to form caps as a result of receptor triggering. In one report, capping of the receptor was shown to occur between 40 and 60 min after triggering, consistent with our data (47). However, two recent reports showed that CD95 caps in a ceramide-dependent fashion as early as 30 s after receptor triggering (6, 9). These two studies were mainly done with type II cells—Jurkat cells and hepatocytes. We did not see a significant difference in the kinetics of CD95 clustering between the type I cells used in our study and the type II Jurkat and CEM cells. However, the CD95 clusters in type II cells were smaller, and clustering could not be inhibited by caspase inhibitors (unpublished results). Localization of CD95 to lipid rafts was required in these studies for efficient signaling. Most of the CD95 receptor clusters detected in our study represent internalized receptor. Only when internalization of CD95 was blocked by treating cells with Ltn A did we detect intense clusters of CD95 on the cell surface (Fig. 5A and B). However, under these conditions recruitment of FADD to the DISC, activation of caspase 8, and the efficiency of apoptosis signaling were all severely impaired (Fig. 5D to F). This is not consistent with the recent reports describing the necessity of recruitment of CD95 and its signaling molecules to clusters in lipid rafts (6, 9). Consequently, pretreatment of cells with the same cholesterol-depleting reagents used in these studies did not affect signaling in the tested cells (Fig. 7).
These discrepancies suggest a different mechanism of signal initiation between type I and type II cells. Our original finding is consistent with this assumption—only type II Jurkat and CEM cells are sensitive to C2-ceramide-induced apoptosis, whereas the type I SKW6.4 and H9 cells are completely resistant (33)—as well as with our recent finding that type II Jurkat and CEM cells do not internalize CD95 upon stimulation (data not shown). These findings could provide an explanation for the contradiction with the two recently published reports on CD95 capping. We are currently investigating the differences in receptor clustering and internalization between type I and type II cells. Alternatively, the differences between our study and two recently published reports (6, 9) could be due to differences in the methods and reagents used to trigger CD95 and/or to detect the capped receptor. In these two reports, cells were stimulated with CH11 (IgM) or sCD95L, and capped CD95 on fixed cells was visualized by labeled secondary antibodies. In contrast, we stimulated cells with anti-CD95 (IgG3) or LZ-CD95L. This stimulation also resulted in CD95 receptor clustering in living cells when a primary labeled anti-CD95 antibody was used.
It has been suggested that the use of higher concentrations of CD95-stimulating agents would artificially cluster CD95 on cells (9), producing an unphysiological triggering of CD95. We have set the titer of our anti-CD95 antibody down to 30 ng/ml with no loss in the ability to induce clustering of the receptor after stimulation. Moreover, the fact that clustering of CD95 could completely be prevented by pretreating cells with caspase inhibitors confirms that CD95 clustering in our experiments is an active process that follows the triggering of the receptor and activation of caspase 8. This is also supported by the fact that the CD95 on the CD95high-expressing BoeR cells does not form clusters or a DISC (33) and does not internalize (data not shown) regardless of how much antibody is added. We can, however, detect SDS-stable microaggregates in these cells (data not shown) after addition of anti-CD95, confirming that the anti-CD95 binds and triggers aggregation of CD95.
No previously published report has tested whether activated CD95 internalizes after clustering. We now demonstrate that triggering CD95 results in receptor internalization shortly after engagement of the receptor by either anti-CD95 MAb or CD95L. Internalization of CD95 was found in lymphoid and nonlymphoid cells, in suspension as well as in adherent cells. Although stimulation-induced receptor internalization of CD95 has not been previously reported, CD95 can be forced to internalize. It has been shown that the adenoviral proteins E3/10.4K to -14.5K induce internalization of CD95 followed by its lysosomal degradation (7, 43). This mechanism was interpreted as a viral strategy to protect its infected host cell through down-modulation of CD95.
The endocytosis machinery is a complex network of different pathways. Cargo can be endocytosed via clathrin-coated pits, by the clathrin-independent pathway, or through caveolae (21). All of these pathways lead to the appearance of cargo in the peripheral early endosomes. We have not yet observed CD95 in coated vesicles in our transmission electron microscopy analysis. However, internalization of CD95 could be prevented by depleting K+, suggesting a mechanism of internalization that depends on coated pits (data not shown). We detected colocalization of internalized CD95 with the lysosomal marker Lamp-1 in some but not all cells (data not shown). Future experiments will determine which pathway of endocytosis is used by the internalized CD95 receptor.
Receptor internalization plays an important role in desensitizing cells to specific extracellular signals and in recycling receptor and/or ligand molecules. Down-modulation of receptors on cells that have received a death stimulus, although seemingly counterproductive, may actually be important physiologically. CD95L bound to surface CD95 may be harmful to neighboring nonapoptosing cells or to macrophages that attempt to engulf apoptotic cells. It may therefore be advantageous and protective for the organism for an apoptosing cell to internalize the death receptor-ligand complex. Growing evidence suggests that the process of receptor internalization is also required to colocalize activated receptors with downstream signaling molecules (reviewed in reference 4). Internalization of a DISC actively producing caspase 8 may therefore be important for targeting the caspase to its intracellular substrates. Future experiments aim to further dissect the initial CD95 signaling events by separating the DISC formation phase from the internalization phase.
Alternatively, activation of caspase 8 does not invariably lead to apoptosis (23). Therefore cells could receive a rescue signal after triggering of CD95 and the surviving cells could potentially recycle internalized CD95 to the surface. We recently showed that MCF7-Fas cells are protected from apoptosis by stable expression of Bcl-xL and that none of the cytosolic caspase substrates is cleaved. However, clustering of CD95 and its internalization occurs and can be prevented by pretreating cells with zIETD-fmk (A. Stegh, B. C. Barnhart, A. Algeciras-Schimnich, and M. E. Peter, submitted for publication), suggesting that very small quantities of active caspase 8 are sufficient to initiate this process. The finding that nonapoptosing cells also internalize CD95 after CD95 stimulation also demonstrates that internalization of CD95 is not just an unspecific event due to changes in the membrane of apoptosing cells. The data rather support our model of both clustering and internalization as active processes that are regulated by activation of caspase 8.
It has recently been demonstrated that another member of the death receptor family, TNF-RI, internalizes upon binding to its ligand TNF-
(36). In this report, the authors demonstrated differences between TNF-RI and CD95. Internalization of TNF-RI was inhibited by the transglutaminase inhibitor MDC. Similar to our study, inhibition of internalization of TNF-RI also inhibited apoptosis induction by the receptor. This inhibition was selective for TNF-RI, since, consistent with our data, MDC did not affect CD95 signaling (36; data not shown).
Interestingly, internalization of TNF-RI cannot be prevented by inhibiting caspases (M. Kroenke, personal communication), confirming that the receptors use different mechanisms for internalization. To our knowledge, CD95 is the first receptor the internalization of which requires activation of caspases. The fact that CD19 was not internalized upon triggering of CD95 suggests that CD95 is internalized in a ligand- and caspase-dependent fashion. However, this does not exclude the possibility that surface molecules in general specifically internalize in a caspase-dependent fashion without being directly stimulated. Activation of caspases could trigger clustering and internalization of receptors, including CD95. Such a mechanism could explain how agents that induce apoptosis by activating caspases could cause clustering of CD95 in a ligand-independent fashion, as has been shown for antitumor ether lipids, UV radiation, and the herpes simplex thymidine kinase-ganciclovir suicide gene therapy system (1, 2, 8).
Caspases may not directly regulate the internalization step of CD95 signaling, since caspase activity is already required for formation of CD95 surface clusters. Caspase-induced surface clustering could, however, be a prerequisite for internalization of CD95. Our data favor caspase 8 as the caspase regulating this process, because MCF7-Fas cells, which do not express caspase 3 and mainly activate caspase 8 during the first 4 h after CD95 triggering (39), very efficiently internalize CD95. Furthermore, the caspase 8-selective inhibitor zIETD-fmk inhibited receptor internalization much more efficiently than the caspase 3- or 7-selective inhibitor zDEVD-fmk. Ligand-induced surface clustering of CD95 followed by internalization of the CD95 DISC may therefore depend on proteolytic degradation of a death substrate by DISC-generated caspase 8, resulting in remodeling of cortical actin. It is unlikely that gelsolin, a regulator of actin dynamics that has been shown to be a caspase substrate (12), triggers the events described in this report, because gelsolin was shown to be cleaved by caspase 3, which is not expressed in MCF7 cells.
Our study has revealed that the signal initiation by CD95 is much more complex than originally expected. The process can now be described as five distinct steps (Fig. 9). First, CD95 resides on the plasma membrane in a preassociated form through its PLAD domain (Fig. 9, step I). Homophilic association through this domain was shown to be a requisite step in recruiting signaling proteins to the DD (37). The next step in CD95 signaling is formation of SDS-stable CD95 microaggregates, as detected by SDS-PAGE (Fig. 9, step II). This step is not yet dependent on activation of caspases or actin filaments, since it is not inhibited by zIETD-fmk or Ltn A. At this stage, the aggregates cannot be seen as receptor clusters by immunofluorescence microscopy and must therefore occur locally as microaggregates. Both kinetics and inhibitor studies suggest that the next step in CD95 signaling is formation of the DISC (Fig. 9, step III). In type I cells, the first recruitment of FADD can be detected seconds after receptor engagement (8). This step requires actin filaments, since it can substantially be inhibited by Ltn A. DISC formation precedes formation of receptor clusters that can be detected by immunofluorescence analysis (Fig. 9, step IV). Surface clusters of CD95 can be observed beginning at 15 min, followed by internalization, which in most cells is complete at around 30 to 60 min after binding of anti-CD95 antibody or CD95L. After pretreatment with Ltn A, surface clustering seems to be intensified, likely due to the absence of actin filaments and the inhibition of receptor internalization. Inhibition of caspase 8 completely blocks CD95 surface clustering, demonstrating that it is an active process and providing an explanation for the delay between formation of the DISC and CD95 surface clustering. The final step of the events at the plasma membrane involves internalization of the activated DISC (Fig. 9, step V). This step, as with many other cases of receptor internalization, can be blocked by inhibiting actin with Ltn A. Future work will be directed at elucidating the mechanism by which actin facilitates formation of the DISC, the identification of the caspase 8 substrate that regulates clustering of CD95, the physiological role of the internalization of activated CD95, and the contribution of the internalized DISC to the execution of apoptosis once inside the cell.
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ACKNOWLEDGMENTS |
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This work was funded by NIH grant GM61712. A. A.-S. is supported by Cancer Biology Training Program 5T32CA09594. B.C.B. is supported by Molecular and Cell Biology Training grant 5T32GM07183.
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FOOTNOTES |
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REFERENCES |
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) or LZ-CD95L (
) per ml for different times as described for panel A, and cells were scored for clustering. One hundred cells per condition were counted in duplicate, and the mean is shown. (E) SKW6.4 cells were treated with different amounts of anti-CD95 for 60 min, and clustering of CD95 was quantified as in panel D.
) transfectants were treated with 1 µg of anti-CD95 per ml for different periods of time. 
