In contrast to positive signaling leading to proliferation, the mechanisms involved in negative signaling culminating in apoptosis after B cell Ag receptor (BCR) ligation have received little study. We find that apoptosis induced by BCR cross-linking on EBV-negative mature and immature human B cell lines involves the following sequential, required events: a cyclosporin A-inhibitable, likely calcineurin-mediated step; and activation of caspase-2, -3, and -9. Caspase-2 is activated early and plays a major role in the apoptotic pathway, while caspase-9 is activated later in the apoptotic pathway and most likely functions to amplify the apoptotic signal. Caspase-8 and -1, which are activated by ligation of the CD95 and TNF-R1 death receptors, are not involved. Apoptosis induced by BCR ligation thus proceeds via a previously unreported intracellular signaling pathway.

Apoptosis, programmed cell death, or the cellular suicide program is a fundamental biological process that plays requisite roles in the development, differentiation, and maintenance of the cells of multicellular organisms. Inappropriate or dysregulated apoptosis, or failure to undergo programmed cell death has been implicated in a number of diseases and pathologic conditions (1). In the immune system, ligation of the Ag receptors on T and B lymphocytes (TCR (1), BCR3) may trigger survival signals leading to proliferation and differentiation or, alternatively, negative signals leading to apoptosis (2). Cellular selection between positive and negative signaling responses is determined by many factors, including cellular maturation state and tissue location, the nature and intensity of TCR or BCR ligation, and coligation of additional cell surface receptors in the appropriate temporal order (3, 4). The importance of apoptosis triggered by TCR ligation in shaping the T cell repertoire in the thymus and in homeostasis of mature peripheral T cells has been well documented (2, 3).

Apoptosis also regulates B cell maturation and differentiation, as well as the development of memory B cells (2, 4, 5). Immature B cells and cell lines representative of immature B cells undergo apoptosis after ligation of the BCR (6), a process that may contribute to the deletion of autoreactive immature B cells in vivo (7). Although mature B cells typically respond to Ag receptor ligation with proliferation, they can also be killed by apoptosis under certain circumstances, such as after extensive BCR cross-linking (8, 9). Death signals triggered by BCR ligation on immature and mature B cells and cell lines can be counteracted, and the cells rescued, by coligation of other cell membrane receptors, such as the IL-4R and CD40 (9, 10) or the CD19/CD21 complex (11, 12). These in vitro studies with cells and cell lines find their in vivo counterpart in the development of B cell responses to T-dependent Ags in germinal centers. B cells with high affinity for Ag presented on follicular dendritic cells in germinal centers differentiate into memory cells or plasma cells in the presence of costimulatory rescue signals (positive selection), whereas those with little or no affinity for Ag die by apoptosis (negative selection) (13, 14).

Considerable progress has been made in identifying the kinases, phosphatases, and signaling events that mediate proliferation after TCR or BCR ligation (15, 16). The same receptor-proximal signaling pathways are triggered in cells that die, rather than proliferate in response to Ag receptor cross-linking (3, 4). Apoptosis induced by TCR or BCR ligation is also undoubtedly additionally dependent on activation of caspase-type proteases, which form a death pathway able to mediate cell death by interfering with critical cellular functions and by disrupting cellular and genomic integrity (17). Although relatively little is known of caspase involvement in TCR- and BCR-induced apoptosis, activation of caspase-3 (Yama, apopain) has been found to precede apoptosis triggered by TCR ligation on murine thymocytes in vitro and in vivo in correlation with negative selection; and caspase-3 inhibitors have been reported to block such apoptosis (18, 19). Evidence for caspase-3 activation after BCR ligation has also been reported (20, 21, 22). The present studies, obtained with human B cell lines with mature and immature phenotypes, indicate that apoptosis induced by BCR ligation proceeds via a previously unreported caspase-2-, -3-, and -9-dependent pathway.

Rat-1 fibroblasts were obtained from Dr. J. Jackson (The Scripps Research Institute, La Jolla, CA). The B104 human B lymphoma cell line was kindly provided by Dr. M. Mayumi (Fukui Medical University, Fukui, Japan) (23). This EBV-negative cell line, derived from a child with malignant lymphoma, expresses CD10, CD19, CD20, CD21, CD35, and CD40, as well as Ia and surface IgM and IgD with κ light chains. The B104 cell line does not express CD23 or surface IgG, IgA, or IgE. The DND Burkitt lymphoma B cell line was also obtained from Dr. Mayumi (24). This EBV-negative cell line expresses CD11a, CD18, CD40, and CD54, as well as surface IgM and IgD; other characteristics have not been reported. The ST486 cell line was purchased from the American Type Culture Collection (ATCC, Manassas, VA). The cell line is EBV negative and expresses surface IgM and IgA with κ light chains, but lacks surface IgG and IgD (25); other characteristics have not been reported. A clonal ST486 cell line (ST486-M) was obtained by limiting dilution. The lymphoma cell lines were cultured in RPMI 1640 containing 10% FCS.

Flag-tagged crmA cDNA was created by PCR amplification from plasmid p8431 (kindly provided by Dr. D. Pickup, Duke University Medical Center, Durham, NC) by PCR using the following primers: 5′-CCG GAA TTC ACC ATG GAC TAC AAA GAC GAT GAC GAC AAG ATG GAT ATC TTC AGG GAA ATC-3′ and 5′-CCT GAA TTC TTA ATT AGT TGT TGG AGA GCA-3′. The amplified flag-tagged crmA fragment was digested with EcoRI and inserted into the EcoRI site of pcDNA3 (Invitrogen, San Diego, CA). The pHook-2 vector was also from Invitrogen. The pRSC-lacZ vector and a dominant-negative (DN) caspase-9 mutant (C287A) in pRSC-lacZ have been described (26). The wt (1) caspase-2 expression plasmid, pβactH372, was provided by Dr. J. Yuan (Harvard Medical School, Cambridge, MA) (27). To generate DN caspase-2, the active site cysteine was mutated to alanine (C303A, TGC→GCC) by site-directed mutagenesis and inserted into pRSV-lacZ and pcDNA3 (Invitrogen, Carlsbad, CA).

Transient expression of crmA in B104 cells was induced by electroporation (250 V, 960 μF). B104 cells (1 × 107 cells) were mixed with 10 μg of vector only, or 10 μg of pcDNA3/flag-crmA together with 1 μg of pHook-2 vector (10:1). Transfected cells expressing a single chain Ab on their membranes were isolated 72 h after transfection by their reactivity with hapten-coated magnetic beads (pHook2; Invitrogen). CrmA expression was determined by the Western blotting procedure using anti-flag M2 mAb (Eastman Kodak, New Haven, CT).

Rat-1 fibroblasts were transiently transfected with wt caspase-2 in the pβact H37Z vector, DN caspase-2 in the pRSV-lacZ vector, wt caspase-2 in the pβact H372 vector plus DN caspase-2 in either the pRSV-lacZ or pcDNA3 vectors, or DN caspase-9 in the pRSV-lacZ vector using lipofectamine (Life Technologies, Gaithersburg, MD). After 24 h, the cells were stained with 5-bromo-4-chloro-3-indolyl β-d-galactoside (X-Gal) (27).

B104 cells were transiently transfected with pRSV-lacZ/DN caspase-2, pRSV-lacZ/DN caspase-9, or both together in the pRSV-lacZ vector by electroporation (250V, 960 μF).

B104, ST486-M, and DND-39 cells were treated with 1 μg/ml purified DA4.4 mAb IgG to human IgM (ATCC) for the times specified in the various experiments. Cell viability was determined by the MTT assay system (Chemicon International, Temecula, CA) and occasionally by trypan blue exclusion. B104 cells transiently transfected with wt and DN caspases in the pRSV-lacZ plasmid were treated with anti-IgM 3 days after transfection. Ten hours later, the cells were stained with X-Gal. Viability of ≥200 blue cells was assessed in each experiment.

Apoptosis was also quantified by flow cytometry using annexin V and propidium iodide (PI) (1) to tag impermeable cells expressing membrane phosphatidylserine (PS) (1) (28). In these studies, DA4.4-treated B104 cells were washed in cold PBS, and 5 × 105 cells were resuspended in 100 μl of buffer (10 mM HEPES, pH 7.4, 140 mM NaCl, 2.5 mM CaCl2) and incubated with 5 μl of Annexin V-FITC (PharMingen, San Diego, CA) together with 10 μl of PI (50 mg/ml in PBS) for 15 min at room temperature in the dark. The samples were analyzed within 1 h by flow cytometry using a FACSort with CELLQuest software (Becton Dickinson, San Jose, CA).

After incubation with anti-IgM at 37°C for the times specified in Results, B cells were washed with cold PBS and lysed in 50 mM Tris-HCl (pH 7.4) containing 1 mM EDTA, 10 mM EGTA, and 10 μM digitonin (Sigma, St. Louis, MO). After 10 min at 37°C, the lysates were centrifuged at 13,000 × g for 20 min, and aliquots (0.5 ml) of the clear cytosolic supernatants were incubated with 2 μl of 50 μM zYVAD-AFC (7-amino-4-trifluoromethyl coumarin) or zDEVD-AFC (Enzyme Systems Products, Dublin, CA), and AFC release was assessed spectrophotometrically (SLM800 spectrophotometer; SLM Instruments, Urbana, IL; excitation wavelength, 400 nm; emission wavelength, 505 nm). Cleavage of Ac-DEVD-pNA (para-nitroanilide) and Ac-VDVAD-pNA (Enzyme System Products) was also evaluated. In these studies, cytosols of B104 cells that had been incubated with anti-IgM for the times specified in the text were prepared by resuspending the cells (1 × 107/ml) in HEPES buffer (20 mM HEPES-KOH, pH 7.5, 10 mM KCl, 1.5 mM MgCl2, 1 mM EDTA, 1 mM sodium EGTA, 5 mM DTT, 0.1 mM PMSF, 1 μg/ml leupeptin, and 1 μg/ml aprotinin) and incubated, followed by repetitive passage (15 times) through a 26-gauge needle. After centrifugation (16,000 × g for 30 min), 100-μl aliquots were dispensed into 96-well plates and incubated with 40 μM Ac-DEVD-pNA or Ac-VDVAD-pNA. Release of pNA assessed at 405 nm in a Spectra Max 250 plate reader (Molecular Devices, Sunnyvale, CA); relative activities were obtained by dividing the protease activity observed at each time point by the time zero values.

Caspase activation was also assessed by the Western blotting procedure. In these studies, B104 cells (5 × 105/sample), which had been incubated with anti-IgM for the times specified in the text, were subjected to SDS-PAGE analysis, and Western blots were evaluated for reactivity with mAb to caspase-1, -2 (PharMingen), or -9 (a kind gift from Dr. Xiaodong Wang, Southwestern Medical Center, Dallas, TX), or with polyclonal Abs to caspase-3 or -8 (PharMingen). Reactivity was visualized via the ECL system (Amersham Life Sciences, Arlington Heights, IL), after the addition of HRP-labeled second Ab (Kirkegaard and Perry Laboratories, Gaithersburg, MD). Double-sized samples (106 cells) and double-thickness (1.5-mm) gels were necessary to detect caspase-9 in B104 cells. Cleavage of PARP (1) was also evaluated in Western blotting studies using mAb C-2-10 (from Dr. G. Poirier, Quebec, Canada). Blots were routinely stripped (100 mM 2-ME, 2% SDS, 62.5 mM Tris-HCl, pH 6.7) at 70°C for 30 min, and reprobed with mouse anti-actin mAb clone C4 (ICN Biochemicals, Aurora, OH) to verify equal loading.

Peptide inhibition studies were of two types. In the first, B cells (2 × 106/ml) were incubated for 2 h with zVAD-fmk, zYVAD-fmk, or zDEVD-fmk (Enzyme Systems Products), or zVDVAD-fmk (Calbiochem, San Diego, CA) at the concentrations indicated in the text before the addition of anti-IgM. A range of concentrations was employed in many studies. The single 100 μM concentration used in some experiments was chosen on the basis of dose-response experiments (10–500 μM) as the concentration that produced maximal effects. Using activity measurements, the Western blotting procedure, or apoptosis, caspase activation was assessed. In the second approach, cytosols prepared 6 h after treatment of B cells with anti-IgM were incubated for 10 min with 1 nM concentration (∼4 times the IC50 concentration) with the irreversible active site inhibitors before assessing caspase-enzymatic activities with synthetic peptide substrates.

In the CsA inhibition studies, B104 cells were incubated with the indicated concentrations of CsA for 2 h before BCR cross-linking. Caspase activation and apoptosis were assessed 16 h later. For the crmA inhibition studies, B104 cells were transiently transfected with crmA and evaluated for crmA expression in Western blotting studies with anti-flag M2 mAb 72 h after transfection. Cells expressing crmA were evaluated for caspase cleavage or apoptosis 16 h after BCR cross-linking.

Cross-linking of mIgM on B104 cells, a human B lymphoma cell line with a mature phenotype, induced rapid cell death, as previously reported (23). Cell death (MTT assay) was detectable at 4 h, and 50% cell loss occurred ∼10 h after BCR ligation (Fig. 1,A). BCR cross-linking on ST486-M cells, a clonal human B lymphoma cell line with an immature phenotype, induced cell death with slower kinetics (Fig. 1 A), as did BCR ligation on DND-39 cells, a human B lymphoma cell line with a mature phenotype (data not shown). BCR-ligated B104 cells were enlarged with electrolucent cytoplasm and eccentric nuclei containing condensed chromatin; the cells did not show DNA fragmentation as earlier reported (29). Dying ST486-M and DND-39 cells, in contrast, exhibited the classic morphologic features of apoptosis, as well as DNA fragmentation.

FIGURE 1.

Characteristics of cell death induced by BCR ligation. A, Kinetics of cell death. Cell death after BCR ligation on B104 and ST486-M cells was evaluated by the MTT assay at the times shown. The error bars represent SDs. B, Effect of caspase inhibitors on cell death. B104 and ST486-M cells were preincubated for 2 h with buffer, zYVAD-fmk, zVAD-fmk, or zDEVD-fmk (100 μM) before BCR ligation. Cell death was evaluated by the MTT assay 18 h (B104 cells) or 36 h (ST486-M cells) later. Bars denote SDs. C, PS exposure on the cell surface after BCR cross-linking on B104 cells. PS exposure was assessed at various times after BCR ligation by flow cytometry on cells stained with FITC-labeled annexin V and counterstained with PI. The proportion of cells in each of the four sectors is indicated.

FIGURE 1.

Characteristics of cell death induced by BCR ligation. A, Kinetics of cell death. Cell death after BCR ligation on B104 and ST486-M cells was evaluated by the MTT assay at the times shown. The error bars represent SDs. B, Effect of caspase inhibitors on cell death. B104 and ST486-M cells were preincubated for 2 h with buffer, zYVAD-fmk, zVAD-fmk, or zDEVD-fmk (100 μM) before BCR ligation. Cell death was evaluated by the MTT assay 18 h (B104 cells) or 36 h (ST486-M cells) later. Bars denote SDs. C, PS exposure on the cell surface after BCR cross-linking on B104 cells. PS exposure was assessed at various times after BCR ligation by flow cytometry on cells stained with FITC-labeled annexin V and counterstained with PI. The proportion of cells in each of the four sectors is indicated.

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Cell death triggered by BCR ligation on these three human B lymphoma cells is clearly apoptotic because it is not passive but, rather, is triggered by ligation of a cell-activating membrane receptor and involves requisite caspase activation, because it was largely inhibited by preincubating the cells with certain irreversible active site caspase inhibitors (Fig. 1,B). Furthermore, BCR ligation led to time-dependent exposure of PS on the membrane of intact cells (Fig. 1 C), an accepted measure of early apoptotic cellular changes (28). The evolution from annexin V- and PI-negative cells, to annexin V-positive and PI-negative cells and, finally, to annexin V- and PI-positive cells is evident.

Programmed cell death induced by many agents is associated with caspase-3 activation, and subsequent cleavage of various substrates by the activated enzyme (30, 31). Evidence for caspase-3 cleavage or activation 4–24 h after BCR ligation on various human B cell lines has been obtained (20, 21, 22). In the present study, initial cleavage of caspase-3 into 20- and 17-kDa fragments was evident 2 h after BCR cross-linking on B104 cells, and maximal cleavage occurred 4–8 h after BCR ligation (Fig. 2,A). The observed cleavage represents activation, because B104 cell extracts acquired the ability to cleave zDEVD-AFC, a synthetic substrate of caspase-3-like enzymes (data not shown), as well as a natural protein substrate, PARP, a DNA repair enzyme (Fig. 2,B), with the same kinetics. Caspase-3 was also cleaved after BCR ligation on ST486-M cells; significant cleavage was apparent 8 h after BCR cross-linking, and maximal cleavage occurred 12–24 h after BCR ligation (data not shown). Caspase-3 activation was blocked by preincubating B104 cells with zDEVD-fmk, an irreversible caspase-3 inhibitor, before BCR ligation (Figs. 2 C), as was PARP cleavage (data not shown).

FIGURE 2.

Activation of caspase-3 after BCR ligation on B104 cells. A, Evaluation of caspase-3 cleavage. Samples taken at intervals after BCR ligation were examined for caspase-3 cleavage with a polyclonal Ab to caspase-3 in Western blotting studies. B, Cleavage of PARP after BCR ligation. The Western blotting analyses employed a mAb to PARP. C, Effect of caspase inhibitors on caspase-3 cleavage. Caspase-3 cleavage was assessed 16 h after BCR ligation. B104 cells that had been preincubated with zYVAD-fmk, zVAD-fmk, or zDEVD-fmk (100 μM) for 2 h. D, Effect of crmA on caspase-3 cleavage. Caspase-3 cleavage was assessed 16 h after BCR ligation on B104 cells transiently transfected with crmA. The results of three independent experiments are shown.

FIGURE 2.

Activation of caspase-3 after BCR ligation on B104 cells. A, Evaluation of caspase-3 cleavage. Samples taken at intervals after BCR ligation were examined for caspase-3 cleavage with a polyclonal Ab to caspase-3 in Western blotting studies. B, Cleavage of PARP after BCR ligation. The Western blotting analyses employed a mAb to PARP. C, Effect of caspase inhibitors on caspase-3 cleavage. Caspase-3 cleavage was assessed 16 h after BCR ligation. B104 cells that had been preincubated with zYVAD-fmk, zVAD-fmk, or zDEVD-fmk (100 μM) for 2 h. D, Effect of crmA on caspase-3 cleavage. Caspase-3 cleavage was assessed 16 h after BCR ligation on B104 cells transiently transfected with crmA. The results of three independent experiments are shown.

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Because the caspase-3 inhibitor, zDEVD-fmk, also potently inhibited cell death induced by BCR ligation on B104 and ST486-M cells (Fig. 1 B), these findings strongly suggest that caspase-3, or a caspase-3-like enzyme, is a required participant in the BCR signaling pathway leading to the death of mature and immature human B cell lines. Of note, these results contrast with two recent reports in which Ac-DEVD-CHO failed to inhibit BCR-induced apoptosis of cloned Ramos B cells (21) and WEHI-231 immature murine B cells (32). The reasons for the different results are not known.

BCR-induced activation of caspase-3 and apoptosis was inhibited by preincubating B104 and ST486-M cells with zVAD-fmk, an irreversible inhibitor of numerous caspases, before the addition of anti-IgM (Figs. 1,B and 2C). These findings provided suggestive evidence for the involvement of another caspase in caspase-3 activation after BCR ligation, because zVAD-fmk is a relatively weak caspase-3 inhibitor (33). Definitive evidence for involvement of an additional caspase upstream of caspase-3 came from the demonstration that the cowpox-encoded caspase inhibitor, crmA, blocked BCR-induced apoptosis after expression in B104 cells. CrmA, although a good caspase-8 (Ki = 950 pM) and caspase-1 (Ki = 10 pM) inhibitor, is not an effective caspase-3 inhibitor (Ki = 500 nM) (34). In these studies, B104 cells were transiently cotransfected with crmA and pHook-2 expression plasmids. In three independent experiments in which high levels of crmA expression were obtained, caspase-3 cleavage (Fig. 2 D) and PARP cleavage and apoptosis (data not shown) were all markedly inhibited.

Caspase-8 and -1 are known activators of caspase-3 (30). Although we considered it unlikely that caspase-8 was involved in BCR-triggered apoptosis of B104 cells because of the absence of death domains (1) in either membrane IgM or the associated CD79a and CD79b proteins, the observed inhibition of caspase-3 activation and apoptosis by crmA, a good caspase-8 and caspase-1 inhibitor (34), suggested the possibility that BCR components interacted directly, or via an intermediate protein(s) with a death effector domain (1)-containing adaptor protein(s), and thereby recruited caspase-8 or caspase-1. However, Western blotting analyses provided no evidence for cleavage of caspase-8 or caspase-1 during the first 8 h after BCR ligation on B104 cells (data not shown). Thus, neither caspase-8 nor caspase-1 was directly activated after BCR ligation. A minor amount of caspase-8 cleavage was detected 12 h after BCR ligation, but this most likely represents secondary or feedback activation, rather than direct activation. Studies using other approaches have also shown that upstream events in the BCR and death receptor apoptotic pathways differ (21, 22).

Because caspase-3 is activated in vitro in the presence of caspase-9, cytochrome c, Apaf-1, and dATP (26, 35), we evaluated the possibility that BCR ligation on B104 cells led to caspase-9 activation. Although caspase-9 is present in very low concentrations in B104 cells, it was not cleaved during the first 8 h after BCR ligation on B104 cells, although modest cleavage became apparent 12 h after BCR ligation (Fig. 3,A). This late cleavage was most likely mediated by caspase-3, or by a caspase-3-like enzyme via a feedback mechanism, because it was blocked by low concentrations of the irreversible caspase-3 inhibitor, zDEVD-fmk (Fig. 3 B). In this regard, caspase-3 has been shown to directly activate caspase-9 in vitro (36).

FIGURE 3.

Caspase-9 plays a minor role in BCR-induced apoptosis of B104 cells. A, Evaluation of caspase-9 cleavage after BCR ligation. Samples were taken at intervals after BCR ligation and evaluated for caspase-9 cleavage by immunoblotting with a polyclonal Ab to caspase-9. Double-sized samples and double-thickness gels were used. B, The caspase-3 inhibitor, zDEVD-fmk, blocks late activation of caspase-9 after BCR ligation. Cells were incubated with various concentrations of zDEVD-fmk for 2 h before BCR ligation, and evaluated for caspase-9 cleavage 16 h later.

FIGURE 3.

Caspase-9 plays a minor role in BCR-induced apoptosis of B104 cells. A, Evaluation of caspase-9 cleavage after BCR ligation. Samples were taken at intervals after BCR ligation and evaluated for caspase-9 cleavage by immunoblotting with a polyclonal Ab to caspase-9. Double-sized samples and double-thickness gels were used. B, The caspase-3 inhibitor, zDEVD-fmk, blocks late activation of caspase-9 after BCR ligation. Cells were incubated with various concentrations of zDEVD-fmk for 2 h before BCR ligation, and evaluated for caspase-9 cleavage 16 h later.

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Transfection studies with DN caspase-9 (26) were conducted to definitively assess the role of caspase-9 in apoptosis of B104 cells triggered by BCR ligation. Apoptosis induced by BCR ligation was evaluated in B104 cells transiently transfected with either DN caspase-9 or lacZ containing vector. DN caspase-9 only modestly inhibited apoptosis of blue cells induced by BCR cross-linking in this experiment (27 ± 6% viability for DN transfected vs 6 ± 5% viability for vector-transfected cells); these data are representative of five identical studies.

Because of the lack of a major role for caspase-9 in caspase-3 activation after BCR ligation, we hypothesized the involvement of another CARD (1)-containing caspase; such caspases include human caspases-1, -2, -4, -5, -8, -9, -10, and -13 (37, 38). Of these, significant roles for caspase-1 and -8 were eliminated by the experiments presented above; and caspase-9 was found to play a minor role. Caspase-10 was considered an unlikely candidate, because it is a receptor-type caspase with death effector domains (39). We focused on caspase-2, because it is closely related to caspase-9 (30) and contains a functional CARD domain (40, 41). In addition, caspase-2 is highly expressed in lymphocytes and participates in certain B and T cell apoptotic reactions (42, 43).

Caspase-2 was cleaved after BCR ligation on B104 cells, with peak generation of the intermediate 33-kDa cleavage product 4–6 h after cross-linking (Fig. 4 A); the Ab used is directed against the prodomain of the molecule, and does not recognize the typical p18 and p14 cleavage products (43). Identical results were obtained with ST486-M cells, except that cleavage occurred several hours later (data not shown). Caspase-2 cleavage most likely corresponded to activation, because cytosols of anti-IgM-treated B104 cells acquired the ability to hydrolyze Ac-VDVAD-pNA, a caspase-2 substrate (44); peak cleavage occurred 4–6 h after BCR ligation (data not shown). Although in vitro studies have shown that caspase-3 can also cleave this substrate (44), the Ac-VDVAD-pNA cleaving activity present in B cell cytosols 6 h after BCR cross-linking was not mediated by caspase-3, because Ac-VDVAD hydrolysis was unaffected by the addition of 1 nM zDEVD-fmk, an irreversible caspase-3 inhibitor, to the cytosols, whereas the addition of 1 nM zVDVAD-fmk blocked cleavage (data not shown). Because the caspase specificity of VDVAD has not been analyzed, it is possible that other caspases cleave and are inhibited by this peptide. However, the similar kinetics of caspase-2 cleavage after BCR ligation with those obtained for the ability of cytosols of BCR-ligated cells to cleave VDVAD, strongly suggest that caspase-2 is activated early after BCR ligation.

FIGURE 4.

Caspase-2 involvement in the BCR-triggered apoptotic pathway. A, Evaluation of caspase-2 cleavage after BCR ligation on B104 cells. Samples were taken at intervals after BCR ligation and evaluated for caspase-2 cleavage by immunoblotting with a mAb to caspase-2. B, Effect of zVDVAD on apoptosis of B104 and ST486-M cells. Cell death was evaluated by the MTT assay 18 h (B104 cells) or 36 h (ST486-M cells) after cross-linking of the BCR on B104 cells preincubated with buffer or various concentrations of zVDVAD-fmk. Bars denote SDs. C, Evaluation of the effects of mutant caspase-2 (C303A) on apoptosis of Rat-1 cells induced by wt caspase-2. Rat-1 cells transiently transfected with wt caspase-2, caspase-2 (C303A), wt caspase-2 plus caspase-2 (C303A), wt caspase-2 plus DN caspase-9, or vector were evaluated for cell death 24 h later. Viability of ≥200 blue cells was evaluated. Bars denote SDs. D, Evaluation of the effect of DN caspase-2 and -9 on BCR-induced apoptosis of B104 cells. B104 cells were transiently transfected with DN caspase-2 (C303A), DN caspase-9, DN caspase-2 plus DN caspase-9, or vector. Viability of ≥200 blue cells was assessed 10 h after BCR cross-linking. Bars denote SDs. E, Effect of irreversible caspase-2- and -3-like inhibitors on caspase-2 cleavage after BCR ligation. Cell lysates were evaluated for caspase-2 cleavage 16 h after BCR ligation on B104 cells that had been preincubated with zVDVAD-fmk or z-DEVD-fmk. F, Effect of the irreversible caspase-2 inhibitor, zVDVAD-fmk, on caspase-3 cleavage after BCR ligation. Cell lysates were evaluated for caspase-3 cleavage 16 h after ligation of the BCR on B104 cells that had been preincubated with various zVDVAD-fmk concentrations.

FIGURE 4.

Caspase-2 involvement in the BCR-triggered apoptotic pathway. A, Evaluation of caspase-2 cleavage after BCR ligation on B104 cells. Samples were taken at intervals after BCR ligation and evaluated for caspase-2 cleavage by immunoblotting with a mAb to caspase-2. B, Effect of zVDVAD on apoptosis of B104 and ST486-M cells. Cell death was evaluated by the MTT assay 18 h (B104 cells) or 36 h (ST486-M cells) after cross-linking of the BCR on B104 cells preincubated with buffer or various concentrations of zVDVAD-fmk. Bars denote SDs. C, Evaluation of the effects of mutant caspase-2 (C303A) on apoptosis of Rat-1 cells induced by wt caspase-2. Rat-1 cells transiently transfected with wt caspase-2, caspase-2 (C303A), wt caspase-2 plus caspase-2 (C303A), wt caspase-2 plus DN caspase-9, or vector were evaluated for cell death 24 h later. Viability of ≥200 blue cells was evaluated. Bars denote SDs. D, Evaluation of the effect of DN caspase-2 and -9 on BCR-induced apoptosis of B104 cells. B104 cells were transiently transfected with DN caspase-2 (C303A), DN caspase-9, DN caspase-2 plus DN caspase-9, or vector. Viability of ≥200 blue cells was assessed 10 h after BCR cross-linking. Bars denote SDs. E, Effect of irreversible caspase-2- and -3-like inhibitors on caspase-2 cleavage after BCR ligation. Cell lysates were evaluated for caspase-2 cleavage 16 h after BCR ligation on B104 cells that had been preincubated with zVDVAD-fmk or z-DEVD-fmk. F, Effect of the irreversible caspase-2 inhibitor, zVDVAD-fmk, on caspase-3 cleavage after BCR ligation. Cell lysates were evaluated for caspase-3 cleavage 16 h after ligation of the BCR on B104 cells that had been preincubated with various zVDVAD-fmk concentrations.

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To assess the importance of a caspase-2-like enzyme for BCR-induced apoptosis, B104 and ST486-M cells were preincubated with various concentrations of zVDVAD-fmk for 2 h before BCR ligation, and cell death was assessed 18 h (B104 cells) or 36 h (ST486-M) later. Apoptosis of both cell types was inhibited by zVDVAD-fmk in a dose-dependent manner (Fig. 4 B).

Transient transfection studies provided definitive evidence for caspase-2 involvement. These experiments were conducted with wt caspase-2 and a caspase-2 construct in which the cysteine at position 303 in the active site (QACRG) had been mutated to alanine (C303A). To demonstrate that mutant caspase-2 (C303A) functioned in a DN manner, Rat-1 fibroblasts were transiently transfected with wt caspase-2, caspase-2 (C303A), or both constructs together in lacZ-containing vectors. Apoptosis of blue cells was induced by wt caspase-2 (90 ± 4% cell death), but not by mutant caspase-2 (6 ± 6% death), and mutant caspase-2 markedly inhibited apoptosis induced by wt caspase-2 (35 ± 7% death) (Fig. 4 C), documenting the DN character of the mutant.

Next, studies were performed in B cells to evaluate the role of caspase-2 in BCR-induced apoptosis. In these experiments, B104 cells were transiently transfected with DN caspase-2 in the lacZ-containing plasmid. Three days after transfection, the cells were treated with anti-IgM, and viability of blue cells was evaluated 10 h later. DN caspase-2 markedly inhibited cell death induced by BCR cross-linking (67 ± 4% viability vs 7 ± 2% for the vector control) (Fig. 4,D). These findings document a major essential role for caspase-2 in BCR-induced apoptosis. In the same experiments, B104 cells were also transfected with DN caspase-9 alone and together with DN caspase-2. DN caspase-9 only modestly inhibited apoptosis induced by BCR ligation (22 ± 6% viability), similar to the values reported above. However, BCR-induced apoptosis was almost completely blocked (80 ± 5% viability) in B104 cells cotransfected with DN caspase-2 and -9 together (Fig. 4 D). These findings, which represent the combined results of three identical experiments, indicate that caspase-2 and -9 are both important in BCR-induced apoptosis, with caspase-2 playing the major role. Blocking both caspases essentially abrogated apoptosis triggered by BCR ligation.

To determine the sequence of involvement of caspase-2 and -3 in BCR-mediated apoptosis, B104 cells were preincubated with several concentrations of zVDVAD-fmk or zDEVD-fmk before BCR cross-linking. The caspase-2-like inhibitor, zVDVAD-fmk, but not the caspase-3 inhibitor, zDEVD-fmk, inhibited caspase-2 cleavage in BCR-ligated B104 cells in a dose-dependent manner (Fig. 4,E). The zVDVAD-fmk peptide also inhibited the generation of the 17- and 20-kDa fragments characteristic of activated caspase-3 (Fig. 4,F), as did zDEVD-fmk (Fig. 2,C) in BCR cross-linked B104 cells. Although a 22–23-kDa caspase-3 cleavage product was formed in the presence of low concentrations of zVDVAD-fmk (Fig. 4 F), this cleavage product lacked caspase-3 activity, as lysates of cells pretreated with zVDVAD-fmk before BCR ligation were unable to cleave either zDEVD-pNA or PARP (data not shown). Presumably, the inactive 22–23-kDa cleavage product is generated by a non-zVDVAD-fmk-inhibitable caspase. These studies indicate that caspase-2 activation precedes and is required for caspase-3 activation after BCR cross-linking.

In the transient transfection studies described earlier, Rat-1 cells were also transfected with wt caspase-2 plus DN caspase-9. Caspase-2-induced apoptosis (90 ± 4% cell death) was blocked in the presence of DN caspase-9 (18 ± 5% death) (Fig. 4 C), suggesting that caspase-9 acts downstream of caspase-2.

CsA (100 ng/ml) completely blocked apoptosis of B104 and ST486-M cells induced by BCR ligation (data not shown), findings that are consistent with previous findings (29, 45). In dose-response studies, very low concentrations of CsA inhibited caspase-2 cleavage (Fig. 5) as well as caspase-3 cleavage (data not shown) after BCR ligation on B104 cells. PARP cleavage was also inhibited (data not shown). Thus, caspase-2 and -3 activation and apoptosis induced by BCR ligation are dependent on an upstream CsA-inhibitable step.

FIGURE 5.

Caspase-2 activation and apoptosis after BCR ligation are dependent on a CsA-inhibitable step. Effect of CsA on caspase-2 cleavage after BCR ligation. Caspase-2 cleavage was assessed 16 h after BCR ligation on B104 cells that had been preincubated with CsA.

FIGURE 5.

Caspase-2 activation and apoptosis after BCR ligation are dependent on a CsA-inhibitable step. Effect of CsA on caspase-2 cleavage after BCR ligation. Caspase-2 cleavage was assessed 16 h after BCR ligation on B104 cells that had been preincubated with CsA.

Close modal

These studies show that BCR ligation triggers a previously undescribed intracellular signaling cascade involving a CsA-inhibitable step and sequential activation of caspase-2, -3, and -9 (Fig. 6). Caspase-8 and -1, upstream caspases activated by ligation of the CD95 and the TNF-R1 death receptors, were not activated by BCR ligation. Although most of the present studies were performed with B104 cells, the results of key experiments conducted with ST486-M cells indicate that the signaling pathway mediating apoptosis after BCR ligation is not unique to the B104 cell line. Because B104 cells exhibit a mature phenotype and ST486-M cells an immature phenotype, the apoptotic pathway is not dependent on the maturation state of the B cell. The signaling pathway also does not correlate with the presence of apoptotic phenotypic characteristics, because ST486-M cells exhibit such features, whereas B104 cells do not.

FIGURE 6.

Key events in the apoptotic pathway triggered by ligation of the BCR on B104 cells.

FIGURE 6.

Key events in the apoptotic pathway triggered by ligation of the BCR on B104 cells.

Close modal

Clear-cut activation of caspase-2 and -3 was evident 2 h after BCR cross-linking on B104 cells, and peak activation occurred 4–8 h after BCR ligation. This time frame correlated with the onset of rapid cell death. Qualitatively identical results were obtained with ST486-M cells, except that caspase-2 and -3 activation and cell death triggered by BCR ligation occurred several hourslater than in B104 cells. Experiments with cell-permeable fmk derivatives of caspase peptide substrates, which are irreversible active site inhibitors, showed that caspase-2 and -3 were required for apoptosis. Studies with B104 cells expressing DN caspase-2 provided definitive evidence for the requirement for caspase-2 for BCR-triggered apoptosis.

Studies with fmk derivatives of caspase substrates also strongly suggested that caspase-2 activation occurs upstream of, and is required for caspase-3 activation after BCR ligation, because a caspase-2 inhibitor blocked caspase-2 as well as caspase-3 activation, whereas a caspase-3 inhibitor prevented caspase-3, but not caspase-2 activation. In other systems, caspase-2 activation has also been reported to occur upstream of caspase-3 activation in apoptosis induced by etoposide, γ-irradiation, serum withdrawal, and treatment with atractyloside, which opens the permeability transition pore (PTP) (1, 46, 47). A caspase-2 to caspase-3 sequence is in agreement with the general conception that caspases with large CARD domains form complexes with specific proteins and are activated within such apoptotic signaling complexes, while caspases with short prodomains are directly activated and act as downstream effector caspases (48). However, a caspase-3 to caspase-2 hierarchy has been reported after CD95 ligation (43), and for caspase-2 cleavage induced in cytosolic extracts by addition of cytochrome c (49); in another study, caspase-2 was shown to be activated by a caspase-3-like activity (43). Whether a caspase-3 to -2 sequence is necessary for apoptosis remains to be determined.

It is likely that there is an intervening caspase(s) between caspase-2 and -3 in the BCR-triggered apoptotic pathway (Fig. 6), because caspase-2 is unable to activate caspase-3 in vitro (41, 50). This possibility is also supported by the finding that caspase-3 cleavage and activation after BCR ligation were blocked by crmA, which is a poor inhibitor of caspase-2 and -3 (27).

The mechanism of caspase-2 activation after BCR ligation was not addressed in these studies. There are, however, several possibilities. First, the presence of similar CARD domains in caspase-2 and -9 suggests the possibility that caspase-2 is activated by a caspase-9-like mechanism involving formation of a ternary complex with cytochrome c and Apaf-1 (26, 35). If so, however, Apaf-1 is not likely to be involved, because the CARD domains of caspase-2 and Apaf-1 do not interact (51). It is possible, however, that caspase-2 interacts with an unidentified Apaf-1-like molecule. A second possibility is that caspase-2 activation is mediated by homotypic interactions between the CARD domains of caspase-2 and an adaptor molecule, in analogy to the activation of caspase-2 by binding to the death domain-containing adaptor molecule, RAIDD/CRADD (40, 41). A third possibility is that caspase-2 is directly activated by proteolytic cleavage by another caspase. Finally, the recent demonstration that procaspase-2 is present in the intermembrane space of liver mitochondria and T cell hybridoma mitochondria, but is released in activated form after PTP opening, provides support for an autocatalytic mechanism of activation (47). Our current studies are addressing the mechanism of caspase-2 activation.

In addition to the demonstrated role of caspase-2, perhaps the most surprising finding in these studies is the lack of a major role for caspase-9 in BCR-triggered apoptosis. Caspase-9 cleavage was not detected until 12 h after BCR ligation on B104 cells, whereas caspase-2 and -3 were activated 2–4 h after BCR ligation on these cells. Thus, caspase-9 involvement is a later event in BCR-induced apoptosis than caspase-2 and -3 activation. Furthermore, DN caspase-9 only modestly inhibited BCR-induced apoptosis, indicating that it plays a relatively minor role in this apoptotic pathway. Although the mechanism of this late activation of caspase-9 was not addressed in this work, it is likely that caspase-3 is responsible, because caspase-9 cleavage after BCR ligation was blocked in cells preincubated with zDEVD-fmk, an active site caspase-3 inhibitor. Furthermore, caspase-3 has been previously reported to possess the ability to cleave caspase-9 in vitro (36). It is likely that caspase-9 functions in this system to enhance apoptosis via a feedback mechanism (Fig. 6).

Because caspase-9 is activated in the presence of cytochrome c and Apaf-1 in vitro (26, 35), the lack of early caspase-9 activation after BCR ligation suggests the possibility that cytochrome c is not released into the cytosol after BCR ligation. However, this would be surprising in view of the rapidly accumulating evidence of mitochondrial involvement and cytochrome c release as early requisite events in the response of intact cells to numerous apoptotic stimuli (38, 52, 53). Indeed, in preliminary studies, we find detectable release of cytochrome c into the cytosol 2 h after BCR ligation on B104 cells, and maximal release 4 h after ligation. The lack of early caspase-9 cleavage in B104 cells after BCR ligation is thus particularly striking, especially in view of the low concentrations of this caspase in B cells, and the recent demonstration that procaspase-9 is present in the intermembrane space in liver mitochondria together with cytochrome c (47). B cells most likely contain Apaf-1, although we have not verified this. A possible explanation for the lack of early caspase-9 cleavage after BCR ligation is that mitochondrial Bcl-XL prevents effective interaction of Apaf-1, or Apaf-1-cytochrome c complexes with the relatively small amounts of caspase-9 in B cells. Such an autoregulatory process would be analogous to the demonstrated binding of overexpressed Bcl-XL to Apaf-1, with resulting inhibition of caspase-9 activation (51, 54). Current studies are addressing this hypothesis.

In confirmation of earlier reports (29, 45), apoptosis triggered by BCR ligation was blocked by CsA. In the present studies, the CsA-dependent step was placed upstream of caspase-2 and -3 cleavage, because activation of these caspases was blocked by treatment of the cells with nanomolar CsA concentrations before BCR cross-linking. CsA binds to two intracellular proteins, cyclophilin A, a cytoplasmic protein, and cyclophilin P, an inner mitochondrial membrane protein. Cyclophilin P represents an attractive target for CsA because of its relationship to the PTP (55), which is associated with apoptotic events (56). Nevertheless, cyclophilin P is not likely to represent the target for the antiapoptotic actions of CsA, because FK506, another immunosuppressant, also inhibits BCR-induced apoptosis in several B cell types (57), but does not alter mitochondrial PTP function (55); FK506, like CsA, targets calcineurin, a calcium- and calmodulin-regulated phosphatase. Therefore, the most likely target for CsA is cyclophilin A. The CsA-cyclophilin A complex functions by binding to and inhibiting the enzymatic activity of calcineurin (58). Inhibition of calcineurin phosphatase activity by CsA and FK506 interferes dramatically with the transcription of IL-2, numerous other cytokines, and various genes involved in cellular activation (59, 60). It is possible that CsA blocks BCR-triggered caspase-2 activation and/or apoptosis by inhibiting the transcription of gene(s) required for these processes. Alternatively, CsA may function by blocking the calcineurin-mediated dephosphorylation of a regulatory protein involved in Bcl-2 or Bcl-XL interactions with death-promoting family members such as BAD (61), another type of caspase-regulating protein, or a critical upstream kinase, such as Akt or Raf-1 (62, 63). In this regard, recent evidence indicates that calcineurin promotes apoptosis by dephosphorylating BAD (64).

The findings presented in this work indicate that the BCR signaling pathway leading to cell death involves a novel calcineurin and caspase-2-, -3-, and -9-dependent pathway (Fig. 6). Further studies are clearly needed to define intervening steps, and to characterize the intracellular mechanisms that regulate cell fate decisions leading to proliferation and cell growth, or, alternatively, to cell death after BCR ligation.

We gratefully acknowledge gifts of reagents from J. Yuan (Harvard Medical School, Cambridge, MA); M. Mayumi, Fukui Medical University (Fukui, Japan); J. Jackson, The Scripps Research Institute (La Jolla, CA); X. Wang, Southwestern Medical Center (Dallas, TX); and D. Pickup, Duke University Medical Center (Durham, NC). We also thank Emanuela Bonfoco for helpful comments and Catalina Hope and Joan Gausepohl for assistance with the manuscript.

1

This work was supported in part by Grant SFP-1247 from the Novartis Corporation (to N.R.C.); Grant AG13487 from the National Institutes of Health (to E.S.A.); and Grant IRG-032 from the American Cancer Society and a grant from the American Association for Cancer Research (to H.-G.W.). W.C. is supported by U.S. Public Health Service Training Grant T32 AI07244. This is publication 11840-IMM from The Scripps Research Institute.

3

Abbreviations used in this paper: BCR, B cell Ag receptor; CARD, caspase recruitment domain; CsA, cyclosporin A; DN, dominant negative; PARP, poly(ADP-ribose) polymerase; PI, propidium iodide; PS, phosphatidylserine; PTP, permeability transition pore; zVAD-fmk, benzyloxycarbonyl-Val-Ala-Asp-fluoromethylketone; zYVAD-fmk, benzyloxycarbonyl-Try-Val-Ala-Asp-fluoromethylketone; zDEVD-fmk, benzyloxycarbonyl-Asp-Glu-Val-Asp-fluoromethylketone; zVDVAD-fmk, benzyloxycarbonyl-Val-Asp-Val-Ala-Asp-fluoromethylketone; wt, wild type; X-Gal, 5-bromo-4-chloro-3-indolyl β-d-galactoside.

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