G. W. Hooper Foundation & Department of Microbiology and Immunology, University of California, San Francisco, CA 94143-0552, USA
Correspondence to: A. L. DeFranco
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Abstract |
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Keywords: apoptosis, BCR, Fas, growth arrest, immature B lymphocytes
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Introduction |
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Fas is known to be essential for eliminating potentially auto-reactive lymphocytes, such as chronically stimulated T cells (9,10) or B cells that are activated via CD40 by T cells but do not react with foreign antigens (1113). The potential role of Fas-induced apoptosis in modulating the selection of developing B or T cells, however, remains less well defined. Initial studies using Fas-deficient mice observed no defect in T cell central tolerance but later studies suggested that Fas has a subtle role in antigen-driven thymocyte selection (1416). Along the B cell lineage, transitional (late immature) B cells were shown to express Fas (17,18). Although BCR-induced apoptosis appears to be normal in Fas-deficient MRL/lpr mice (19,20), auto-antibodies are present in these mice, indicating a role of Fas in eliminating autoreactive B cells. It is unclear whether Fas may functionally participate in B cell selection at some point during B cell development. It is noteworthy in this regard that transitional B cells, expressing Fas, have been shown to be a major stage for negative selection (17,21,22).
The molecular mechanisms of Fas-induced apoptosis are much better characterized than those of BCR- or TCR-induced cell death (23,24). Upon ligation by its trimeric ligand, Fas, as well as other members of the death receptor family, induces apoptosis primarily through assembly of a protein complex including Fas, the adaptor protein FADD and caspase-8 or -10 (23,24). The assembly of this complex leads to the cleavage and activation of recruited caspases. In addition to caspase activation, Fas ligation activates other intracellular events, such as the Jun N-terminal kinase (JNK/SAPK) pathway (25,26). Fas signaling is negatively regulated by FLIP, Btk and probably other proteins as well (2729). Fas-induced apoptosis can also be counteracted by the pro-survival members of Bcl-2 family proteins (30). In contrast, the mechanisms of BCR- or TCR-induced death are not well established. Antigen receptor ligation activates multiple signaling pathways, such as those involving phospholipase C/protein kinase C/Ca2+, phosphotidylinositol-3-kinase, Ras and Rho family GTPases (for reviews, see 31,32). However, it is not clear which of these signaling events activate the apoptotic program. To understand how BCR signaling induces cell death, the murine lymphoma cell line WEHI-231 has been a frequently employed model system. These cells exhibit major biochemical and functional markers of immature B lymphocytes (33) and readily undergo growth arrest and cell death in response to BCR stimulation (34,35). Specific pathways mediating apoptotic induction by the BCR remain to be understood, but may involve the down-regulation of c-Myc expression and NF-B activity (36,37). Activation of caspases by BCR stimulation has been reported in this and other similar in vitro systems (3844). However, the caspase(s) that is first activated by the BCR signaling remains unknown, as is the relationship of BCR-induced growth arrest to BCR-induced caspase activation and apoptosis.
During our studies of BCR-induced apoptosis of WEHI-231, we identified a subclone of this cell line that retained the apoptotic response to BCR stimulation, but acquired susceptibility to Fas-induced apoptosis. We report here the characterization of the apoptotic responses of these cells to stimulation via the BCR and Fas, and the effects of CD40 stimulation on these responses. Our results revealed a functional interplay among these signals different from that seen in mature B cells. We further demonstrated that BCR-induced growth arrest in WEHI 231 cells is a reversible event and is separable from the apoptotic death induction, a process inhibited by the caspase inhibitor BOC-Asp-FMK (BD). The direct comparison of BCR- and Fas-induced apoptosis reported here represents the first (to our knowledge) carried out in the molecular context of a mouse immature B cell line. Finally, a subset of the splenic immature B cells was found to express Fas but they failed to exhibit Fas-mediated apoptosis in vitro.
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Methods |
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Plasmids and cDNAs
The cDNA of the dominant-negative-acting truncation mutant FADD/Mort1 (dnFADD) containing amino acids 81205 of the mouse FADD/Mort1 and anti-FADD/Mort1 rabbit antiserum were kindly provided by Dr Zhang of Dr A. Winoto's laboratory (University of California at Berkeley). The mutant cDNA sequence as a BamHI fragment (45) was subcloned into a mammalian expression plasmid following the promoter of human elongation factor 1a (nucleotide 3731561, GenBank accession no. J04616) and the plasmid also confers hygromycin B resistance by the hygroR cassette from p3'SS (Stratagene, La Jolla, CA). The plasmid DNA was introduced into WM cells by electroporation (200 V, 960 µF, Gene Pulser; BioRad, Hercules, CA). Hygromycin B-resistant clones were obtained and examined for dnFADD expression using the anti-FADD/Mort1 antiserum.
Cells and cell culture
The murine B cell line WEHI-231 and its variant WM were cultured in RPMI 1640 medium supplemented with 5% FCS, 2 mM sodium pyruvate, 1 mM glutamine and 50 µM 2-mercaptoethanol. Cultures were maintained at 15x105 cells/ml. WM cells were initially isolated as one of many clones showing relatively high sensitivity to BCR-induced cell death compared with long-passaged parental WEHI 231 cells. They were later found incidentally to be susceptible to Fas-induced apoptosis as well.
Cell stimulation and assay for viable cells
The viable cell assay used in this study measured the number of viable cells by propidium iodide (PI) exclusion in a stimulated culture and then compared it to the number of viable cells in the control culture (identically seeded at the beginning of the experiment) as a percentage. This assay was designed to examine the overall effects of BCR stimulation which include both growth arrest and cell death, while avoiding counting cell fragments or apoptotic bodies by the flow cytometer as apoptotic cells. To distinguish growth arrest and apoptosis, special attention was given to the ratios of dead versus viable cells within each population. For experiments examining effects of caspase inhibitors, such as those in Fig. 3, the BCR- or Fas-stimulated cultures in the presence of the inhibitors were compared with the unstimulated cultures containing the respective inhibitors.
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Annexin V staining for phosphatidylserine on the cell surface
To examine the externalization of phosphatidylserine as an early apoptotic marker, the Annexin VFITC apoptotic detection kit (Oncogene, Cambridge, MA) was used. The staining was carried out as recommended by the manufacturer (RAPID Annexin V binding procedure). The stained cells were resuspended in PBS with 2% FCS and 1 µg/ml PI (propidium iodide), and examined by flow cytometry with a FACScan (Becton Dickinson, San Jose, CA).
Immunoblotting
Stimulated cells were pelleted by centrifugation, washed with cold PBS, and lysed on ice for 5 min in ice-cold lysis buffer consisting of 20 mM Tris (pH 8.0), 140 mM NaCl, 10% glycerol, 2 mM EDTA, 1% Triton X-100, 1 mM Na3VO4, 10 mM NaF, 1 mM EGTA, 1 mM PMSF, 1 mM aprotinin and 1 mM leupeptin. Following high-speed centrifugation to remove detergent-insoluble material, the protein concentration of the lysate was determined using the bicinchoninic acid assay (Pierce, Rockford, IL). Lysates containing 50 µg protein were subjected to SDSPAGE and separated proteins were transferred onto an Immobilon-P (PVDF) membrane. The membrane was blocked with 3% dried non-fat milk or 3% BSA in Tris-buffered saline with 0.5% Tween 20 (TBST) and incubated with a primary antibody (1 µg/ml) for >1 h at room temperature. After washing the membrane, a horseradish peroxidase-coupled secondary antibody was added for an additional 1 h incubation. After extensive washes, the membrane was subjected to Renaissance chemiluminescence detection (NEN Life Science Products, Boston, MA) and subsequently exposed to autoradiographic films.
Immunostaining
Cells (510x105) were resuspended in 0.5 ml staining buffer (PBS/1% BSA/0.1% sodium azide) with a primary antibody (12 µg) and rotated at 4°C in the dark for 30 min. Cells were pelleted, washed once with staining buffer and resuspended in 0.5 ml staining buffer with the secondary agents (fluorochrome-conjugated secondary antibody or streptavidin). After 30 min binding at 4°C in the dark, cells were pelleted, washed twice with staining buffer and resuspended in 0.5 ml staining buffer. PI was added at 1 µg/ml if necessary. Cells were then analyzed by flow cytometry with a FACScan after compensation was adjusted with unstained and singly stained samples. Acquired data were analyzed with CellQuest software.
Cell cycle analysis
BrdU (Sigma) was added to the medium to a final concentration of 10 µM and the cultures were incubated at 37°C for an additional 30 min. Cells (510x105) were pelleted by centrifugation, resuspended in 100 µl PBS and 100 µl 1% paraformaldehyde in PBS was then added. Following a 20 min incubation on ice, cells were pelleted by centrifugation and resuspended in 50 µl PBS/1% BSA, and 0.5 ml 3 M HCl with 0.5% Tween 20 was then added. After 20 min on ice, cells were pelleted and resuspended in 0.5 ml 0.1 M disodium tetraborate (pH 8.5) to neutralize the residual HCl. Cells were pelleted again, washed once with PBS/1% BSA, resuspended in 100 µl PBS/1% BSA with an additional 20 µl anti-BrdUFITC (Becton Dickinson, San Jose, CA) and incubated on ice for 20 min. Cells were then washed twice, resuspended in 0.5 ml PBS/1% BSA with 1 µg/ml PI and analyzed by flow cytometry.
Mouse splenocyte preparation and in vitro culture
Spleens of BALB/c-J mice (Jackson Laboratory, Bar harbor, ME) were ground against a cell strainer (70 µm) with a syringe plunger and the pass-through cell suspension was centrifuged and resuspended in red blood cell lysis buffer (ammonium chloride-based; Sigma, St Louis, MO). After 5 min incubation at room temperature to facilitate lysis of red blood cells, cells were centrifuged and washed twice with cold PBS. Finally, cells were resuspended in regular culture medium at 1x106/ml for in vitro culture experiments or in PBS with 1% BSA for antibody staining for flow cytometry.
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Results |
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BCR-induced apoptosis was not affected by dnFADD in WM cells
We were interested in understanding if BCR-induced apoptosis may utilize death domain-mediated pathways employed by Fas and other death receptors. For example, the BCR could induce the expression of a ligand for a death receptor on the B cells, as occurs in T cells (48,49). FADD is critically involved in apoptotic induction by the death receptors such as Fas and tumor necrosis factor (TNF) receptor (TNFR)1 (24,50). It interacts with Fas through its death domain and with pro-caspase-8 through its death effector domain. A truncated version of FADD containing the death domain but lacking the death effector domain can compete with the endogenous FADD and act as a dominant interfering molecule for FADD-dependent apoptosis (45,51). To determine if FADD-mediated events participate in BCR-induced apoptosis, a truncated dnFADD was introduced via electroporation into WM cells. Individual transfectant clones were examined for expression of dnFADD by anti-FADD immunoblotting. Four transfectant clones which expressed dnFADD at levels varying by <2-fold (data not shown) and two transfectant clones which did not express detectable levels of dnFADD were examined for their apoptotic response to Fas and BCR stimulation. The two non-expressor transfectants of WM cells were efficiently killed by Fas stimulation as expected (Fig. 2A). All four dnFADD-expressing clones exhibited strong resistance to Fas killing, consistent with previous observations in other systems that dnFADD blocked Fas-induced apoptosis (45,51). In contrast, both the expressor clones and non-express clones efficiently underwent apoptosis upon BCR stimulation. Examination of the cultures after 48 h revealed that the BCR-stimulated cultures contained predominantly dead cells and broken-down cell fragments (data not shown) and there were no noticeable differences in responses to BCR stimulation between the two groups of clones. As truncated FADD blocked Fas-induced apoptosis and not BCR-induced apoptosis, it appears that the latter process does not require FADD.
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The enhancing effect of Fas and the BCR on each other in apoptotic induction in WM cells is in contrast to mature B cells in which Fas-induced apoptosis is inhibited by BCR signaling and promoted by CD40 activated by Th cells (57). The effect of CD40 stimulation on Fas- and BCR-induced apoptosis in WM cells was also examined in the experiment described above (Fig. 2B, CD40 stimulation conditions used here were maximally protective against BCR-induced death; data not shown). CD40 stimulation alone had only minor effects on proliferation of these cells as measured by the number of viable cells. As expected, co-stimulation via CD40 substantially inhibited the negative effects of BCR signaling and restored the cell growth to ~90% of the control level. In contrast, CD40 did not affect the Fas response in either WM cells or the parental WEHI 231 (Fig. 2B
and data not shown). Further experiments (data not shown) demonstrated that CD40 treatment did not alter the dose response of WM cells to varying concentrations of anti-Fas and did not cause WEHI 231 to become Fas sensitive. Therefore in WM and WEHI 231, the Fas pathway is not responsive to CD40 signaling in the way it is in mature B cells.
Fas induces apoptosis through FADD, but Fas can also activate other signaling events such as JNK activation (25,26) To assess the contribution of FADD-mediated pathways to the observed synergistic death induction by Fas and the BCR, we examined the WM transfectants expressing dnFADD for their response to BCR stimulation in the presence or absence of co-stimulation via Fas. As shown in Fig. 2(C), without Fas co-stimulation, all three transfectants (two expressing dnFADD, Dn1 and Dn2, and one not, N7) underwent apoptosis similarly in response to varying concentrations of anti-IgM, all reaching maximum killing starting at 0.6 µg/ml of anti-IgM. In the transfectant not expressing dnFADD (N7), Fas co-stimulation caused a dramatic shift of the anti-IgM killing curve to low doses and the synergistic death induction by Fas and the BCR was readily seen at 0.2 µg/ml anti-IgM. In contrast, in both of the dnFADD-expressing clones tested (Dn1 and Dn2), Fas co-stimulation did not significantly affect the BCR killing curves. These results demonstrate that it is the FADD-mediated signaling pathways of Fas that confer a synergistic death-inducing effect with the BCR. FADD-independent signaling pathways induced by Fas, if any, did not appear to enhance BCR-induced apoptosis in these cells.
Apoptotic induction by the BCR and by Fas exhibit differential sensitivity to caspase inhibitors
Different apoptotic stimuli can induce cell death by activating different caspases. Fas and several other death receptors initiate cell death primarily by activating caspase-8 or -10, whereas growth factor withdrawal or DNA-damaging agents typically induce cell death through the mitochondrioncaspase-9 pathway (for review, see 52). To probe the involvement of different caspases in BCR- or Fas-induced apoptosis, we investigated the effects of the commercially available peptide inhibitors of caspases. WM cells were subjected to Fas or BCR stimulation in the presence or absence of the peptide caspase inhibitors and then examined for cell survival. In the case of Fas, all the inhibitors tested showed significant protective effects albeit to different degrees (Fig. 3A). The two pan-caspase inhibitors, BD and VAD, effectively protected cells from Fas killing, achieving >90% reversal of the Fas effects. BD displayed evident protective effects even at the concentration of 10 µM (data not shown). DEVD, an inhibitor with relative specificity for caspase-3 and -7, showed partial protection, and this is consistent with the current understanding that caspase-3 participates in Fas-induced apoptosis downstream of caspase-8 and -10 (53). LEHD, an inhibitor with relative specificity for caspase-4, -5 and -9, also provided limited protection. Control experiments indicated that the non-peptidyl FMK moiety of the inhibitors did not interfere with cell death induction by either the BCR or by Fas (data not shown).
The effects of these inhibitors on BCR-induced apoptosis were dramatically different from their effects on Fas-induced apoptosis. BD showed significant pro-survival effects and its presence abrogated the synergistic effects of co-stimulation via Fas and the BCR (Fig. 3A). As will be described further below, BD inhibited manifestations of apoptosis such as loss of plasma membrane integrity as judged by PI staining; however, cell cycle progression remained blocked. VAD, another pan-caspase inhibitor, showed a small but detectable pro-survival effect, which was further confirmed in repeated experiments. DEVD and LEHD, while partially protecting cells from Fas-induced apoptosis, did not exhibit significant effects on BCR-induced cell death. These results suggest that the BCR induces apoptosis through activation of caspases, but the DEVD- and LEHD-inhibitable caspases such as caspase-3, -4, -5, -7 and -9 may not play critical roles. The dramatically different effects of these caspase inhibitors on death induction by the BCR and Fas suggest that the two death processes are mediated by different subsets of caspases, which provides an explanation for the synergistic effect of Fas and the BCR in death induction as described in the previous section.
Several groups have reported the protective effect of VAD on BCR-induced apoptosis in different B cell lines (38,41,54,55); therefore, we compared the effect of BD and VAD in protecting WEHI 231 from BCR-induced apoptosis. Cells were stimulated via the BCR in the presence of BD or VAD (100 µM). The cultures were sampled 25 and 42 h later, and examined by PI-staining for cell viability and by Annexin V staining for exposure of phosphatidylserine on the cell surface, an early marker of apoptosis (Fig. 3B and C). At 25 h, the viable cells in the BCR-stimulated culture numbered 52% of those in the control culture (Fig. 3B
). The presence of VAD and BD improved the viable cell number to 63 and 73% when compared to their respective controls (i.e. BD- or VAD-containing cultures without BCR stimulation). The anti-apoptotic effects of VAD and BD were more evident at the 42 h time pointthe numbers of viable cells in BCR-stimulated cultures were 21% of its control in the absence of any caspase inhibitors, and 36 and 40% of their controls in the presence of VAD and BD respectively. Some of the drop in viable cell number in the presence of BD is due to BCR-induced growth arrest (see below). Further examination of the effects of BD and VAD on the appearance of phosphatidylserine on the cell surface revealed that, without the presence of BD or VAD, BCR stimulation caused elevated Annexin V staining in ~10% of the viable cells (i.e. PI-excluding cells) (Fig. 3C
) at both the 25 and 42 h time points. Both VAD and BD completely blocked the BCR-induced Annexin V staining of the surviving (PI-excluding) cells at both time points (Fig. 3C
). Indeed, VAD and BD can lower the background Annexin V staining in non-stimulated cells (Fig. 4C
and data not shown). Thus both BD and VAD are capable of inhibiting BCR- as well as Fas-induced apoptosis, with BD being more effective. Doseresponse experiments (data not shown) indicated that the concentration of 100 µM was maximally effective for both BD and VAD, in agreement with a previous report (41).
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BCR-induced cell cycle arrest is reversible and the killing process is mediated by BD-inhibitable factors
As the presence of BD or VAD allowed Fas-stimulated cells to proliferate similarly to the control cells (Fig. 3A), Fas signaling apparently did not perturb cell cycle progression in WM cells other than through caspase activation. In contrast, BCR-stimulated WEHI-231 cells are known to undergo growth arrest, followed by apoptotic death (33). Therefore we hypothesized that the pro-survival effect of BD on the BCR-stimulated cells, as shown in Fig. 3
(A and B), may reflect the inhibition of apoptosis by BD but not inhibition of growth arrest. In agreement with this idea, the number of apoptotic cells resulting from BCR engagement was decreased in the presence of BD (Fig. 4A
). The cell cycle status of the PI-excluding cells in the BCR-stimulated culture was examined by a 30-min BrdU pulse to allow cells in S phase to incorporate BrdU, followed by anti-BrdU staining to detect the cells in S phase and by PI staining for cellular DNA content. The BrdU incorporation assay revealed that BCR-stimulated cells arrested in G1/G0 phase even in the presence of BD (Fig. 4B
, II). Thus the increase in the number of viable cells seen in the BCR-stimulated culture in the presence of BD (Fig. 4A
, BCR + BD), compared to the BCR-stimulated culture without BD (Fig. 4A
, BCR), was primarily due to inhibition of apoptosis. The low number of viable cells in the BCR-stimulated culture with BD (BCR + BD), compared to the no-stimulation control (BD), was due to the lack of proliferation as well as to the incomplete inhibition of apoptosis.
To examine whether the WEHI 231 cells that were growth arrested by BCR stimulation in the presence of BD were capable of re-entering the cell cycle or whether they were terminally damaged without apoptotic manifestations, anti-IgM was removed from stimulated cells by washing and cells were re-cultured in fresh medium without BD. The cells were then examined for BrdU incorporation 0, 2, 6 and 18 h later. No change in cell cycle profile was detected after 2 or 6 h anti-IgM-free incubation (Fig. 4B, III and IV). However, after 18 h, a significant cell population (21%) was in S phase with a concomitant decrease in the G0/G1 population to 74% (Fig. 4B
, V). For comparison, cells that had not received BCR stimulation but were otherwise treated identically (Fig. 4B
, VI) had 52% of cells in S phase. Therefore a substantial fraction of the growth arrested cells were able to resume cell cycle progression upon removal of BCR stimulation. The BCR-stimulated cultures without BD had very few viable cells left at the end of the stimulation, making a BrdU incorporation assay difficult and its results hard to interpret. In a similar experiment in which cells were stimulated by anti-IgM and then anti-IgM was removed, the cells were further cultured for 52 h and assayed for the number of viable cells by PI staining and flow cytometry. More than twice as many live cells were recovered from the (BCR + BD) culture than from the BCR-alone culture. The PI-excluding cells were similar in size to normally proliferating cells as judged by forward light scatter (data not shown). BD treatment alone had no detectable effects on cell cycle progression. However, BD did show limited inhibition of cell proliferation over long-term (days) as judged by the number of cells in the BD-containing culture compared to untreated cells (~15% fewer cells were present after a 48 h culture period, data not shown). This small effect presumably reflects a non-specific effect of BD.
WM cells stimulated via Fas in the presence of BD had similar numbers of viable cells compared to the control cultures (Fig. 3A). The BrdU incorporation profile of these cells was similar to that of the control (Fig. 4D
), indicating that cell cycle progression was not perturbed by Fas treatment if apoptosis was blocked by the pan-caspase inhibitor BD.
Splenic immature B cells express heterogeneous levels of Fas but Fas-induced apoptosis was not detected in vitro
The expression pattern of WM cells, IgMhi, IgDlo/int and Fas+, is reminiscent of that of the peripheral immature (transitional) B cells, a population highly tolerizable through a BCR-dependent but poorly defined mechanism (17,18,21,22,47,57). As Fas dramatically enhanced BCR-induced apoptosis in WM cells (Fig. 2B and C), we investigated whether Fas might also enhance BCR-induced apoptosis in the splenic immature B cells. First we characterized the expression of Fas on these cells. Splenocytes from BALB/c mice (46 weeks old) were stained for Fas, HSA and CD19. The Fas staining was controlled by an isotype-matched antibody. Following flow cytometry, the immature B cells were identified as HSAhi/CD19hi/int and mature B cells as HSAint/CD19hi (Fig. 5A
). Both the immature and mature B populations showed low but detectable levels of Fas staining as indicated by the shift of peaks from the control staining. About 11% of the immature B cells (17% with anti-Fas 6% with control) were Fas+ (as indicated by the marked region, Fig 5A
), compared with 4% of the mature B cells (62%) being Fas+. Therefore, Fas+ cells were found in both immature and mature B populations, with the immature B population having a higher percentage of Fas+ cells. A similar conclusion was reached when IgM, instead of CD19, was used as the B cell marker to identify the immature and mature B cells (17,18,21,47) (data not shown). Two-color analysis using anti-IgM and anti-CD19 to stain splenocytes revealed a pattern of co-expression between IgM and CD19 as expected (99% IgM-expressing cells were CD19+ and 93% CD19-expressing cells were IgM+) (data not shown).
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Discussion |
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Activation-induced cell death of T cells is mediated by Fas, which requires FADD for killing (48,50). By analogy, it seems possible that BCR-induced apoptosis might also involve a Fas or TNFR-mediated process. However, when a dnFADD was expressed in WM cells we found that it blocked Fas- but not BCR-induced apoptosis, thus ruling out a necessary role of Fas/TNFR death domain pathways involving FADD in BCR-induced apoptosis, at least in WM cells. Consistent with our result, Lens et al. reported (43) that expression of a similar dnFADD in a derivative of the human B cell line Ramos also did not affect BCR-induced apoptosis. In addition, another group also reported that WEHI 231 cells do not require FADD for BCR-induced apoptosis (58). In agreement with this conclusion, we found that expression of v-FLIP (equine herpes virus 2/E8), a death effector domain protein that can inhibit Fas killing by blocking recruitment of pro-caspase-8 to FADD, also did not interfere with the apoptotic induction by the BCR (obtained from Dr J. Tschopp. University of Lausanne, Switzerland) (data not shown) Thus it appears that the Fas/TNFRFADDcaspase-8 pathways are not necessary for BCR-induced apoptosis in WM cells.
We found that Fas and the BCR synergized in inducing apoptosis in WM cells (Fig. 2B and C). This is in dramatic contrast to the interplay between BCR and Fas in mature B cells and mature B cell lines. Mature B cells interacting with Th cells during an immune response are rendered susceptible to Fas-mediated apoptosis via CD40CD40L interaction (13,59,60), whereas signals from the engaged BCR result in resistance to Fas killing (57). In contrast to mature B cells, WM cells did not increase Fas sensitivity upon CD40 stimulation and WEHI 231 cells did not respond to CD40 stimulation by acquiring Fas susceptibility. These findings reinforce the notion of WEHI 231 being immature B cells in nature. On the contrary, Ramos cells, a human B cell line also showing BCR-induced apoptosis, was markedly sensitized by CD40 for killing by Fas (43), thus resembling mature B cells. Similar to Ramos cells, a subclone of WEHI 231 was recently reported to respond to CD40 stimulation by acquiring susceptibility to Fas-induced apoptosis (58). It seems that the WEHI 231 cell line may be capable of giving rise to sublines with different phenotypes.
The broad spectrum caspase inhibitors, BD and VAD, provided maximal protection against Fas-induced death: cells stimulated via Fas in the presence of either of the two inhibitors were able to proliferate as well as the control cells did. This indicates that signals from ligated Fas probably only cause the activation of caspases and otherwise do not interfere with cell cycle progression. Two other caspase inhibitors, DEVD (relatively specific for caspase-3 and -7) and LEHD (relatively specific for caspase-4, -5 and -9) (61), were also partially protective. The DEVD result is consistent with the current understanding that caspase-8 activates caspase-3 which plays a major role in executing Fas-induced apoptosis (24). The LEHD result suggests that caspase-9 and mitochondria may be involved in Fas-induced apoptosis in WM cells (23).
For BCR-induced apoptosis, BD and VAD also provided the strongest protection but this was incomplete (Fig. 3A). Comparison of the two inhibitors indicated that BD was more effective than VAD (Fig. 3B and C
). VAD has been reported by other groups to inhibit BCR-induced death in different B cell lines (3841,54). Detailed study of the BD effects (Fig. 4A
) indicated that its protection against BCR-induced apoptosis resulted from inhibition of the apoptotic death of the stimulated cells, which in the presence of BD still underwent cell cycle arrest. A number of groups (3843) have observed caspase activation and changes in mitochondrial physiology during BCR-induced apoptosis in different cell lines. In WM cells, the DEVD and LEHD inhibitors did not show any inhibitory effects on BCR-induced apoptosis. These two inhibitors have strong specificity to caspase-3 and -9 (61), and were clearly inhibitory to Fas-induced apoptosis (Fig. 3A
). Our results are in contrast to reports implicating caspase-3 and -9 in BCR-induced apoptosis in two human B cell lines (42,43,62). Given the amplifying nature of caspase activation, it is conceivable that multiple caspases are activated during BCR-induced apoptosis. In fact, we also observed that caspase-3 was activated during BCR-induced apoptosis (data not shown). However, our inhibitor study suggests that caspase-3 activation does not play a critical role during BCR-induced apoptosis. Further studies are needed to ascertain which caspase species are indispensable in inducing or executing BCR-induced apoptosis. Additionally, the role of a particular caspase in mediating apoptosis may vary among different cell types. Caspase-9, for example, is known to play differential roles in Fas-induced apoptosis in different cell types (23,63).
Although the caspase inhibitor BD inhibited BCR-induced apoptosis of WM cells, the cells still exhibited cell cycle arrest (Fig. 4A and B). Moreover, a significant fraction of arrested cells resumed cell cycle progression when BCR stimulation was withdrawn. This result indicates that BD not only prevented various manifestations of cell death, such as loss of the plasma membrane permeability barrier and loss of phosphatidylserine asymmetry in the membrane, but also prevented other cell damage that would fatally wound cells and prevent them from progressing through the cell cycle. It should be pointed out that the BD inhibition of BCR-induced apoptotic death was not complete, as shown by the increased percentage of PI+ cells (and cell fragments) in BD-containing BCR-stimulated culture when compared with the no-stimulation control (Fig. 4A
). Also in the presence of BD, the number of cells surviving BCR stimulation decreased over time (data not shown). Nonetheless, the number of cells that resumed proliferation after removal of anti-IgM was significantly higher for cells treated with BD (Fig. 4B and C
), indicating that caspase inhibition by BD uncoupled the link between BCR-induced growth arrest and apoptotic death induction. Thus BCR-induced growth arrest and apoptotic death are separately controlled processes.
Compared with the parental WEHI 231, WM cells have similar levels of surface IgM, but up-regulated Fas expression and down-regulated IgD expression (IgMhi, IgDint/lo, Fas+). Immature or transitional B cells are also IgMhi, IgDlo/null and have been reported to express Fas (17,21,22,64,65). Developing immature B cells are tolerized in the bone marrow through the mechanisms of receptor editing, apoptosis and perhaps anergy. Yet most immature B cells that migrate to the spleen do not reach maturity and are generally presumed to be eliminated by BCR-mediated selection (17,57,64,66,67). We and others (17,18) have found that a substantial fraction of immature B cells (1015%) have acquired Fas expression. However, when splenocytes were stimulated in vitro via Fas we did not detect loss of immature B cells (Fig. 5B). A straightforward explanation is that the Fas antigen on the immature B cells does not transmit an apoptotic signal, as reported on human thymocytes (15). However, as mentioned above, cell culture conditions appeared to influence the sensitivity of WM cells to Fas-induced apoptosis. By analogy, the immature B cells could be sensitive to Fas-induced apoptosis in vivo. However, when they were cultured in vitro, their sensitivity to Fas may have been decreased or abolished due to the presence of survival factors or lack of death-promoting factors. Future experiments will be needed to address the mechanism by which Fas expression is induced on immature/transitional B cells and the functional role it may play in the further development of these cells.
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Acknowledgments |
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Abbreviations |
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BD BOC-Asp-FMK |
DEVD Z-Asp-Glu-Val-Asp-FMK |
JNK Jun N-terminal kinase |
LEHD Z-Leu-Glu-His-Asp-FMK |
PARP poly(ADP-ribose) polymerase |
PI propidium iodide |
TNF tumor necrosis factor |
TNFR tumor necrosis factor receptor |
VAD Z-Val-Ala-Asp-FMK |
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Notes |
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Received 14 August 2000, accepted 10 January 2001.
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References |
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