Apoptosis induced by the antigen receptor and Fas in a variant of the immature B cell line WEHI-231 and in splenic immature B cells

Maoxin Tim Tian, Chih-Hao Gilbert Chou and Anthony L. DeFranco

G. W. Hooper Foundation & Department of Microbiology and Immunology, University of California, San Francisco, CA 94143-0552, USA

Correspondence to: A. L. DeFranco


    Abstract
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Signaling by the BCR causes proliferation and resistance to Fas-induced apoptosis in mature B cells, but growth arrest and apoptosis in immature B cells. We have identified a variant of the immature B cell line WEHI 231 that retains the apoptotic response to the BCR but has acquired susceptibility to Fas-induced apoptosis. The Fas susceptibility was associated with increased Fas expression on the cell surface and down-regulated IgD expression. These cells exhibited a distinctive functional relationship in response to signals from the BCR, Fas and CD40: BCR stimulation markedly promoted Fas-mediated apoptosis (and vice versa) and Fas-induced apoptosis was not subject to modulation by CD40 signaling. While BCR-induced apoptosis was effectively rescued by CD40, it was not affected by the expression of a dominant-negative FADD. The mechanistic distinctions between BCR- and Fas-induced apoptosis were further characterized by the differential effects of different caspase inhibitors on these two processes which imply the involvement of different subsets of caspases. For BCR-induced apoptosis, we provide evidence that the final apoptotic destruction phase can be inhibited by the pan-caspase inhibitor BOC-Asp-FMK (BD) and that, in the presence of BD, the BCR only induces growth arrest which is reversible. The striking enhancing effects of Fas on BCR-induced apoptosis seen in the variant cells prompted us to examine if a similar cooperation in induction of apoptosis occurs in the highly tolerizable immature B cells of the spleen. We found that the splenic immature B population contains a significant number of Fas-expressing cells, but neither Fas-induced apoptosis nor an enhancing effect of Fas on BCR-induced apoptosis of these cells was detected in vitro.

Keywords: apoptosis, BCR, Fas, growth arrest, immature B lymphocytes


    Introduction
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
The immune system uses apoptosis of self-reactive lymphocytes as a critical molecular mechanism to establish self-tolerance (1,2). A high rate of de novo production of both T and B cells in the thymus and bone marrow creates an antigen receptor repertoire of great diversity, but subsequent selection processes for establishing self-tolerance permit only a small fraction of the generated lymphocytes to mature into the long-lived peripheral pool. A substantial portion of the developing lymphocytes bear self-reactive antigen receptors and the resulting persistent signaling from these BCR or TCR induces these cells to undergo apoptotic cell death (3). Receptor editing and anergy are additional means of avoiding self-specificity (4). Upon reaching maturity, lymphocytes respond to the stimulation of the BCR or TCR by proliferation and, for B cells, by acquiring resistance to Fas-mediated apoptosis (57). The mechanisms that govern the differential outcomes of the antigen receptor stimulation between immature and mature lymphocytes remain incompletely understood (8).

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-{kappa}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.


    Methods
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 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Culture media, antibodies and other reagents
RPMI 1640, DMEM and FSC were purchased from Gibco/BRL (Rockville, MD). Goat anti-mouse IgM (µ chain specific) was purchased from Jackson ImmunoResearch (West Grove, PA) and used at 10 µg/ml unless otherwise indicated. Hamster anti-mouse Fas monoclonal (Jo2), mouse anti-hamster IgG–FITC, rat anti-mouse IgM–biotin (R6-60.2), rat anti-IgD–FITC (11–26c.2a), rat anti-HSA–biotin (M1/69), hamster anti-Fas–FITC (Jo2) and its isotype-matched IgG (2, {lambda})–FITC; rat anti-HSA–FITC (M1/69) and rat anti-CD19–phycoerythrin (1D3) were all from PharMingen (San Diego, CA). Goat anti-IgM–phycoerythrin was from Southern Biotechnology Associates (Birmingham, AL). Streptavidin–TriColor was from Caltag (San Francisco, CA). Horseradish peroxidase-conjugated secondary antibodies for immunoblotting were from Amersham (Arlington Heights, IL). Rabbit anti-mouse Fas and rat anti-mouse Fas (7C10) were from Santa Cruz Biotechnology (Santa Cruz, CA) and Upstate Biotechnology (Lake Placid, NY) respectively. Mouse anti-poly(ADP-ribose) polymerase (PARP) antibody (C2.10) was from Enzyme Systems Products (Livermore, CA). Rabbit F(ab')2 anti-mouse IgM (µ chain-specific) was purchased from Zymed (South San Francisco, CA). Murine CD40 ligand (CD40L) was kindly provided as soluble recombinant gp39 produced by COS cells in DME medium by Dr D. Hollenbaugh (Bristol-Myers Squibb, Seattle, WA). The peptide inhibitors of caspases, including BD [(N-benzyloxycarbonyl-Asp-fluoromethylketone (BOC-Asp-FMK)], VAD [Z-Val-Ala-Asp-FMK (Z-VAD-FMK or Z-VAD)], DEVD [Z-Asp-Glu-Val-Asp-FMK (Z-DEVD-FMK)] and LEHD [Z-Leu-Glu-His-Asp-FMK(Z-LEHD-FMK)] were purchased from Enzyme Systems Products and dissolved in DMSO.

Plasmids and cDNAs
The cDNA of the dominant-negative-acting truncation mutant FADD/Mort1 (dnFADD) containing amino acids 81–205 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 373–1561, 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 1–5x105 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. 3Go, 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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Fig. 3. Differential effects of caspase inhibitors on Fas- and BCR-induced cell death. (A) Effects of different inhibitors. Identical WM cultures were seeded at 1x105 cells/ml in medium containing caspase inhibitors (100 µM) or solvent (DMSO) and stimulated with anti-Fas (1 µg/ml) or anti-IgM (10 µg/ml) or both for 38 h, stained with PI and assayed for viability by flow cytometry. Data are averages of duplicates with deviations within duplicates shown. *Values <0.5%. (B and C) Comparison of the effects of BD and VAD on BCR-induced apoptosis in WEHI 231. Cells were stimulated with 10 µg/ml anti-IgM in presence or absence of BD or VAD (100 µM). After 25 and 42 h, cells were analyzed for survival by PI exclusion (B) and for phosphatidylserine exposure on the surface of surviving (PI) cells by Annexin V staining (C). Data are averages of duplicates with deviations within duplicates shown. (D) BD and VAD inhibited the apoptotic breakdown of PARP. Cells were stimulated with anti-IgM (10 µg/ml) or anti-Fas (1 µg/ml) in medium containing caspase inhibitors (100 µM each) or solvent (DMSO) for 24 h, lysed and analyzed by anti-PARP immunoblotting (the 85 kDa protein represents a cleavage product of PARP whereas the full length is 116 kDa).

 
Cells were identically seeded at the start of an experiment at a density of 1x105 cells/ml in fresh medium with appropriate stimuli. After the indicated period of incubation, all cultures were pelleted, resuspended in equal volumes of PBS containing 1% BSA (or 2% FCS) and 1 µg/ml PI (Sigma, St Louis, MO), and analyzed by flow cytometry (FACScan; Becton Dickinson, San Jose, CA) for a fixed period (20 s). For limited Fas stimulation, cells were seeded at 2x105 cells/ml in DME medium containing 10% FCS. The apoptotic effects of stimulation with anti-Fas in WM cells were somewhat variable from experiment to experiment, presumably as a result of changes in culture conditions. High serum content in the medium and/or high seeding density of cells appeared to inhibit the Fas effects. The death induction by the BCR was also sensitive to the prior growth history and the culture conditions of the cells, but to a lesser degree than Fas. The relative effects of stimulations on apoptosis reported here represent consistent observations and representative experiments are shown. For CD40 stimulation experiments, CD40 ligand as soluble gp39 in DME was added to the regular RPMI 1640 medium in a ratio (1:40) that was determined in prior experiments with WEHI-231 cells (1x105 cells/ml) to produce saturating protection from BCR-induced apoptosis.

Annexin V staining for phosphatidylserine on the cell surface
To examine the externalization of phosphatidylserine as an early apoptotic marker, the Annexin V–FITC 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 SDS–PAGE 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 (5–10x105) were resuspended in 0.5 ml staining buffer (PBS/1% BSA/0.1% sodium azide) with a primary antibody (1–2 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 (5–10x105) 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-BrdU–FITC (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.


    Results
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 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
A subclone of WEHI-231 that is susceptible to apoptotic induction by Fas and by the BCR
During our study of BCR-induced apoptosis, we discovered unexpectedly a subclone of WEHI 231 in the lab (called WM) to be not only sensitive to apoptotic killing by the BCR but by Fas as well, while the parental WEHI 231 cell line is resistant to Fas-induced apoptosis (Fig. 1Go). When these cells were stimulated with 20 ng/ml anti-Fas antibody (Jo2), the number of viable cells decreased to <50% of the control culture within 25 h. The simplest hypothesis to explain the susceptibility of WM and the resistance of the parental WEHI-231 cells to Fas stimulation is that they differ in the expression of Fas on the cell surface. Therefore cells were stained with anti-Fas antibody and fluorescence-conjugated secondary antibody and analyzed by flow cytometry. As shown in Fig. 1Go(B and C), WEHI-231 cells had little or no surface Fas expression, but WM cells clearly expressed surface Fas. Thus the susceptibility to Fas apoptosis may be the result of the surface Fas expression on these cells. The reason for elevated surface Fas expression in these susceptible cells remains to be answered. Anti-Fas immunoblotting detected similar levels of Fas in the lysates of WM and WEHI-231 cells (data not shown), suggesting that post-translational processes may contribute to the surface expression of Fas on WM cells, as suggested previously in muscle cells (46).



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Fig. 1. Sensitivity of the WM cells to Fas-induced apoptosis and their surface Fas, IgD and IgM expression. (A) WM and WEHI 231 cells seeded at 105/ml in media with 1% FCS were stimulated by varying concentrations of anti-Fas antibody for 25 h, stained with PI and analyzed by flow cytometry. The number of viable cells (i.e. cells excluding PI) was determined and is presented as percentage of viable cells compared to untreated cultures. Shown is a representative of more than three experiments and data are averages of duplicates (variation within each duplicate was <15%). (B and C) cells were incubated with anti-Fas (Jo2) (filled peaks) or PBS (open peaks) and then stained with FITC-conjugated secondary antibody. Median fluorescence intensities, after subtracting the values for control staining, were 0.7 for WEHI 231 and 9.7 for WM cells. (D and E) WEHI 231 and WM cells were examined for IgD and IgM expression by staining with anti-IgD–FITC (D) or anti-IgM–biotin and streptavidin–TriColor (E), followed by flow cytometry.

 
WEHI 231 is generally regarded as an in vitro analog of immature B cells based on its negative response to BCR signaling, i.e. growth arrest and apoptosis (33,35). Since the response to Fas is a phenotype of activated mature B cells (11), we wanted to determine if the Fas expression and susceptibility of the WM cells may be indicative of changes of other developmental markers as well, such as IgM and IgD. WM cells and the parental WEHI 231 cells were stained with FITC-conjugated pan-IgD antibody and biotin-conjugated IgM, and analyzed by flow cytometry. WM and parental WEHI 231 showed similarly high levels of IgM expression, but the IgD expression on WM cells was significantly lower than on WEHI 231 cells (Fig. 1D and EGo). The cell surface expression profile of WM cells (IgMhi, IgDlo/int and Fas+) is reminiscent of that of the late immature (transitional) B cells (17,18,21,47).

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. 2AGo). 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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Fig. 2. Functional interactions among Fas, BCR and CD40 in modulating apoptosis. (A) FADD does not mediate BCR-induced apoptosis. Stable transfectant clones of WM expressing dnFADD (Dn1–Dn4) or not expressing it (N7 and N10) were stimulated via Fas [1 µg/ml anti-Fas (Jo2)] or the BCR (10 µg/ml anti-IgM) for 48 h and then analyzed by flow cytometry for viability by PI exclusion. The number of viable cells (PI) in stimulated cultures was compared with that of identically seeded control cultures and presented as a percentage. Differences among replicate cultures were <10%. (B) Fas and the BCR synergistically induce cell death and CD40 does not significantly affect Fas sensitivity. WM cells were seeded at 105 cells/ml in 10% FCS medium and stimulated via Fas (1 µg/ml anti-Fas) or the BCR (0.2 µg/ml anti-IgM) (both being submaximal stimulation conditions) or CD40 (2% v/v gp39-containing medium) or in combination for 47 h. Cultures were then stained with PI and assayed for viability by flow cytometry. Data are averages of duplicates with deviations within duplicates indicated by the error bars. *0.1%. (C) FADD-dependent pathway(s) mediates the promoting effect of Fas on BCR-induced apoptosis. A non-expressing transfectant and two dnFADD-expressing transfectants (see A) of WM were stimulated with varying concentrations of anti-IgM in the presence or absence of Fas co-stimulation (1 µg/ml) for 47 h. Cultures were assayed for viability by flow cytometry.

 
Fas and the BCR act synergistically to induce cell death
Fas induces cell death by activation of multiple caspases such as caspase-8 and -3, and the BCR also employs caspases to induce cell death (38,4143). Therefore we were interested in whether the apoptotic pathways of Fas and the BCR may converge at some point. We reasoned that if Fas and the BCR utilize overlapping pathways to induce cell death, then combining limited stimulation via Fas and the BCR would be additive in cell death induction. Limiting stimulation of Fas or the BCR alone resulted in reduction of viable cell number to ~50% of the control (Fig. 2BGo). This reduction was accompanied by an increase in the number of dead cells and dissembled cell fragments (PI+ and heterogeneous in size, data not shown). Around 25% viable cells would be expected in the doubly stimulated cultures if Fas and BCR were additive in death induction. Instead, simultaneous stimulation of Fas and BCR resulted in survival reduction to 0.1% of the control (Fig. 2BGo). Similarly, as shown in Fig. 2Go(C), the presence of limited Fas stimulation dramatically sensitized a WM subclone (N7, see Fig. 2AGo) to BCR-induced death: 0.2 µg/ml anti-IgM alone or anti-Fas alone reduced the number of surviving cells to 65 and 70% of the control respectively, but their co-stimulation resulted in the death of virtually all the cells. The synergistic death induction by Fas and the BCR implies that the overall apoptotic pathways by Fas and by the BCR are distinct.

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. 2BGo, 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. 2BGo 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. 2Go(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 mitochondrion–caspase-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. 3AGo). 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. 3AGo). 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 CGo). At 25 h, the viable cells in the BCR-stimulated culture numbered 52% of those in the control culture (Fig. 3BGo). 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 point—the 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. 3CGo ) 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. 3CGo). Indeed, VAD and BD can lower the background Annexin V staining in non-stimulated cells (Fig. 4CGo 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. Dose–response 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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Fig. 4. BCR-induced growth arrest is separable from apoptotic death and is reversible. (A) BD inhibited BCR-induced apoptotic breakdown. WEHI 231 cells were stimulated by anti-IgM (10 µg/ml) with or without BD (100 µM) for 47 h, stained with PI and examined by flow cytometry. Percentage of apoptotic cells and cell fragments (ungated regions) is noted. (B) BCR-induced growth arrest in the presence of BD was reversible. WEHI 231 cells in BD-containing medium were treated with anti-IgM (10 µg/ml) or not for 44 h and aliquots of the cultures were analyzed for BrdU incorporation (B, I and II) and PI staining (which showed 75% surviving cells in stimulated culture and 90% in control). Parallel cultures at this time were washed twice with PBS and resuspended in fresh medium without BD or anti-IgM and incubated for 2, 6 or 18 h as indicated. Cultures at different time points were examined for BrdU incorporation (100 µM, 30 min), stained with anti-BrdU–FITC antibody for DNA synthesis and PI for DNA content, and analyzed by flow cytometry. The upper, lower-left and lower-right regions represent cells in S, G0/G1 and G2/M phase of the cell cycle respectively. The numbers are the percentages of cells in gated regions. (C) In extended culture, more cells recovered from BCR killing in the presence of BD. WEHI 231 cells were treated with anti-IgM (10 µg/ml) with or without BD (100 µM) for 40 h. After removing anti-IgM by washing, cells were incubated (with or without BD) for an additional 52 h, stained with PI and examined by flow cytometry (timed counting). Relative number of recovered live cells (PI) during timed acquisition is shown (the experiment was in triplicate). (D) In contrast to BCR stimulation, Fas stimulation did not perturb cell cycling in the presence of BD. WM cultures with BD (100 µM) were stimulated via Fas or BCR for 46 h and then examined for cell cycle status by BrdU incorporation and PI staining.

 
PARP, a well-characterized caspase substrate which is often cleaved early during apoptosis (56), was chosen to evaluate biochemically the effects of the caspase inhibitor treatment (Fig. 3DGo). Cells incubated with caspase inhibitors (BD, VAD and DEVD) were stimulated with anti-IgM or anti-Fas and lysates were subjected to SDS–PAGE and immunoblotting with anti-PARP antibody. PARP cleavage was evident in both BCR- and Fas-induced apoptosis (the 85 kDa protein is one of the cleavage products of the intact 116 kDa protein). PARP cleavage was significantly inhibited by BD or VAD in both types of death induction, demonstrating that these inhibitors were effective in blocking the activity or activation of the caspases responsible for PARP cleavage. DEVD, which did not protect cells from BCR-induced apoptosis but partially protected from Fas apoptosis, did not inhibit PARP cleavage for reasons not yet clear.

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. 3AGo), 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. 3Go(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. 4AGo). 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. 4BGo, II). Thus the increase in the number of viable cells seen in the BCR-stimulated culture in the presence of BD (Fig. 4AGo, BCR + BD), compared to the BCR-stimulated culture without BD (Fig. 4AGo, 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. 4BGo, 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. 4BGo, V). For comparison, cells that had not received BCR stimulation but were otherwise treated identically (Fig. 4BGo, 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. 3AGo). The BrdU incorporation profile of these cells was similar to that of the control (Fig. 4DGo), 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 CGo), 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 (4–6 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. 5AGo). 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 5AGo), compared with 4% of the mature B cells (6–2%) 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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Fig. 5. Splenic immature B cells: expression of Fas on the cell surface (A) and responses to BCR- and/or Fas-induced apoptosis (B). (A) Splenocytes from BALB/c mice were stained with anti-HSA–biotin and streptavidin–TriColor, CD19–PE and anti-Fas–FITC or IgG–FITC (hamster isotype-matched control antibody). The lymphocyte population (based on SSC/FSC profile; not shown) was gated for CD19/HSA profile as shown above (A and B). The immature B cells were identified as CD19hi/int/HSAhi and mature B cells as CD19hi/HSAint. These two populations were further gated to display anti-Fas or control staining in histograms. Similar results were obtained when anti-IgM–PE instead of anti-CD19–PE was used in the three-color analysis on the same cells. (B) Fas stimulation did not affect the survival of splenic immature B cells nor did it promote BCR-induced apoptosis. Splenocytes were identically seeded (1x106/ml) and then stimulated with varying concentrations of anti-IgM [rabbit F(ab')2] with or without anti-Fas (1 µg/ml). Following a 18 h incubation, cells were stained with anti-HSA–FITC and anti-CD19–PE, and counted via flow cytometry (30 s). The number of immature and mature B cells acquired during the 30 s cytometry for each culture was determined, and is expressed as the percentage of the control culture. This is a representative of three similar experiments.

 
To examine whether Fas stimulation induces apoptosis in splenic immature B cells or whether Fas promotes BCR-induced apoptosis, splenocytes were stimulated with varying concentrations of anti-IgM with or without co-stimulation via Fas. After 18 h, cells were stained with anti-HSA and anti-CD19, and analyzed by flow cytometry using the mode of timed acquisition (30 s). The immature and mature B cells were identified using HSA and CD19 as described above (Fig. 5AGo), and their numbers determined and expressed as a percentage of the control culture. As shown in Fig. 5Go(B), BCR stimulation alone effectively reduced the number of immature B cells (HSAhi, CD19+) when compared to the control culture, presumably as a result of BCR-induced apoptosis. The mature B population was much less sensitive to the negative effects of BCR stimulation. However, Fas stimulation alone did not reduce the survival of either immature or mature B cells, even though it effectively killed WM cells (Fig 1Go). Co-stimulation via Fas did not cause a detectable change in the dose–response curve to anti-IgM. We concluded that a significant fraction (10–15%) of the splenic immature B cells expressed Fas, but under the conditions used, Fas-induced apoptosis was not detected in these cells.


    Discussion
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
WEHI 231 cells are normally resistant to Fas-induced cell death and have little Fas expression on the cell surface (Fig. 1AGo) (58). We show here that WM cells express significant levels of Fas on the cell surface and are susceptible to Fas-induced apoptosis. WM cells were initially examined as a subclone of WEHI 231 showing relatively high sensitivity to BCR-induced cell death and were found unexpectedly to be sensitive to Fas killing as well. It is not clear if the increase in Fas level in WM cells was entirely responsible for their sensitivity to Fas-induced apoptosis as Fas susceptibility may depend on other factors as well, such as the expression levels of FLIP, Bcl-2 and Btk (2729). The availability of this cell line allowed us to compare the mechanisms of BCR- and Fas-induced apoptosis, and to explore the functional relationship of the BCR and Fas.

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/TNFR–FADD–caspase-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 CGo). 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 CD40–CD40L 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. 3AGo). Comparison of the two inhibitors indicated that BD was more effective than VAD (Fig. 3B and CGo). 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. 4AGo) 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. 3AGo). 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 BGo). 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. 4AGo). 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 CGo), 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 (10–15%) have acquired Fas expression. However, when splenocytes were stimulated in vitro via Fas we did not detect loss of immature B cells (Fig. 5BGo). 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.


    Acknowledgments
 
We would like to thank Drs J. Zhang and A. Winoto (University of California, Berkeley, CA) for supplying the dnFADD cDNA and the anti-FADD serum, Dr D. Hollenbaugh (Bristol-Myers Squibb Pharmaceutical Research Institute, Seattle, WA) for the soluble gp39 (CD40L), and Dr J. Tschopp (University of Lausanne, Lausanne, Switzerland) for v-FLIP DNA. We are particularly indebted to Drs Cliff Lowell, Calvin Yu, Neetu Gupta and Malcolm Lowry for advice and critical review of the manuscript. This research project was supported by NIH grant AI20038. C.-H. G. C. was supported by an Arthritis Foundation Fellowship.


    Abbreviations
 
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

    Notes
 
Transmitting editor: S. J. Korsmeyer

Received 14 August 2000, accepted 10 January 2001.


    References
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 

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