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. Author manuscript; available in PMC: 2021 Jul 1.
Published in final edited form as: J Immunol. 2020 May 20;205(1):113–120. doi: 10.4049/jimmunol.2000359

Maintenance of Germinal Center B cells by Caspase-9 through Promotion of Apoptosis and Inhibition of Necroptosis

Jingting Zhang *, Srikanth Kodali *,†, Min Chen ‡, Jin Wang *,§
PMCID: PMC7992968  NIHMSID: NIHMS1598878  PMID: 32434938

Abstract

In response to T cell-dependent antigen encounter, naïve B cells develop into germinal center (GC) B cells, which can further differentiate into antibody-secreting plasma cells or memory B cells. GC B cells are short-lived and are prone to caspase-mediated apoptosis. However, how apoptotic caspases regulate GC B cell fate has not been fully characterized. Here we show that mice with B cell-specific knockout of caspase-9 (B/Casp9−/−) had decreases in GC B cells and antibody production after immunization. Caspase-9-deficient B cells displayed defects in caspase-dependent apoptosis but increases in necroptosis signaling. Additional deletion of receptor interacting protein kinase-3 (Ripk3) restored GC B cells and antibody production in B/Casp9−/− mice. Our results indicate that caspase-9 plays an important role in the maintenance of antibody responses by promoting apoptosis and inhibiting necroptosis in B cells.

Introduction

Germinal centers (GCs) are special regions within secondary lymphoid organs that are seeded by B cells following T cell-dependent antigen activation (1, 2). GCs are segregated into dark zones and light zones (3–5). In the dark zones, B cells undergo proliferation and somatic hypermutation of their immunoglobulin genes, after which they migrate to the light zones for antigen-affinity-based selection (3–5). The selected B cells can return to the dark zones for further proliferation and somatic hypermutation, or exit the GCs and differentiate into plasma cells or long-lived memory B cells. However, only a small fraction of B cells survive the competition and selection in the GC to become plasma cells or memory B cells (1, 6, 7). Instead, a majority of GC B cells are cleared by apoptosis (7).

Caspases are endoproteases that play essential roles in the execution of apoptosis (8, 9). Apoptotic caspases are subdivided into initiators and effectors based on their ordering in the apoptosis signaling cascade (10). Caspase-8 is an initiator caspase of the extrinsic apoptosis pathway mediated by the tumor necrosis factor receptor (TNFR) family death receptors (11, 12), while caspase-9 is the initiator caspase in the intrinsic apoptosis pathway mediated through disruption of mitochondria (13–15). The activation of the initiator caspases results in the activation of downstream effector caspases, including caspase-3, caspase-6 and caspase-7 (8, 16). While signaling from Fas and B cell receptors (BCR) can mediate apoptosis during negative selection of GC B cells (17, 18), caspase-9 is also involved in BCR-induced apoptosis in this process (18).

In addition to the induction of apoptosis, caspases have been shown to regulate lymphocyte survival and development (19, 20). Caspase-3 and caspase-6 can regulate cell cycling in B cells that is important for B cell activation and differentiation into plasma cells (21, 22). Caspase-8 is able to cleave and inactivate Ripk1 to inhibit necroptosis (23, 24), a form of programmed cell death through necrosis (25). Necroptosis signaling involves the activation of Ripk1, Ripk3 and its substrate, mixed lineage kinase like (MLKL) (26–32). Phosphoglycerate mutase family member 5 (PGAM5) can be activated by the Ripk1-Ripk3-MLKL complex to induce Drp1-mediated mitochondrial fragmentation and necroptosis (33, 34). Necroptosis plays an essential role in immune cells in response to infection (35, 36). Cell membrane rupture may cause cell contents to spill into the organ, releasing damage-associated molecular patterns (DAMPs). Immune cells are then recruited into the damaged tissues (37). Caspase-8 deficiencies lead to decreased cell proliferation in T cells in humans and mice (38–40). Such defects in T cell proliferation in caspase-8-knockout mice have been shown to be rescued by additional deletion of an essential necroptosis gene, Ripk3 (40). This suggests that caspase-8 signaling is important for the protection of cell survival and proliferation by inhibiting necroptosis.

Knockout studies suggest that caspase-9 is essential for mitochondrion-dependent apoptosis induced by various stimuli (41, 42). Caspase-9 can also inhibit inflammatory responses by promoting immunologically silent apoptosis (43, 44), suggesting that caspase-9 may also contribute to the protection of cell survival. In this study, we show that GC B cell numbers and primary antibody responses are decreased in mice with B cell-specific deletion of caspase-9 (B/Casp9−/−). B cells deficient in caspase-9 displayed elevated levels of necroptosis signaling, while deletion of Ripk3 rescued GC B cells in B/Casp9−/− mice. Our results suggest that caspase-9 protects GC B cells and antibody responses by inhibiting necroptosis.

Materials and Methods

Mice

Two loxP sites were inserted into both sides of exon 6 of caspase-9 in a 9.3 kB Hind III genomic DNA from a BAC clone (BACPOAC Resources). The construct was used for homologous recombination in embryonic stem cells to generate caspase-9-flox mice. CD19-cre mice (The Jackson Laboratory) were crossed with caspase-9-flox mice for B cell-specific deletion of caspase-9 (B/Casp9−/−). Ripk3−/− mice (45) were crossed with B/Casp9−/− mice to generate Ripk3−/−B/Casp9−/− mice. Sex- and age-matched mice, 6-8 weeks old and on the C57BL/6 background, were used for all experiments. Mice were bred and maintained in a specific pathogen-free facility at Houston Methodist Research Institute. Experiments were performed according to federal and institutional guidelines and with the approval of the Institutional Animal Care and Use Committee.

Immunization

Mice were immunized with 100 μg of NP-KLH (Biosearch Technologies, Novato, CA) precipitated with 100 μl Imject Alum adjuvant (Thermo Fisher, Waltham, MA) by intraperitoneal injection. Mice were used for various analyses two weeks after immunization (46).

Flow cytometric analysis

Spleen cells were isolated and red blood cells were lysed with ammonium chloride lysing buffer (0.15 M NH4Cl, 10 mM KHCO3, 0.1 mM EDTA). Splenocytes were then used for various analyses. To detect NP-specific GC B cells, splenocytes were stained with V450-anti-IgM (BD Biosciences, San Jose, CA), Pacific Blue-anti-CD4, Pacific Blue-anti-CD8a, Pacific Blue-anti-CD11b, Pacific Blue-anti-CD11c, Pacific Blue-anti-Gr-1, Pacific Blue-anti-IgD, Brilliant Violet 421-anti-CD138, APC/Fire-750-anti-B220, Alexa Fluor 647-anti-IgG1, PerCP/Cy5.5-anti-GL-7, PE/Cy7-anti-CD38 (BioLegend, San Diego, CA), and 4-Hydroxy-3-nitrophenylacetyl (NP) conjugated with-PE (PE-NP; Santa Cruz Biotechnology, Dallas, TX) for 15 min on ice. For intracellular staining of active caspases, splenocytes were stained with surface markers, fixed with 1% methanol-free formaldehyde (Thermo Fisher, Waltham, MA) for 15 min on ice, permeabilized with the Permeabilization Buffer (Thermo Fisher, Waltham, MA) for 15 min on ice, and incubated with rabbit antibodies specific for mouse cleaved caspase-9 or cleaved caspase-3 (Cell Signaling Technology, Danvers, MA) for 15 min on ice. Cells were then stained with Alexa Fluor 647-goat anti-rabbit IgG (Thermo Fisher, Waltham, MA) for 15 min on ice. The cells were analyzed by flow cytometry on an Fortessa flow cytometer (BD Biosciences, San Jose, CA).

B cell purification and Western Blot

B cells were purified from splenocytes using CD19 Microbeads (Miltenyi Biotec, Bergisch Gladbach, Germany) and stimulated with 20 ng/mL IL-4 and 1 μg/mL LPS in RPMI 1640 complete medium for 3 days (47). Cells were then cultured in RPMI 1640 medium with 0.5% FBS and harvested at 0 h, 8 h, and 16 h for lysis in sample buffer (50 mM HEPES, 150 mM NaCl, 1 mM EDTA, 1% Triton X-100, 10 μM zVAD-FMK and 1× protease/phosphayase inhibitor cocktail ( Cell Signaling Technology, Danvers, MA). Cell lysates were used for Western blot and probed with rabbit antibodies against caspase-9, cleaved caspase-9, caspase-3, cleaved-caspase-3, caspase-1, cleaved-caspase-1, gasdermin D, cleaved gasdermin D, Ripk1, Phospho-Ripk3, Ripk3 (Cell Signaling Technology, Danvers, MA), MLKL (WuXi AppTec, Shanghai, China), or Pgam5 (Abcam, Cambridge, UK) followed by incubation with HRP-conjugated goat anti-rabbit IgG (Abcam, Cambridge, UK) and development with SuperSignal Western Pico enhanced chemiluminescence substrate (Pierce Chemical Co, Dallas, TX). The blots were also probed with anti-β-Actin (Santa Cruz Biotechnology, Dallas, TX) as loading controls. Wild type and caspase-9−/− B cells were also cultured with 1 μg/ml biotinylated anti-Fas (BD Biosciences, San Jose, CA) and 0.1 μg/ml streptavidin (Jackson ImmunoResearch, West Grove, PA) for the indicated time. Cells were also used for staining with FITC-conjugated anti-Fas (BD Biosciences, San Jose, CA) for flow cytometry, or lysed for Western blot to detect caspases. Alternatively, activated B cells were treated with or without 20 μM Z-IETD and 20 ng/ml TNF-α (R&D System, Minneapolis, MN) for 4 h. Z-IETD was added 30 min prior to TNF-α. Cells were lysed for Western blot to detect necroptosis-related proteins.

Cell death analyses

To determine spontaneous cell death, splenocytes were collected and cultured in RPMI medium with 0.5% fetal bovine serum for 4 h as described (46). Cells were then incubated with FITC-DEVD-FMK (SM Biochemicals LLC, Anaheim, CA) at 37°C for 30 min and then stained with biotin anti-CD4, biotin anti-CD8a, biotin anti-CD11b, biotin anti-CD11c, biotin anti-Gr-1, biotin anti-IgD, biotin anti-CD138, Alexa Fluor 700 anti-B220, Alexa Fluor 647 anti-IgG1, PerCP/Cy5.5 anti-GL-7, and PE/Cy7 anti-CD38, followed by staining with APC/Fire-750 streptavidin (BioLegend, San Diego, CA). Cells were then labeled with Alexa Fluor 350 Annexin V conjugate (Thermo Fisher, Waltham, MA) at room temperature for 15 min. Cells were analyzed by flow cytometry on an Fortessa (BD Biosciences, San Jose, CA). To determine necroptosis-related cell death, activated B cells were cultured in the presence of 20 μM Z-IETD and 20 ng/ml TNF-α (R&D System, Minneapolis, MN) with or without 20 μg/ml Necrostatin-1 (Enzo Life Science, Farmingdale, NY) for 20 h. Propidium iodide (PI) was added and the cells were analyzed by flow cytometry. PI− viable B cells were quantified and the percentage of cell death was calculated as described (48): % cell death = (untreated-treated)/untreated x 100%.

ELISA

Clear polystyrene 96-well microplates (Corning Inc, Corning, NY) were coated with 5 μg/mL NP5-BSA or NP25-BSA (Biosearch Technologies, Novato, CA) overnight at 4°C. After blocking with PBS containing 0.05% Tween-20 at 37 °C for 2 h, diluted serum samples were added and incubated at 4 °C overnight. The plates were incubated with HRP-conjugated goat anti-mouse IgG1 (Southern Biotechnology, Birmingham, AL) for 1 h, followed by development using TMB Peroxidase EIA Substrate Kit (Bio-Rad Laboratories, Hercules, CA). The reactions were stopped with 1N H2SO4 and the absorbance of each well was measured using a Synergy H1 Hybrid Multi-Mode Microplate Reader (BioTek) at 450 nm. Antibody titers were calculated as described (46).

ELISPOT

MultiScreen 96-well Filtration plates (Millipore, Burlington, MA) were pre-coated with 20 μg/mL NP5-BSA or NP25-BSA at 4 °C overnight. After blocking with RPMI 1640 complete medium for 2 h at 37 °C, cells (1-5×105 /well) were then added to the plates and incubated at 37 °C overnight. The cells were lysed with H2O, and then incubated in RPMI 1640 complete medium for 2 h at room temperature. The plates were incubated with HRP-conjugated goat anti-mouse IgG1 (Southern Biotechnology, Birmingham, AL), followed by development with 3-amino-9-ethylcarbazole (Sigma-Aldrich, St. Louis, MO) and stopped by washing with PBS. The stained dots on the filters were quantified using an Immunospot S6 ULTIMATE Analyzer (ImmunoSpot, Cleveland, OH).

Statistical analyses

Statistical analyses were performed by two-tailed Student’s t-test, or by Dunnett’s test (in Fig. 8) using GraphPad Prism 8 (GraphPad, San Diego, CA). The results of ELISA, ELISPOT and flow cytometry were presented as the mean ± SD, with P<0.05 considered statistically significant.

FIGURE 8.

FIGURE 8.

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Deletion of Ripk3 rescues antigen-specific Casp9−/− GC B cells. (A) ELISA for high-affinity (NP5-BSA) anti-NP IgG1 and total (NP25-BSA) anti-NP IgG1 relative titer in WT, B/Casp9−/−, Ripk3−/−, and DKO mice 2 weeks after immunization with NP-KLH.

(B) ELISPOT for NP-specific IgG1 ASCs at 2 weeks post-immunization with NP-KLH.

(C) Flow cytometric analysis of DUMP−B220+IgG1+NP+GL-7+ CD38− GC B cells in the spleen of WT, B/Casp9−/−, Ripk3−/−, and DKO mice at 2 weeks post-immunization with NP-KLH. (D) Schematic mode of GC B cell death pathway. In response to cytochrome c release from mitochondria, caspase-9 cleaves executioner caspases to drive apoptosis, while also inhibiting Ripk1/Ripk3 signaling to block necroptosis. In the absence of caspase-9, GC B cells shift from apoptotic to necroptotic cell death. Data in this Figure were analyzed by Dunnett’s test from three independent experiments with 3 or more mice per group. *P<0.05, **P<0.01.

Results

Caspase-9 signaling in GC B cells

GC B cells are prone to caspase activation and apoptosis in the absence of survival signals (46). To examine caspase-9 signaling in GC B cells, we cultured total spleen B cells from unimmunized mice under low serum conditions to induce spontaneous cell death as described (46). We found that naturally occurring IgG1+ GC B cells, but not naïve or IgG1+ memory B cells, displayed the activation of caspase-9 and downstream caspase-3 (Fig. 1A). We also examined caspase signaling in B cells from mice immunized with 4-hydroxy-3-nitrophenylacetyl-keyhole limpet hemocyanin (NP-KLH). Consistently, the activation of caspase-9 and caspase-3 was detected in NP-specific IgG1+ GC B cells but not memory B cells (Fig. 1B). These data indicate that caspase-9 signaling can be rapidly activated in GC B cells.

FIGURE 1.

FIGURE 1.

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GC B cells are prone to caspase-9 signaling. A, B Intracellular staining of active caspase-9 and caspase-3 in (A) B220+CD23hiIgD+IgMlo naïve B cells, IgD−IgM−CD11b−CD11c−Gr-1−CD4−CD8a−CD138− (DUMP−) B220+IgG1+GL-7+CD38− GC B cells, and DUMP−B220+IgG1+GL-7−CD38+ memory B cells in unimmunized mice and (B) NP-specific GC and memory B cells in mice immunized with NP-KLH. Cells were cultured in vitro for 0 h (gray background) or 4 h (black line). Percentages of naïve, GC, and memory B cells with active caspase-9 or caspase-3 are also shown (lower panels). Data are representative of three independent experiments. **P<0.01 (n=3).

Reduced GC B cells and antibody production in the absence of caspase-9

To investigate the function of caspase-9 in immune regulation, we generated mice with a floxed allele of caspase-9 (Fig. 2A). These mice were bred with CD19-cre mice for B cell-specific deletion of caspase-9 (B/Casp9−/−). B/Casp9−/− mice exhibited no significant differences in the numbers of transitional 1 (T1), transitional 2 (T2), and mature naïve B cells in the spleen (Fig. 2B), indicating that caspase-9 is dispensable for normal B cell development. To determine whether GC B cell numbers were changed in the absence of caspase-9, we immunized wild type and B/Casp9−/− mice with NP-KLH. Surprisingly, we found that NP+IgG1+ GC B cells were decreased in B/Casp9−/− mice despite impaired caspase-9-dependent signaling (Fig. 2C). These data suggest that caspase-9 is not required for B cell development, but important for maintaining normal GC B cell numbers after immunization.

FIGURE 2.

FIGURE 2.

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GC B cells are decreased in the absence of caspase-9. (A) CD19-cre-mediated excision of the Casp9 gene to generate a conditional KO mouse model. Western blot of purified splenic B cell extracts from WT and B/Casp9−/− mice. (B) Flow cytometric analysis of DUMP−B220−CD23+IgMhiIgDlo T1, DUMP−B220+CD23−IgMhiIgDhi T2, and DUMP−B220+CD23+IgMloIgDhi M B cells in the spleens of unimmunized mice. Bar charts show the number of T1, T2, and M B cells per 106 spleen cells. (C) Flow cytometric analysis of DUMP−B220+IgG1+NP+GL-7+CD38− GC B cells in the spleens of B/Casp9−/− or WT mice 2 weeks after immunization. Bar charts show the number of NP-specific GC B cells per 106 spleen cells. Data in (B) and (C) are representative of three independent experiment with 3 to 4 mice per group as indicated. ns, statistically not significant. **P <0.01.

We next measured antibody production in the absence of caspase-9. Consistent with a decrease in NP-specific GC B cells, B/Casp9−/− mice exhibited a decrease in the production of NP-specific antibodies two weeks after immunization with NP-KLH, including the production of high-affinity and total anti-NP IgG1 (Fig. 3A). Since class-switched antibody-secreting cells (ASCs) are generated by the GC reaction (49), we next examined the numbers of these cells in the absence of caspase-9. We found that B/Casp9−/− mice produce fewer antigen-specific ASCs by ELISPOT (Fig. 3B). Consistently, antigen-specific CD138+ plasma cells were reduced in B/Casp9−/− mice (Fig. 3C). Taken together, our data indicate that caspase-9 deficiency leads to a decrease in GC B cells, resulting in reduced antibody response after immunization.

FIGURE 3.

FIGURE 3.

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Impaired primary antibody response in the absence of caspase-9. (A) ELISA for high-affinity (NP5-BSA) anti-NP IgG1 and total (NP25-BSA) anti-NP IgG1 relative titers at 2 weeks post-immunization. (B) ELISPOT for NP-specific IgG1 antibody-secreting cells (ASCs) at 2 weeks post-immunization. Bar charts show the numbers of ASCs per 106 spleen cells. (C) Flow cytometric analysis of DUMP−B220+CD138+ IgG1+ NP+ plasma and plasmablast cells in the spleen at 2 weeks post-immunization. Data are representative of three independent experiments with 3 to 5 mice per group as indicated. *P<0.05, **P<0.01.

Decreased apoptosis in GC B cells in the absence of caspase-9.

To determine whether the caspase signaling cascade is indeed absent in caspase-9-deficient B cells, we purified and cultured B cells from wild type and B/Casp9−/− mice with reduced serum to induce cell death in vitro according to our previous protocol (46). We detected active caspase-9 and caspase-3 in wild type B cells, but not in in caspase-9-deficient B cells (Fig. 4A). Moreover, caspase-9-deficient GC B cells showed less apoptotic activities compared to wild type controls as indicated by cleavage of DEVD (Fig. 4B). These data indicate that caspase-9 deficiency leads to decreased apoptosis under the condition of serum starvation conditions in vitro.

FIGURE 4.

FIGURE 4.

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Caspase-9-deficiency leads to decreased apoptosis in GC B cells. (A) Western blot analysis of apoptosis-related proteins in purified wild type (WT) and Casp9−/− (KO) B cells after 3 days of stimulation and culture in low serum for 0 h, 8 h, and 16 h. Data are representative of three independent experiments. (B) Flow cytometric analysis of DUMP−B220+IgG1+GL-7+CD38− GC B cells stained by DEVD-FITC after 9 h of low serum culture. Quantification of the percentage of cells positive for DEVD cleavage in WT and Casp9−/− GC B cells after 3, 6 or 9 h of culture. Data are representative of three independent experiments with 3 mice per group. *P<0.05, **P<0.01.

Caspase-9-deficient B cells display decreased cleavage of Ripk1

Caspase-8 can inhibit necroptosis by cleavage of Ripk1, and caspase-8 deficiency leads to increased necroptotic cell death (50, 51). We therefore tested whether caspase-9 could similarly suppress necroptosis by examining necroptotic signaling molecules (35). We detected the cleaved and inactive form of Ripk1 in wild type B cells, but not in caspase-9-deficient B cells after 8 to 16 hours of in vitro culture (Fig. 5A). Moreover, full-length Ripk1 was higher in caspase-9-deficient B cells after in vitro culture (Fig. 5A), indicating that necroptosis is more active in the absence of caspase-9.

FIGURE 5.

FIGURE 5.

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Necroptosis signaling in Casp9−/− B cells. (A) Western blot for necroptosis-related proteins in purified WT and Casp9−/− (KO) B cells. Relative levels of cleaved Ripk1 were analyzed by quantification of the density of the protein bands with NIH ImageJ software. The ratios of intensities of cleaved Ripk1 versus β-Actin were: 0.20 (0 h WT), 0.05 (0 h KO), 0.54 (8 h WT), 0.09 (8 h KO), 0.79 (16 h WT) and 0.09 (16 h KO); full length Ripk1 versus β-Actin: 0.79 (0 h WT), 0.82 (0 h KO), 0.27 (8 h WT), 0.44 (8 h KO), 0.22 (16 h WT) and 0.43 (16 h KO). (B) Western blot for pyroptosis-related proteins in samples as in (A). **Degraded or non-specific band. (C) WT and Casp9−/− B cells were stained with FITC-anti-Fas (dark line) and analyzed by flow cytometry. Grey line, unstained control. (D) Induction of cell death in WT and Casp9−/− B cells after crosslinking with anti-Fas. ns, statistically not significant (n=3). (E) Western blot for caspases-8 and caspase-3 in WT and Casp9−/− B cells after different time of Fas-crosslinking. Western blot data are representative of two independent experiments.

Caspase-1, an inflammatory caspase, has been shown to trigger pyroptosis, a form of inflammatory cell death mediated by cleavage of gasdermin D (52–54). We determined whether pyroptosis might be responsible for reduced GC B cell numbers in the absence of caspase-9. However, we did not detect the activation of caspase-1 or cleavage of gasdermin D in caspase-9-deficient B cells or wild type controls following induction of spontaneous cell death during in vitro culture (Fig. 5B). These results indicate that pyroptosis does not serves as an alternate cell death pathway in caspase-9-deficient B cells.

It has been established that B cells are susceptible to Fas-dependent killing by Fas-ligand-expressing T cells (48, 55, 56). Because caspase-8 and caspase-9 are the initiator caspases in the extrinsic and intrinsic apoptosis pathways, respectively, caspase-9 deficiency is unlikely to affect caspase-8-dependent extrinsic apoptosis mediated by Fas. We therefore determined whether Fas-mediated activation of caspase-8 signaling is affected in caspase-9-deficient B cells. We observed that Fas expression and Fas-mediated cell death were comparable between wild type and caspase-9-deficient B cells (Fig. 5, C and D). After Fas-crosslinking, the activation of caspase-8 and caspase-3 was similar between wild type and caspase-9-deficient B cells (Fig. 5E), suggesting that Fas-mediated apoptosis is normal in caspase-9-deficient B cells. Together, our data indicate that caspase-9-deficient B cells display defects in intrinsic apoptosis but are normal Fas-mediated apoptosis. Caspase-9 deficiency does not cause the activation of caspase-1-dependent pyroptosis in B cells but decreases the cleavage of Ripk1, leading to increased Ripk1 levels.

Increased necroptosis signaling in the absence of caspase-9.

TNF-induced necroptosis regulated by Ripk3 is the primary pathway for necroptosis (26, 29). It has been shown that TNF-α can induce necroptosis in the presence of caspase inhibitors (57, 58). We found that TNF-α could induce cell death in wild type B cells in the presence of a caspase-8 inhibitor, Z-IETD (Fig. 6A). Interestingly, caspase-9-deficient B cells underwent increased cell death (Fig. 6A). Moreover, a necroptosis inhibitor necrostatin-1 (59), significantly inhibited the cell death in caspase-9-deficient B cells (Fig. 6A), indicating cell death in caspase-9-deficient B cells involve necroptosis. Consistently, we found that phosphorylation of Ripk3 was increased in caspase-9-deficient B cells compared with wild type B cells (Fig. 6B), indicating an increase in necroptosis signaling. These results suggest that B cells display increased necroptosis in the absence of caspase-9.

FIGURE 6.

FIGURE 6.

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Necroptosis in Casp9−/− B cells. (A) Quantification of the percentage of cell death in WT and Casp9−/− (KO) B cells after incubation with Z-IETD and TNF-α with or without necrostatin-1 (Nec-1) for 20 h. Data are representative of two independent experiments with 3 mice per group. **P<0.01.

(B) Western blot analysis of necroptosis-related proteins in WT and Casp9−/− (KO) B cells after 3 days of stimulation and treated with Z-IETD and TNF-α for 4 h. Data are representative of two independent experiments.

Deletion of Ripk3 rescues antigen-specific caspase-9-deficient GC B cells.

The above data suggest that decreased cleavage of Ripk1 in caspase-9-deficient B cells results in an increase in full-length Ripk1 compared to wild type controls (Fig. 5A). Moreover, necroptosis is elevated in caspase-9-deficient B cells as shown by increase phosphorylation of Ripk3 (Fig. 6B). It has been shown that necroptosis can be triggered in caspase-8-deficient T cells to compensate for impaired apoptosis (40). We investigated whether necroptosis might contribute to the decreased numbers of caspase-9-deficient GC B cells in vivo. Because Ripk3 is a key mediator of necroptosis (29, 30), we crossed B/Casp9−/− mice with Ripk3−/− mice to generate mice with double knockout (DKO) of caspase-9 and Ripk3 in B cells. We first determined whether of Ripk3 affects GC B cell apoptosis. We found that Ripk3−/− and WT GC B cells showed similar levels of cell death after serum starvation in vitro (Fig. 7). Moreover, GC B cells from B/Casp9−/− and DKO mice also showed similar levels of reduction in cell death (Fig. 7). This indicates that GC B cell apoptosis after serum starvation in vitro is affected by caspase-9 but not Ripk3. We then immunized B/Casp9−/−, Ripk3−/−, DKO mice and wild type controls with NP-KLH and examined anti-NP GC B cell numbers and primary antibody responses. We found that deletion of Ripk3 rescued anti-NP IgG1 antibodies in Ripk3−/−B/Casp9−/− mice (Fig. 8A). Moreover, the numbers of NP-specific ASCs were also rescued in Ripk3−/−B/Casp9−/− mice compared to B/Casp9−/− mice (Fig. 8B). Consistent with these results, NP-specific IgG1+ Casp9−/− GC B cells were also rescued by the deletion of Ripk3 (Fig. 8C). These results indicate that caspase-9 plays a protective role for GC B cells by limiting necroptotic cell death. Our results reveal an important role for caspase-9 in maintaining a balance between apoptosis and necroptosis to protect the homeostasis of GC B cells in antibody responses (Fig. 8D).

FIGURE 7.

FIGURE 7.

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Deletion of Ripk3 has no effect on GC B cell apoptosis. Flow cytometric analysis of DUMP−B220+GL-7+CD38− GC B cells stained by DEVD-FITC and Annexin V after 6 h of low serum culture. Quantification of the percentage of DEVD+Annexin V+ cells in WT, B/Casp9−/−, Ripk3−/−, and DKO GC B cells after 0, 3, 6 or 9 h of culture. Data are representative of two independent experiments with 3 mice per group.

Discussion

In our study, we found that GC B cell numbers are decreased in caspase-9-deficient mice despite a defect in apoptosis. We observed that the cleaved inactive form of the necroptosis mediator Ripk1 was decreased in caspase-9-deficient B cells. Moreover, caspase-9-deficient B cells display increased potential for necroptosis signaling. By comparison, we did not detect a significant cleavage of the pyroptosis signaling molecules, including caspase-1 and gasdermin D in wild type and caspase-9-deficient B cells. Consistent with increased necroptosis, deletion of an essential necroptosis gene, Ripk3, rescued GC B cell numbers and antibody production in B/Casp9−/− mice. Our findings suggest that caspase-9 promotes apoptosis and inhibits necroptosis to maintain the homeostasis of GC B cells.

GC B cells can undergo apoptosis through either the cell-intrinsic or extrinsic pathways. After engagement by Fas ligand expressed by activated T cells, Fas on GC B cells can trigger the extrinsic apoptosis pathway through activation of caspase-8 (48, 55, 56, 60). We found that the expression of Fas, as well as Fas-mediated caspase-8 activation and apoptosis were normal in caspase-9-deficient B cells (Fig. 7C-E). This suggests that caspase-8-dependent extrinsic apoptosis of B cells is not affected by the loss of caspase-9. Caspase-9 that mediates the intrinsic pathway has been implicated in BCR-induced apoptosis (18). Therefore, caspase-9 may be required for BCR-induced intrinsic apoptosis but not Fas-mediated extrinsic apoptosis.

In addition to their long-established roles as mediators of the pro-apoptotic proteolytic cascade, caspases have more recently been recognized for other functions. Caspase-9 has been shown to inhibit type I interferon signaling, a mechanism of anti-viral immunity that results in inflammatory cell death (43). Caspase-8 has been demonstrated to negatively regulate necroptosis in T cells, with its absence resulting in decreased T cell numbers, a phenotype that is rescued by ablation of Ripk3 (40). Interesting, however, GC B cell-specific deletion of Fas or caspase-8 has been shown to cause GC B cell accumulation (60, 61). This suggests that the extrinsic and intrinsic apoptotic caspases have distinct, non-redundant functions in GC B cell homeostasis.

Necroptosis is a programmed form of necrosis that triggers inflammation (35). It may be favorable for a virus-infected cell to die by necroptosis, both to eliminate the viral reservoirs and to provoke a host anti-viral immune response. However, excessive inflammation may be detrimental to the host. Apoptosis is an immunologically silent form of cell death. Caspase-8 has been showed to play an important role in the suppression of necroptosis (23, 51). Our results suggest that caspase-9 also plays an important role in limiting necroptosis in germinal center B cells.

Programmed cell death plays an essential role in the regulation of germinal center responses. This may help to prevent the entry of deleterious GC B cell clones generated from somatic hypermutation into the long-lived memory B cell and plasma cell pools. We have observed that GC B cells rapidly induce caspase-9 signaling during in vitro culture. However, the contributions of apoptotic and non-apoptotic cell death in the regulation of GC B cell homeostasis is not clear. This study suggests caspase-9 not only help to delete GC B cells by apoptosis, but also prevents excessive necroptosis to maintain the proper numbers of GC B cells for antibody responses. Our data reveal an important role for caspase-9 in maintaining a balance between apoptosis and necroptosis to protect GC B cell survival and antibody production.

Key points:

Caspase-9 promotes apoptosis in GC B cells

Caspase-9 inhibits necroptosis in GC B cells.

Acknowledgments

We thank Min Li, Wei Liu and Francis Chan for technical support, Vishva Dixit for Ripk3−/− mice, and Xiulong Xu for discussions.

This study was supported by funding from the NIH R01AI116644 and R01AI123221 to J.W., the Cancer Prevention and Research Institute of Texas RP160384 to J.W. and M.C.

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