Differing Requirements for MALT1 Function in Peripheral B Cell Survival and Differentiation

Activation of NF-κB has emerged as one of the most crucial steps in mounting an effective immune response, regulating a wide array of genes essential for immune cell survival and function. NF-κB signaling can occur via the canonical route, in which the major players are NF-κB dimers composed of RelA, c-Rel, and p50, or via the alternative non-canonical route that is mediated by the Relb and p52 heterodimers (1). The canonical NF-κB pathway becomes transiently activated after Ag receptor engagement via the assembly of a signaling platform composed of scaffold proteins CARMA-1 and BCL-10, and the paracaspase MALT1 (termed the CBM complex), which relays signals from proximal kinases and adaptors to the core IκB kinase complex (2, 3). Gene-targeting studies have revealed that mice deficient in any of the components of the CBM complex show defective B cell and T cell activation, resulting in an inadequate adaptive immune response (4). In the B cell compartment, Malt1^−/− mice exhibit a reduction in the marginal zone (MZ) and B1 B cell subsets, decreased serum IgM and IgG3 levels, as well as impaired Ab responses to both T dependent (TD) and T independent (TI) Ags (5, 6). A subsequent study unraveled a new role for MALT1 in mediating BAFF-induced non-canonical NF-κB signaling specifically in MZ B cells, where MALT1 deficiency resulted in diminished BAFF-induced p100 processing and cell survival (7). Although the detailed mechanism has not been completely elucidated, the defect in MALT1-deficient B cells has been associated with nuclear translocation of c-Rel upon triggering of the BCR (8).

Germinal centers (GCs) are specialized microenvironments within the secondary lymphoid tissues that are formed at the height of an ongoing immune response and are critical for the optimization of TD humoral immune responses. Ag-activated B cells entering the GC proliferate rapidly and undergo somatic hypermutation and affinity maturation, ultimately leading to the generation of memory B and plasma cells producing high-affinity Abs (9). This process requires intricate interactions among Ag-specific B and T cells, and follicular (FO) dendritic cells (10). The initiation of the GC reaction is also dependent on a multitude of B cell-intrinsic factors modulating the BCR signal and, notably, deletion or mutation of NF-κB family members often abrogates GC formation (11). For example, c-Rel–deficient mice exhibit a severe impairment in GC formation upon immunization, due to its critical role in regulating B cell survival and cell cycle progression (12, 13). Also, it has recently been shown that c-Rel is essential for the maintenance of GC B cells via activation of a metabolic program that promotes cell growth (14). Histologic staining of the spleen in Malt1^−/− mice immunized with TD Ag revealed a complete absence of GC formation (5). Furthermore, MALT1-deficient animals are devoid of spontaneously formed GC B cells in the Peyer’s patches, which are chronically exposed to Ags because of their anatomical location (15). The lack of GC B cells also correlates with a severe reduction in T FO helper (T_FH) cells (15), because it has been shown that GC B cells are required to sustain the T_FH phenotype (16).

For the initiation and progression of the GC response, T cell help is crucial. T cells can participate through the engagement of CD40 on B cells and via signals delivered through costimulatory molecules and cytokines (17, 18). It has been consistently reported that Malt1^−/− T cells exhibit impairment in activation, proliferation, and IL-2 production as well as NF-κB activation in response to TCR ligation (5, 6). However, it remains unclear whether the inability of the Malt1^−/− animals to mount GC reactions can be entirely attributed to the defective T cell response. Moreover, the role of MALT1 in B cell activation remains controversial with respect to the requirement of MALT1 in BCR-stimulated proliferation and NF-κB activation (5, 6). Moreover, despite the established link to c-Rel translocation, Malt1^−/− animals do not fully phenocopy mice with c-Rel deletion, suggesting that MALT1 may regulate additional mechanisms in B cells.

Loss-of-function mutations in MALT1 have also been discovered in human patients with combined immunodeficiency disorders, resulting in abnormal T cell proliferation and failure to activate NF-κB following treatment with the direct protein kinase C (PKC) activator PMA (19–21). Yet, the contribution of B cells to the disease manifestations in these patients is still unclear. In contrast, deregulated expression of MALT1 has been implicated in B cell lymphomagenesis, and a recent study demonstrated the oncogenic potential of MALT1 in driving B cell lymphopoiesis (22). Consequently, understanding the role of MALT1 in B cells is crucial from a clinical perspective, for the management of diseases including immunodeficiency and lymphoma.

To our knowledge, in this study we provide the first evidence that MALT1 is required in B cells for the GC response. MALT1 is critical for efficient proliferation and survival in response to BCR signaling. Consequently, initiation of the GC response upon immunization is impaired in the absence of MALT1. We also show that T cell-derived help fails to rescue the proliferation and survival defects in MALT1-deficient B cells. Lastly, we demonstrate that ectopic expression of Bcl-2 in MALT1-deficient B cells can partially rescue the GC phenotype by prolonging the survival of the B cells, but there remains an intrinsic block in plasma cell differentiation, as well as Ab production.

The Malt1^−/− mice have been previously described (5) and were kindly provided by Dr. Vishva Dixit (Genentech). Littermate controls of Malt1^+/+ or Malt1^+/− genotypes were generated along with Malt1^−/− mice by interbreeding heterozygous animals. Bcl-2 transgenic mice (Eμ-bcl-2-22) used for breeding with Malt1^−/− animals were purchased from The Jackson Laboratory. The mice were housed in a pathogen-free environment in the animal facility at the Sanford Burnham Prebys Medical Discovery Institute. All experiments conformed to the ethical principles and guidelines approved by the Institutional Animal Care and User Committee.

For generation of B cell specific wild type (WT) and Malt1^−/− mice, μMT mice (The Jackson Laboratory) were sublethally irradiated (5-Gy dose) and reconstituted with bone marrow (BM) from Malt1^+/+ or Malt1^−/− mice. Mixed chimeras were generated as follows: μMT animals were exposed to 10-Gy of irradiation and reconstituted with a 1:1 mix of BM from CD45.1 and Malt1^−/− mice. BM from one to two donor mice was divided among four to six recipient animals.

Citrated SRBCs (Colorado Serum Company) were washed twice with PBS and resuspended in PBS to a final concentration of 10% (v/v). Animals were injected i.p. with 0.2 ml of SRBC suspensions. Sera were collected on day 0 and day 7 postimmunization for measuring the levels of SRBC-specific IgM and IgG1 levels using a flow cytometry-based method. Briefly, SRBCs were incubated with varying dilutions of the serum samples, washed and detected with fluorescent conjugated Abs against either IgM (clone II/41; eBioscience) and IgG1 (cloned A85.1; BD Biosciences). Mean fluorescence intensities of the SRBC-bound αIgM and αIgG1 Abs were plotted against the dilution factors, and the values in the linear range were used for comparing the relative Ab titers.

Splenic B cells were isolated from CD45.1 or Malt1^−/− animals by CD43 depletion and mixed at a 1:1 ratio. Then 1–2 × 10⁶ B cells were transferred into the μMT mice via the tail vein. After 24 h, the μMT mice were immunized with SRBCs and analyzed on day 7 postimmunization.

Single-cell suspensions prepared from spleens and Peyer’s patches were blocked with anti-CD16/32 (clone 2.4G2; BD Biosciences) and stained with the indicated combination of conjugated Abs for 30 min on ice. Live cells were assessed by forward and side scatter profiles. All cells were acquired on a FACSCanto flow cytometer using the FACSDiva software (BD Biosciences) and data were analyzed using FlowJo software (Tree Star). The following Abs were obtained from eBioscience: anti-B220 (RA3-6B2), -CD3e (145-2C11), -CD21/35 (8D9), -CD23 (B3B4), -CD45.1 (A20), -CD4 (RM4-5), -IgD (11–26), -CD86 (PO3.1), -PD1 (J43), -ICOS (7E.17G9), -IgM (II/41), -CD5 (53.753), -CD62L (MZL-14), -CD86 (PO3.1), -MHC class II (M5/114.15.2), -CD25 (PC61.5), and -CD69 (H1.2F3). Anti-GL7, -FAS (JO2), -CD45.2 (104), -CD138 (281-2), -IgG1 (A85.1), and -CD80 (16-10A1) Abs were purchased from BD Biosciences.

B cells were isolated from splenocytes by negative magnetic-based sorting of cells labeled with CD43 microbeads (Miltenyi Biotec). For the proliferation assay, purified B cells were labeled with eFluor670 (eBioscience) according to the manufacturer’s protocol and cultured in 96-well plates at a density of 10⁶ cells/ml in complete RPMI 1640 (Cellgro; Corning) supplemented with 10% FBS (Sigma), 1× Penicillin/Streptomycin (Cellgro; Corning), 2 mM GlutaGro (Cellgro; Corning), 1× MEM non-essential amino acids (Cellgro; Corning), 1 mM sodium pyruvate (Cellgro; Corning), and 50 μM β-mercaptoethanol (Life Technologies), with or without various stimuli. The following stimuli were used at the indicated concentrations: 25 ng/ml recombinant murine BAFF (R&D Systems), 10 ng/ml recombinant IL-4 (eBioscience), 5 μg/ml αCD40 (eBioscience), 10 μg/ml αIgM F(ab′)₂ (Jackson ImmunoResearch), and 10 μg/ml LPS (Sigma). To assess the effect of caspase inhibition on proliferation, B cells were stimulated in the presence of the pan caspase inhibitor IDN-6556 at 10 μM. For cell cycle analysis, splenic B cells were stimulated with the indicated stimuli and duration, and pulsed with 10 μM of BrdU (Life Technologies) during the last hour of incubation prior to harvest. The cells were fixed with 70% ice-cold ethanol, treated with 2 M HCl to denature the DNA, and washed with 0.1 M sodium tetraborate (pH 8.5) to neutralize the acid. The cells were then incubated with αBrdU (BD Biosciences), resuspended in a mixture containing RNase at 100 μg/ml and 7-aminoactinomycin D (7-AAD) at 5 μg/ml, and analyzed by flow cytometry. For combined apoptosis and proliferation assays, eFluor670-labeled B cells were cultured as indicated and stained with Annexin-V-FITC (BioVision) according to the manufacturer’s instructions.

Spleens were embedded in Tissue-TEK OCT compound (Sakura Finetek) and frozen at −80°C. Frozen tissue blocks were sectioned, mounted on Superfrost/Plus slides (Fisher Scientific), fixed in ice-cold acetone, and blocked with PBS with 5% FBS, plus the inhibitor E-64 at 10 μM to block endogenous cysteine proteases for the detection of MALT1 activity. The sections were stained with the following reagents: αIgD, αCD45.1, αCD45.2, αKi67 (clone S01A15) (eBioscience), peanut agglutinin (Vector Labs), and the MALT1 activity probe Cy5-LVSR-AOMK (23). Imaging was acquired on a Zeiss Axio ImagerM1 microscope using the Slidebook software (Intelligent Imaging Innovations). GNU Image Manipulation Program was used for overlaying images.

CD43-depleted naive B cells were cultured in the presence of 40LB cells (24), in complete RPMI 1640 supplemented with recombinant mouse IL-4 (eBioscience). On day 5, the cells were harvested and analyzed by flow cytometry.

CD43-depleted splenic B cells were rested for at least 60 min and stimulated with 1 μg/ml αIgM F(ab′)₂ for the indicated times at 37°C. Total RNA was isolated from B cells using Trizol (Thermo Fisher Scientific) and cDNA was prepared using the MMLV Reverse Transcriptase cDNA Advantage kit (Clontech), according to the manufacturers’ instructions. Quantitative real-time PCR was performed with the SsoAdvanced SYBR Green Supermix (Bio-Rad) on a CFX384 Touch Real-Time PCR Detection system (Bio-Rad). The primers were designed with the help of PrimerBank (25) and the sequences are: Bcl2 forward 5′-ATGCCTTTGTGGAACTATATGGC-3′, Bcl2 reverse 5′-GGTATGCACCCAGAGTGATGC-3′, Bcl2l1 forward 5′-GACAAGGAGATGCAGGTATTGG-3′, Bcl2l1 reverse 5′-TCCCGTAGAGATCCACAAAAGT-3′, Bcl2ala forward 5′-GGCTGAGCACTACCTTCAGTA-3′, Bcl2a1a reverse 5′-TGGCGGTATCTATGGATTCCAC-3′, Rel forward 5′-AGACTGCGACCTCAATGTGG-3′, Rel reverse 5′-GCACGGTTGTCATAAATTGGGTT-3′, Irf4 forward 5′-TCCGACAGTGGTTGATCGAC-3′, Irf4 reverse 5′-CCTCACGATTGTAGTCCTGCTT-3′, E2f3 forward 5′-AAACGCGGTATGATACGTCCC-3′, E2f3 reverse 5′-CCATCAGGAGACTGGCTCAG-3′, Gapdh forward 5′-CATGGCCTTCCGTGTTCCTA-3′, Gapdh reverse 5′-CCTGCTTCACCACCTTCTTGAT-3′. The expression level of each target gene was normalized to that of Gapdh, and the changes were calculated using the ∆∆ cycle threshold method.

Freshly isolated B cells were cultured in complete RPMI 1640 with or without stimuli. After 24 h, cells were collected and lysed with RIPA buffer (PBS, 1% NP-40, 0.5% deoxycholate, 0.1% SDS, and 10 mM EDTA) plus 10 mM sodium fluoride and complete protease inhibitor mixture (Roche). The protein content of cleared lysates was measured using the BCA Protein Assay kit (Thermo Fisher Scientific). Lysates were resolved on 4–12% or 10% polyacrylamide Bis-Tris gel (Bio-Rad or Invitrogen) and transferred onto polyvinylidene difluoride membrane (EMD Millipore). The membrane was probed for the indicated proteins. The following Abs were purchased from Cell Signaling: anti-MALT1, -Bcl-2 (D17C4), -β-actin (13E5), -phospho-Rb (Ser807/811) (D20B12), -cyclin D3 (DCS22), -c-Myc (D84C12), -survivin (71G4B7), and -total Akt. Anti-Mcl-1 was purchased from Rockland Immunochemicals, anti-cyclin D2 (M-20) from Santa Cruz, and anti-Bcl-xL from BD Biosciences.

Primary Abs were then detected with HRP-labeled donkey anti-rabbit or anti-mouse Abs (Jackson ImmunoResearch) and developed with the SuperSignal West Pico chemiluminescence kit (Thermo Fisher Scientific).

Bar graphs represent the mean values and the error bars represent the SD. Statistical significance was calculated using two-tailed unpaired Student t test or one-way ANOVA using GraphPad Prism (GraphPad Software). Only significant differences are indicated in the figures by asterisks as follows: *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.

Although the importance of MALT1 for GC formation and subsequent Ab production is undisputed (5, 6, 15), it remains unclear whether the defects associated with MALT1 deficiency arise from perturbed B cell or T cell activation, or a combination thereof. To resolve this issue, we generated chimeric animals by reconstituting sublethally irradiated B cell-deficient μMT mice, with BM from control or Malt1^−/− mice. The recipients bearing B cells that were MALT1 sufficient (μMT/WT) or deficient (μMT/knockout [KO]) were immunized with SRBCs to assess whether they can mount an effective immune response. At 7 d postimmunization, the presence of GC B cells in both the spleens and Peyer’s patches was assessed by flow cytometry. Similar to the defects reported for Malt1^−/− mice, μMT/KO animals exhibited a ∼7-fold reduction in B220⁺GL7⁺Fas⁺ GC B cells (Fig. 1A). Histology of spleen sections from the immunized μMT/KO chimera also revealed almost no PNA⁺ GC cells in the follicles, despite the presence of organized CD35⁺ FO dendritic cell networks (Fig. 1B). The reductions in the total splenic B cell pool and MZ B compartment were also recapitulated in μMT/KO mice (Fig. 1C). Interestingly, μMT/KO mice displayed a modest but significant reduction in FO B cells, which we did not observe in Malt1^−/− mice (data not shown). The defect in GC reactions in μMT/KO animals also correlated with abrogated production of SRBC-specific Abs (Fig. 1D). These findings indicate that B cells intrinsically require MALT1 for both the GC response and differentiation into MZ B cells.

One explanation for the lack of GCs in μMT mice reconstituted with Malt1^−/− BM is a failure of B cells to induce or respond to T cell help. To address this possibility, we adoptively transferred a 1:1 mixture of WT (CD45.1) to Malt1^−/− (CD45.2) splenic B cells into μMT recipients, followed by SRBC immunization. This model also enabled us to directly compare the competitive fates of Ag-activated control versus Malt1^−/− B cells in an identical environment. The WT and Malt1^−/− B cells can be tracked by flow cytometry using Abs specific for CD45.1 or CD45.2, respectively (Fig. 2A). Following immunization with TD Ags SRBCs, B cells derived from the CD45.1 donor accounted for more than 80% of the splenic GC compartment in the recipients (Fig. 2B), whereas the distribution of CD45.1⁺ and CD45.2⁺ populations within the total B cells remained equal.

The adoptive transfer approach yielded only a small number of donor-derived B cells, rendering further characterization difficult. Therefore, we also employed a congenic mixed chimera approach by reconstituting lethally irradiated μMT mice with a 1:1 mixture of CD45.1 to Malt1^−/− BM, yielding μMT/mixed animals. Histologic analysis of spleen sections from the SRBC-immunized μMT/mixed chimeras was in agreement with what we have observed with the adoptive transfer model, revealing a predominance of CD45.1⁺ cells in the GC region defined by peanut agglutinin (PNA) positivity (Supplemental Fig. 1A, 1B). We also included the Ki67 marker to detect cycling B cells and noted that its expression mainly colocalized with the CD45.1-stained cells within the GC (Supplemental Fig. 1B).

In addition to its scaffolding role in promoting NF-κB activation, MALT1 also possesses a functional protease domain that shows activity toward a number of substrates and fine-tunes NF-κB signaling through substrate cleavage (26). Using a newly developed probe that can selectively label active MALT1 (23), we were able to monitor MALT1 protease activity in situ in the spleens of the immunized μMT/mixed chimeric animals. Interestingly, cells exhibiting MALT1 activity were mostly confined to the GC region and did not appear in the FO mantle composed primarily of naive B cells (Fig. 2C, Supplemental Fig. 1C). In addition, MALT1 activity mostly colocalized with the CD45.1⁺ cells, thus demonstrating the specificity of the probe (Fig. 2C). Taken together, our data suggest that the B cell-intrinsic GC defect associated with MALT1 deficiency cannot be compensated for even in the presence of WT B cells that provide transactivating signals to cognate T cells.

The reduction in GC B cells observed at the peak of the response could have resulted from either reduced entry of B cells into the GC or a failure of the early GC cells to properly expand and differentiate. To distinguish between these possibilities, we used an analytic approach that we had previously developed (27) involving immunization with SRBCs, which induces a robust TD response with well-defined kinetics, and examination of the progression of GC B cell differentiation in the μMT/mixed chimera setting. First, PNA was used as a marker to encompass all stages of GC B cell development. After immunization, the proportion of PNA⁺ B cells that were of Malt1^−/− (CD45.2) origin was diminished (Fig. 2D, upper panel). We next incorporated GL7 and IgD staining to resolve GC B cell maturation, which proceeds from an IgD⁺GL7⁻ to an IgD⁻GL7⁺ stage (27). Focusing on the CD45.1⁺ population, there was an expansion of IgD⁻GL7⁺ GC B cells, accompanied by a decrease in IgD⁺GL7⁻ early GC B cells. In contrast, the majority of the PNA⁺ B cells within the CD45.2 gate retained the IgD⁺GL7⁻ phenotype, and <10% of the cells matured into IgD⁻GL7⁺ GC B cells (Fig. 2E). Collectively, these results suggest that GC initiation is defective in Malt1^−/− animals and that the few PNA⁺IgD⁺GL7⁻ early GC B cells that are present also fail to mature and expand. The defect in GC initiation paralleled the reduced frequency of Malt1^−/−-derived B220loCD138⁺ plasma cells on days 7 and 10 (Fig. 2F). Interestingly, we also observed a decreased T_FH population of Malt1^−/− origin on day 7, suggesting that MALT1-deficient T cells harbor an intrinsic block in T_FH differentiation (Fig. 2G).

The inability to mount a GC response translates into a failure to generate memory B cells and long-lived plasma cells. To determine how extensive the defect in GC formation is, we adopted an in vitro culture system that has been previously described (24) to propagate GC phenotype B cells. When naive WT B cells were cultured on the 40LB feeder cells, which are BALB/c 3T3 fibroblasts stably transfected with CD40L and BAFF, they underwent massive expansion and acquired the GL7⁺FAS⁺ GC phenotype, and are termed induced GC B (iGB) cells (Fig. 3, left panels). Supplementing the culture with IL-4 also induced BCR class switching from IgM to IgG1 (Fig. 3, middle panels). Strikingly, we were able to efficiently induce the Malt1^−/− B cells to differentiate into GL7⁺FAS⁺ GC-like B cells after 5 d in culture with 40LB cells. Similar to the iGB cells derived from WT B cells, ∼50% of the Malt1^−/− iGB cells expressed IgG1, indicating that the class-switching machinery in MALT1-deficient B cells remains intact. Although a small fraction of the Malt1^−/− iGB cells differentiated into B220loCD138⁺ plasmablasts, we noted that this population was significantly reduced relative to WT iGB cells (Fig. 3, right panels). Thus, Malt1^−/− FO B cells proliferate normally to T cell-derived stimuli to adopt a GC B cell phenotype, but are impaired in maturation toward the plasma cell lineage.

The role of MALT1 in B cell proliferation in response to BCR engagement remains unclear due to conflicting results in the literature (5, 6). We observed that FO (MZ B cell depleted) Malt1^−/− B cells proliferated robustly, comparable to WT B cells, in response to LPS or αCD40 plus IL-4, but were hyporesponsive to αIgM stimuli (Supplemental Fig. 2). To investigate the nature of this proliferative defect, we performed cell cycle analysis using BrdU incorporation in combination with the DNA dye 7-AAD. We observed a significant reduction in S-phase entry of Malt1^−/− B cells at 24 h after αIgM treatment. However, by 48 h poststimulation, the percentage of Malt1^−/− B cells in the S as well as G2/M phase increased to levels comparable to WT B cells (Fig. 4A). Normal B cells induce expression of both cyclins D2 and D3 upon stimulation with αIgM (28). The two d-type cyclins activate the G1 kinases and target the retinoblastoma gene product for phosphorylation, thus freeing the E2F protein to drive transcription of genes required for the transition from the G1 to S phase. In line with the cell cycle analysis, we observed upregulation of cyclins D2 and D3, and subsequent induction of phospho-Rb in both WT and Malt1^−/− B cells upon αIgM treatment (Fig. 4B). Taken together, these results suggest there is a delay rather than lack of proliferative response of Malt1^−/− B cells to BCR crosslinking.

The finding that the proliferation machinery remains functional in Malt1^−/− B cells raised the possibility that most Malt1^−/− B cells undergo apoptosis before they have a chance to divide, although it is unclear whether this occurs preferentially during the S or G2/M phase of the cell cycle. This notion prompted us to examine the viability of MALT1-deficient B cells in response to a panel of stimuli. There was a reduction in viability when Malt1^−/− B cells were cultured in media alone or stimulated with αIgM (Fig. 4C). Although the addition of BAFF or αCD40 to αIgM treatment was able to enhance the viability of WT B cells, it failed to achieve the same effect on Malt1^−/− B cells. We previously noted that LPS- or αCD40-induced proliferation remained intact in Malt1^−/− B cells and, consistently, Malt1^−/− B cells survived just as well as WT B cells when stimulated with either factor, suggesting that both the proliferative and survival defects are restricted to signaling through the BCR. Although the addition of BAFF failed to increase the viability of αIgM-stimulated Malt1^−/− B cells compared with cells in media, BAFF treatment alone improved the survival of Malt1^−/− B cells. Altogether, these findings raise the intriguing possibility that in the absence of MALT1, BCR engagement induces a dominant proapoptotic signal.

To examine more closely the interplay of proliferation and survival, we used combined proliferation dye and Annexin-V labeling to track cell division and apoptosis induced by αIgM treatment. We noted that in the case of the Malt1^−/− B cells, the majority of the Annexin-V⁺ apoptotic cells did not dilute the proliferation dye, suggesting that the cells underwent apoptosis before cell division took place (Fig. 4D). In line with a recent report (15), the addition of IL-4, but not BAFF, was able to induce proliferation of Malt1^−/− B cells and improve their viability. LPS alone was found to provide strong mitogenic signals to both WT and Malt1^−/− B cells, and we did not observe any significant difference in proliferation and survival when LPS was administered in combination with αIgM. In contrast, WT and Malt1^−/− B cells responded differently to combined αIgM and αCD40 treatment. Although the addition of αCD40 was able to rescue BCR-induced proliferation in Malt1^−/− B cells to some extent, there were fewer rounds of division compared with WT B cells stimulated under the same conditions, and there were far fewer viable Annexin-V Malt1^−/− B cells. This result is consistent with a dominant proapoptotic effect of BCR signaling in the absence of MALT1.

Next, we sought to determine if rescuing survival can overcome the impairment in proliferative response to αIgM stimulation. When treated with the broad spectrum caspase inhibitor IDN-6556, the viability of Malt1^−/− B cells in media alone or stimulated with αIgM was increased to a level comparable with that of WT B cells (Fig. 4E). Moreover, in the presence of IDN-6556, Malt1^−/− B cells stimulated with αIgM exhibited a proliferative profile similar to WT B cells (Fig. 4F). Collectively, our findings indicate that the delay in cell cycle entry and the increased propensity to caspase-dependent apoptosis account for the impaired proliferative response of Malt1^−/− B cells to BCR stimulation.

To rule out the possibility that Malt1^−/− B cells are simply anergic to BCR crosslinking, the levels of various activation markers on the cell surface were assessed by flow cytometry. When stimulated with αIgM, Malt1^−/− B cells upregulated CD5, CD86, MHC II, CD69, and CD80 to levels comparable with WT B cells (Fig. 5A). Notable differences were observed in the induction of CD25 and downregulation of CD62L on WT B cells and, surprisingly, enhanced basal expression of MHC II on Malt1^−/− B cells. During TD immune responses, Ag-activated B cells migrate to the B–T zone boundary to seek help from cognate T cells. Accessory signals, notably CD40 and CD40L interactions, are necessary to drive the initial B cell proliferation and induction of GC formation. Profiling of activation markers on BCR-activated Malt1^−/− B cells suggests that they retain the ability to prime T cells. Although we found that the combination of αIgM and αCD40 stimuli were able to partially restore αIgM-induced proliferation in Malt1^−/− B cells (Fig. 4D), Malt1^−/− B cells did not survive well under the combined stimulation. These findings prompted us to examine the fate of Malt1^−/− B cell upon receiving T cell help. We mimicked the events in the GC response by pretreating B cells with αIgM, followed by αCD40 stimulation. It has been shown that washing out αIgM after 24 h did not prevent the first division, but most of the cells died instead of completing further divisions (29). However, subsequent engagement of CD40 was sufficient to induce further proliferation of WT B cells and rescue cell death (Fig. 5B, 5C). In contrast, Malt1^−/− B cells failed to proliferate and were more prone to apoptosis, verifying that CD40 signals delivered by the T cells cannot correct the proliferation and survival defects in BCR-activated Malt1^−/− B cells. Interestingly, Malt1^−/− B cells primed with αCD40 were able to complete one round of cell division upon subsequent αIgM stimulation, but their viability was also markedly reduced compared with that of WT B cells. In the presence of IL-4, Malt1^−/− B cells underwent robust proliferation regardless of the stimulating conditions (Fig. 5B). Thus, the impairment in both proliferative and survival response to signaling through BCR and CD40 may explain why GC initiation is defective in Malt1^−/− B cells.

PKCβ is situated upstream of the CBM complex and controls assembly of the complex via phosphorylation of CARMA1 upon BCR engagement (30). PKCβ-deficient animals share many of the defects observed in Malt1^−/− mice. In particular, BCR-dependent proliferation and survival in Pkcb^−/− B cells are impaired due to defective expression of Bcl-2 and Bcl-xL (31, 32). Thus, we examined the levels of various anti-apoptotic proteins in BCR-activated WT and Malt1^−/− B cells. Although WT B cells upregulated Bcl-xL when cultured in the presence of αIgM alone or in combination with BAFF or αCD40, Bcl-xL expression was reduced in Malt1^−/− B cells after stimulation (Fig. 6A). Intriguingly, whereas Bcl-2 is abundant in unstimulated B cells there was a reduction in Bcl-2 expression in Malt1^−/− B cells upon stimulation. In contrast, the Mcl-1 expression was comparable between WT and Malt1^−/− B cells. Malt1^−/− B cells also induced c-Myc expression upon stimulation, indicating that they were responsive to stimulation. Bcl2l1 is a direct target of the transcription factor c-Rel (33), which was found to be selectively activated by MALT1 (8). This prompted us to monitor the transcript levels of several known c-Rel target genes in WT versus Malt1^−/− B cells after αIgM treatment by time-course real-time PCR. With the exception of Bcl2l1, the profiles of all the other genes in response to αIgM stimulation were similar between WT and Malt1^−/− B cells (Fig. 6B). Thus, the reduction in Bcl-2 is likely regulated at the level of protein stability. These data suggest that the impaired survival in BCR-activated Malt1^−/− B cells is due to reduced transcription of Bcl2l1 and impaired stability of Bcl-2.

Given that treatment with a caspase inhibitor was able to restore proliferation and survival of BCR-activated Malt1^−/− B cells, we sought to determine whether ectopic expression of Bcl-2 could rescue MZ B cell development and TD responses in Malt1^−/− mice. Malt1^−/− mice were interbred with Eμ-Bcl-2-22 transgenic mice, which specifically express the human Bcl2 transgene in the B lineage (34), and chimeric mice were generated by transferring BM from Bcl-2 Tg × Malt1^+/− (μMT/Bcl-2 Tg × Malt1^+/−) or Bcl-2 Tg × Malt1^−/− (μMT/Bcl-2 Tg × Malt1^−/−) littermates into sublethally irradiated μMT mice. Remarkably, immunization with SRBC elicited GC responses in μMT mice reconstituted with Bcl-2 Tg × Malt1^−/− BM, albeit not as robust as in μMT/Bcl-2 Tg × Malt1^+/− (Fig. 7A, top panels). The μMT/Bcl-2 Tg × Malt1^−/− recipients reconstituted 34% of the GL7⁺FAS⁺ cells as a proportion of total B220⁺ B cells relative to μMT/Bcl-2 Tg × Malt1^+/− mice. A similar proportion of the GC B cells in both groups stained positive for IgG1 (Fig. 7A, middle panels), suggesting that MALT1 is not essential for class-switch recombination. However, the frequency of plasma cells was drastically reduced in μMT/Bcl-2 Tg × Malt1^−/− animals (Fig. 7A, lower panels). In addition, we found that forced Bcl-2 expression did not rescue the defect in MZ B cell differentiation observed in Malt1^−/− animals (Fig. 7B). Although the two groups showed comparable frequencies of splenic CD4⁺ T cells, μMT/Bcl-2 Tg × Malt1^−/− animals exhibited a 2-fold reduction in T_FH population (Fig. 7C), which correlated with the reduction in GC B cells.

To investigate whether the reduction in GC B cells observed at the peak of GC response was due to impaired maturation of early GC founder cells, PNA, GL7, and IgD staining was incorporated to resolve subsets of GC B cells in the immunized μMT chimeras. The entire GC compartment was first identified by staining with the pan-GC marker PNA, which showed a 3-fold reduction in μMT/Bcl-2 Tg × Malt1^−/− animals (Fig. 7D, upper panels). However, there was no appreciable difference in the proportions of mature IgD⁻GL7⁺ and early IgD⁺GL7⁻ GC B cells in μMT/Bcl-2 Tg × Malt1^−/− relative to μMT/Bcl-2 Tg × Malt1^+/− mice (Fig. 7D, middle and bottom panels). When histology was performed on μMT/Bcl-2 Tg × Malt1^−/− spleen cross-sections, we were able to detect PNA⁺ GC clusters in all the follicles, but the size of the GC clusters was smaller compared with those seen in the μMT/Bcl-2 Tg × Malt1^+/− animals (Fig. 7E). The effect on the Ab response was also assessed, and as expected, SRBC immunization led to the production of SRBC-specific IgM and IgG Abs in μMT/Bcl-2 Tg × Malt1^+/− animals. Consistent with the block in plasma cell differentiation, the levels of SRBC-specific Abs were severely diminished in immunized μMT/Bcl-2 Tg × Malt1^−/− chimeras (Fig. 7F). Taken together, in the presence of WT T cells, forced expression of Bcl-2 in B cells partially restored GC formation in MALT1-deficient animals, but failed to rescue plasma cell differentiation or MZ B cell formation.

To investigate whether the Bcl2 transgene rescues the response to BCR stimulation, proliferation and apoptosis of splenic B cells were monitored by combined Annexin-V and eFluor670 labeling. Similar to WT B cells, Bcl-2 Tg × Malt1^+/− B cells underwent several rounds of division after 72 h in culture in response to BCR triggering, and proliferation was further enhanced by the addition or BAFF or αCD40 (Fig. 8A). Notably, Bcl-2 Tg × Malt1^−/− B cells cycled at most two rounds to BCR ligation, although viability was increased ∼6-fold by ectopic Bcl-2 expression compared with Malt1^−/− B cells (see Fig. 4D). Although the addition of BAFF or αCD40 had a minimal effect on αIgM-induced proliferation in Bcl-2 Tg × Malt1^−/− B cells, the inclusion of either IL-4 or LPS triggered a robust proliferative profile that was similar to what Bcl-2 Tg × Malt1^+/− B cells exhibited. The survival of Bcl-2 Tg × Malt1^+/− and Bcl-2 Tg × Malt1^−/− B cells in response to other stimuli was also explored, and the incubation period was extended from 72 to 96 h to determine if there was a greater differential. Viability was similar for Bcl-2 Tg × Malt1^+/− and Bcl-2 Tg × Malt1^−/− B cells when cultured under resting conditions or in the presence of stimuli other than αIgM (Fig. 8B). In contrast, despite the enforced Bcl-2 expression, Bcl-2 Tg × Malt1^−/− B cells exhibited more cell death when stimulated with αIgM, and the addition of BAFF or αCD40 failed to enhance viability. Lastly, we examined the differentiation of plasma cells under defined in vitro conditions to rule out other B cell extrinsic variables. In support of our in vivo findings, the generation of B220loCD138⁺ Ab-secreting cells was significantly reduced in Bcl-2 Tg × Malt1^−/− and Malt1^−/− B cells after 3 d of culture with LPS or αCD40⁺IL-4, compared with their respective littermate controls (Fig. 8C, 8D). Our collective results thus suggest that MALT1 is selectively required for the survival of resting and BCR-activated B cells, and that forced expression of Bcl-2 can partially overcome this proapoptotic effect but it does not enable the differentiation of Ab-secreting cells.

MALT1 has been recognized as an essential component of the Ag receptor signaling pathway, regulating NF-κB activity via both scaffolding (35) and protease functions (36–39). However, its specific role in B cells has not been fully characterized. In this study, we provide several lines of evidence for a cell-intrinsic role of MALT1 in regulating B cell immune function. Using chimeric mouse models, we demonstrated that there is a B cell-intrinsic requirement for MALT1 in the GC response to TD Ags. Central to BCR-induced NF-κB signaling is Bruton’s tyrosine kinase (Btk), which is primarily responsible for the activation of PLCγ2 and subsequently PKCβ, leading to the assembly of the CBM complex. Mutation or inhibition of Btk has been shown to result in a compromised GC response (40, 41). In accordance with the BCR-Btk pathway, deficiency in any of the components of the CBM complex led to impaired TD humoral response (5, 6, 31, 42–44). It is thought that NF-κB is dispensable for GC B cell proliferation, because GC B cells fail to express most NF-κB target genes and NF-κB pathway components (45). This notion is supported by a later study showing that BCR signaling is dampened in GC B cells, due to phosphatase activity, and increases again in the light zone (46). A recent study dissecting the differential roles of the NF-κB subunits RelA and c-Rel in the GC response also revealed that although c-Rel is required for GC maintenance, activation of RelA is indispensable at later stages for the generation of plasma cells (14). Thus, we postulate that BCR-induced NF-κB induction is more associated with GC initiation as a consequence of Ag activation. Consistently, our results indicate that the block in Malt1^−/− animals occurs at the early stage of the GC reaction before proliferative expansion takes place. Interestingly, using an activity-based probe, we showed that MALT1 proteolytic activity can only be detected within the GC region. A recent report on MALT1 protease-inactive mice revealed that although PMA-induced NF-κB activation remains intact in B cells, the lack of MALT1 proteolytic activity partially affected the GC response (15). Whether MALT1-mediated cleavage events are necessary for the maintenance of GC reactions and GC exit would require the characterization of additional inducible deletion mouse models. Further studies are also needed to elucidate the mechanism of how MALT1 protease activity is regulated during the GC response and the in vivo consequences of MALT1 substrate cleavage. In this sense, it would be interesting to determine whether MALT1 protease activity mirrors that of NF-κB signaling.

Importantly, our data demonstrate that the requirement for MALT1 is mainly restricted to BCR-induced signaling. Divergent views on the involvement of MALT1 in mitogen-induced proliferation exist since the first Malt1^−/− mice were described. Whereas Ruefli-Brasse et al. (5) reported that proliferative responses to αIgM, αCD40, or LPS were all impaired in Malt1^−/− B cells, Ruland et al. (6) argued that MALT1 is dispensable for proliferation induced by any of the three stimuli. Our results, using eFluor670 dye dilution to directly monitor cell division instead of ³H-thymidine incorporation to measure DNA replication, are intermediate and more in line with observations made by Ferch et al. (8). Also consistent with recent findings reported by Bornancin et al. (15), we showed that costimulation of IL-4 rectified both the proliferative and survival defects in BCR-activated Malt1^−/− B cells, most likely via induction of the STAT6 pathway (47). A similar defect in BCR-induced proliferation was seen in mice deficient in CARMA-1, BCL-10, Btk, and PKCβ, thus supporting the requirement of the CBM complex downstream of the BCR-Btk axis (43, 48–50). Although CD40 signaling remains intact in Malt1^−/− B cells, when αCD40 treatment was provided at the same time, or preceding BCR stimulation, it had a minimal effect on survival and only partially rescued proliferation. Costimulation with BAFF also shared a similar proliferation/survival profile as stimulation of BCR alone, suggesting that the defect in BCR-induced signaling in the absence of MALT1 is dominant over BAFF-induced signaling.

Our results indicate that reduced viability of BCR-activated Malt1^−/− B cells is due to reduced expression of Bcl-2 and lack of Bcl-xL upregulation, on both mRNA and protein levels. Interestingly, there was no difference in viability between Bcl-2 Tg × Malt1^−/− and Bcl-2 Tg × Malt1^+/− B cells that were left in media alone, but viability of Bcl-2 Tg × Malt1^−/− B cells was reduced in all stimulating conditions involving αIgM stimulation, suggesting that the protective mechanism of MALT1 following BCR activation also includes a Bcl-2 independent component. Although overexpression of either Bcl-2 or Bcl-xL leads to accumulation of mature B cells, Bcl-xL but not Bcl-2 becomes rapidly upregulated upon activation via surface Ig, LPS, or CD40 stimulation (51, 52). Furthermore, within the GC, CD40-CD40L interactions rescue centrocytes from apoptosis primarily via the induction of Bcl-xL (53), which can also be mimicked by the ectopic expression of Bcl-2 (54). More definitive experiments are required to verify whether Bcl-xL expression in B cells can completely reverse the defects associated with MALT1 deficiency. How MALT1 regulates the expression of both Bcl-2 and Bcl-xL also awaits further analysis. Previous studies have identified c-Rel as a protector of B cells from BCR-mediated apoptosis, partly via induction of Bcl-xL and A1 (13, 33). In contrast, the link between Bcl-2 expression and NF-κB activation in B cells is not firmly established.

Our results indicate that the Bcl2 transgene, together with the presence of WT T cells, led to a partial rescue of GC formation by prolonging the survival of MALT1-deficient B cells. Our μMT/mixed chimera model showed that even in the same environment as WT T cells, MALT1-deficient T cells less readily differentiate into the T_FH phenotype. T_FH cells have been acknowledged as the key cell type required for GC formation and the generation of memory B cells (55). Although T_FH cells provide help to Ag-activated B cells, reciprocal signals from B cells are also important for the maintenance of T_FH differentiation, leading to the induction of the master regulator transcription factor Bcl-6 in either cell type (56, 57). Consequently, the reduction in T_FH phenotype in Malt1^−/− mice may reflect the inability of MALT1-deficient B cells to successfully prime T_FH differentiation. It has been reported that sustained Ag presentation by dendritic cells facilitates early T_FH formation in the absence of B cells (58). These findings suggest that the block in T_FH differentiation may also arise from a cell-intrinsic defect in Malt1^−/− T cells. The MALT1 substrate Roquin is a likely candidate because it inhibits ICOS, and the loss of both Roquin-1 and Roquin-2 has been reported to result in spontaneous T_FH differentiation and GC development (59, 60).

Nevertheless, there remains a B cell-intrinsic requirement for MALT1 for the GC response, plasma cell differentiation, and Ig production. The partial decline in GC B cells observed in the μMT/Bcl-2 × Malt1^−/− animals can be explained by the intrinsic block in BCR-induced proliferation associated with MALT1 deficiency, thereby restricting the expansion of centroblasts during early GC development. Although defective c-Rel translocation has been implicated in Malt1^−/− B cells (8), we cannot completely rule out the involvement of MALT1 in regulating activation of RelA as well, provided that RelA is essential for the generation of GC-derived plasma cells (14). Our results indicate that MALT1 is also indispensable for plasmablast differentiation induced by BCR-independent stimuli. The fact that LPS- and αCD40⁺IL-4–induced proliferation remains intact in Malt1^−/− B cells seems paradoxical, but our analysis of in vitro B cell culture revealed an impairment in plasmablast generation. Given that differentiation into plasma cells is a stochastic event that is tightly associated with cell divisions and is governed by a distinct gene regulatory network (61, 62), MALT1 may promote plasma cell differentiation by either regulating cell cycle progression or the induction of transcription factors including Blimp-1, XBP-1, and Irf4. These functions of MALT1 are likely independent of its scaffolding role as part of the CBM signalosome during BCR stimulation.

Humoral responses to TI Ags were also found to be defective in Malt1^−/− animals (5, 6), most likely due to the compromised B1 and MZ compartment and BCR-induced apoptosis of mature recirculating B cells (63). It has been shown that MZ B cells proliferate better than FO B cells in response to LPS stimulation, presumably because the MZ B cells express higher level of TLR4 (64, 65). Moreover, MZ B cells are essential for the initiation of TI immune response by differentiating into plasmablasts upon interaction with other APCs (66). Intriguingly, unlike mice with mutations in components of the BCR-Btk pathway that exhibit a loss of FO B cells but normal MZ structure (67, 68), deletion of any of the CBM complex components results in a substantial decrease in MZ B cells (44, 69). Based on our findings, it appears that the reduction in MZ B cells is B cell autonomous and is not due to impaired survival, because enforced Bcl-2 expression failed to rescue the MZ compartment. Although canonical NF-κB signaling plays a role in lymphocyte development, it has been shown that c-Rel or RelA deficiency only partially affects MZ B cell development (70). In contrast, MZ B cell development is dependent on BAFF-induced signaling, and this requirement cannot be overcome by constitutive canonical NF-κB signaling (71).

MALT1 protease activity has been implicated in both activated B cell-like diffuse large B cell lymphomas (ABC-DLBCL) and MALT lymphomas. In ABC-DLBCL cells, which are characterized by a preassembled CBM complex and constitutive NF-κB activation, targeting of the MALT1 caspase-like domain with the use of small molecule inhibitors or peptides has proven effective in inducing apoptosis (72–75). A subset of MALT lymphoma patients harbor the t (11, 18) (q21;q21) translocation, which results in the fusion of the Birc3 (Ciap2) and Malt1 genes. The resulting product, cIAP2-MALT1, is a potent activator of NF-κB, due to spontaneous oligomerization through its N-terminal cIAP2-derived sequence with the MALT1 C terminus (76). Strikingly, the MALT1 portion of the fusion protein retains functionality and it has been shown that cIAP2-MALT1 contributes to the pathogenesis of MALT lymphoma by recruiting and cleaving substrates including NF-κB inducing kinase and LIM domain and actin-binding protein 1, which do not normally interact with the WT MALT1 protease (77, 78). Consequently, therapeutic inhibition of MALT1 proteolytic activity has emerged as a promising approach for the treatment of B cell lymphomas with deregulated NF-κB activation. Furthermore, MALT1 inhibition has advantages over targeting of upstream protein kinases in ABC-DLBCL tumors with lesions that are located downstream or in parallel pathways, such as CARMA1 or MYD88, respectively, because they are unlikely to respond to inhibition of Syk, Btk, or PKCβ.

The authors have no financial conflicts of interest.