|Home | About | Journals | Submit | Contact Us | Français|
BCR signaling plays a critical role in B-cell tolerance and activation. Here, we show that mice with B cell-specific ablation of both c-Cbl and Cbl-b (Cbl-dko) manifest systemic lupus erythymatosis (SLE)-like autoimmune disease. The Cbl-dko mutation results in a significant increase in marginal zone (MZ) and B1 B cells. The mutant B cells are not hyperresponsive in terms of proliferation and antibody production upon BCR stimulation; however, B-cell anergy to protein antigen appears to be impaired. Concomitantly, BCR-proximal signaling, including tyrosine phosphorylation of Syk, PLCγ-2, and Vav and Ca2+ mobilization, are enhanced, whereas tyrosine phosphorylation of BLNK is significantly attenuated in the mutant B cells, suggesting that the loss of coordination between these pathways is responsible for the impaired B-cell tolerance induction. Thus, Cbl proteins control B cell-intrinsic checkpoint of immune tolerance, possibly through coordinating multiple BCR-proximal signaling pathways during anergy induction.
B-cell development, activation, and tolerance are interconnected processes controlled by signals delivered by the B-cell antigen receptor (BCR) (Healy and Goodnow, 1998; Rajewsky, 1996; Reth and Wienands, 1997). Paradoxically, the same BCR can either signal immunogenically, stimulating the proliferation and differentiation of B cells specific for foreign antigens, or signal tolerogenically to eliminate or silence cells that bind to self-antigens. Although divergent hypotheses exist as to how precisely BCR signaling is triggered by antigen and how this signaling is quantitatively and differentially altered in tolerized B cells (Healy et al., 1997; Vilen et al., 2002), the developmental timing when B cells encounter antigens may determine the final outcomes (Cancro, 2004; Chung et al., 2003). In particular, evidence indicate that triggering of the antigen receptors on bone marrow (BM) immature and peripheral transitional (T1 or T2) B cells leads to B-cell tolerance in the absence of T-cell help (Allman et al., 1992; Carsetti et al., 1995; Fulcher and Basten, 1994). These findings thus support the idea that the immature stages of B-cell development may represent a time window during which B-cell tolerance is established. After these stages, binding of antigens to the BCR on mature B cells results in B-cell activation.
The BCR complex is composed of antigen binding chains, the Ig molecules and a non-covalently associated signal transduction complex, Ig-α/Ig-β, containing in its cytoplasmic domain immunoreceptor tyrosine-based activation motifs (ITAMs) (Cambier, 1995b; Campbell, 1999; Reth, 1989; Reth, 1992). Cross-linking of the BCR results in tyrosine phosphorylation of the ITAMs by Src family tyrosine kinase Lyn followed by recruitment and activation of Syk tyrosine kinase (Cambier, 1995a; Reth and Wienands, 1997). Recruitment and activation of Syk by the phosphorylated BCR is a key event in the assembly of the BCR signalosome composed of the adaptor protein BLNK and downstream signaling components PLCγ-2, Bruton’s tyrosine kinase (Btk) and Vav (Kurosaki, 2002; Pierce, 2002). These components coordinately induce Ca2+-influx and activate nuclear signals, including NF-AT, AP-1, and NF-κB that are essential for B-cell development and activation (Campbell, 1999; Kurosaki, 2000).
Cbl proteins were recently identified as E3 ubiquitin ligase (Joazeiro et al., 1999). They interact with E2-ubiquitin conjugating enzyme (Ubc) through their ring figure (RF) domain, and regulate the signaling of a broad range of receptors by promoting ubiquitination of the components involved in these receptor signaling (Duan et al., 2004; Liu and Gu, 2002; Thien and Langdon, 2005). In mammals, the Cbl family of proteins has three members, c-Cbl, Cbl-b, and Cbl-3, among which c-Cbl and Cbl-b are expressed in hematopoietic cells (Duan et al., 2004). Recent genetic studies from our and several other laboratories have revealed a critical role of Cbl proteins in T-lymphocyte development and activation (Bachmaier et al., 2000; Chiang et al., 2000; Murphy et al., 1998; Naramura et al., 2002; Naramura et al., 1998). The role of Cbl in B-cell development and function requires further investigation. The involvement of Cbl proteins in BCR signaling has been reported in several papers, in which c-Cbl and Cbl-b were shown to regulate PLCγ-2 activation and Ca++ response (Sohn et al., 2003; Yasuda et al., 2000; 2002). Cbl proteins associate with Syk and BLNK upon BCR stimulation, suggesting that they are part of the BCR signalosome. Cbl-b deficiency leads to an enhanced tyrosine phosphorylation of Syk and Ca++ response in mouse B cells, despite of normal BCR-induced proliferation of Cbl-b−/− B cells (Sohn et al., 2003). However, the precise signaling and physiological function of Cbl proteins in B-cell biology has not yet been fully addressed, to some extent as a result of functional redundancy between c-Cbl and Cbl-b.
In order to understand the biochemical and physiological functions of Cbl proteins in B cells, we have generated a mouse model in which c-Cbl and Cbl-b are simultaneously inactivated in B-lineage cells. Our study revealed that these mice manifested systemic lupus erythematosus (SLE)-like disease. The mutation significantly increased the rate of B-cell maturation and impaired B-cell anergy. These results thus indicate that Cbl proteins control the checkpoint of B-cell tolerance, possibly by extending the duration of B-cell maturation, providing sufficient time for B-cell tolerance induction.
Our previous data demonstrate that inactivation of the germline c-cbl or cbl-b gene alone results in a negligible impact on the development and function of B cells; however, the simultaneous ablation of both c-cbl and cbl-b genes in germline leads to embryonic lethality (Naramura et al., 2002), suggesting that c-Cbl and Cbl-b may have a redundant role in intracellular signaling. To assess whether c-Cbl and Cbl-b have a redundant function in B cells, we generated mutant mice in which the c-cbl and cbl-b genes were simultaneously inactivated only in B cells. These mice carried the homozygous c-cblf/f (c-cbl gene flanked by loxP sequences) alleles and cbl-b−/− (cbl-b null) alleles and a CD19-cre transgene (Tg). Since deletion of the c-cblf/f alleles in a given cell by the Cre recombinase results in the Cbl-dko mutation, we expected that in these mice the Cbl-dko mutation would occur only in B cells, because CD19-cre transgene was expressed specifically in B-lineage cells (Rickert et al., 1997). Indeed, we found that in these mice the c-cblf/f alleles were deleted efficiently in B but not T cells (Supplementary Fig. 1a, 1b, and 1c and data not shown). Hereafter we will refer to c-cblf/f cbl-b−/− CD19-cre Tg mice as Cbl-dko mice.
Cbl-dko mice were born normal and fertile, and exhibited no gross abnormality in major organs (data not shown). To determine whether the Cbl-dko mutation altered B-cell development, we analyzed B-cell compartments of the bone marrow (BM), spleen, lymph nodes, and peritoneal cavity from the mutant mice by flow cytometry (Fig. 1a). Cbl-dko and WT control mice possessed comparable numbers of BM B (B220+) cells, as well as similar representation of BM B-cell subsets, including pro/pre (B220lo IgM−), immature (B220lo IgM+), and mature re-circulating (B220hi IgM+) B cells. These observations were expected as CD19-Cre-mediated deletion occurred in less than 40% of pro/pre-B cells whereas almost complete deletion was found only in mature B cells (Supplementary Fig. 1d). On the contrary, we found that the mutant mice possessed approximately 30% more B (B220+) cells than did the WT mice and altered representations of B-cell subsets in spleen and peritoneal cavity (Fig. 1a and 1c), suggesting that the Cbl-dko mutation affected peripheral B-cell development. To determine which subsets of peripheral B cells were affected by the Cbl-dko mutation, we analyzed the cellularity of splenic B-cell subsets by flow cytometry. We found that Cbl-dko mice possessed approximately 2-folds more splenic T1 (B220+ AA4.1hi HSAhi CD21lo CD23lo), B1 (B220+ AA4.1lo HSAlo CD21− CD23−), and marginal zone (MZ) (B220+ AA4.1lo HSAlo CD21hi CD23−) B cells as compared to WT mice; however, the total numbers of follicular (FO) (B220+ AA4.1lo HSAlo CD21+ CD23+) and T2 (B220+ AA4.1hi HSAhi CD21hi CD23hi) B cells were comparable between the mutant and WT mice (Fig. 1b and 1c). Cbl-dko mice also had an increased number (up to 20–40% more) of B1 B cells in the peritoneal cavity than did the WT mice (Fig. 1a and data not shown). Based on these results we conclude that the Cbl-dko mutation alters the development of multiple B-cell subsets in the periphery.
Our inspection revealed that while the WT control mice (10/10) remained normal beyond 10 months of age, 50% (6/11) of Cbl-dko mice became moribund during the same period. The mutant mice also possessed a significantly (3–5 folds) higher level of serum IgM antibodies as compared to WT, c-Cbl−/−, and Cbl-b−/− mice; however, the serum levels of the other Ig isotypes in the mutant mice appeared to be similar to that in control mice (Fig. 2a). To determine whether immune tolerance was impaired in the mutant mice, we first examined the titers of serum autoantibodies of anti-double stranded-DNA (anti-dsDNA) and presence of anti-nuclear antigen (ANA) by ELISA and immunofluorescent-staining, respectively. We found that a high percentage of Cbl-dko mice but not WT littermates possessed anti-dsDNA and ANA of both IgG (Fig. 2b, 2c) and IgM (data not shown) isotypes in the sera, suggesting that the mutant mice developed autoimmune diseases. To directly assess whether Cbl-dko mice manifested autoimmune diseases, we performed histopathological analysis on various tissues from the mutant and control mice. Comparing to the WT or Cbl-b−/− (data not shown) mice in which the tissue infiltration of leukocytes were absent, we found massive infiltrations of leukocytes in liver, lung, kidney, and salivary grand of Cbl-dko mice that have developed the diseases (Fig. 2d). Immunohistological analysis on kidney sections of the mutant mice revealed greatly enlarged glomeruli (average diameter: 76.02 ± 12.72 vs 49.12 ± 8.28 μM, (n=40 glomeruli), p < 0.001) and severe glomerular deposits of IgG and IgM antibodies than controls (Fig. 2e). These results thus suggest that the Cbl-dko mutant mice developed SLE-like autoimmune disorders. Since our Cbl-dko mice carry the Cbl-dko mutation only in B cells and Cbl-b−/− mice did not develop spontaneous autoimmune diseases (Chiang et al., 2000), we propose that the simultaneous ablation of c-Cbl and Cbl-b in B cells disrupt B cell-intrinsic program for immune tolerance induction.
Impaired B-cell tolerance is frequently associated with B-cell hyperactivity and/or resistance to apoptosis. To determine whether the Cbl-dko mutation affected B-cell activation in vivo, we examined T-dependent (TD) and T-independent (TI) antibody responses in these mice. We immunized WT and Cbl-dko mice either with hapten nitrophenyl-acetyl (NP)-conjugated keyhole lymphocyte hemaglutinin (KLH) for TD antibody responses, or with NP-Ficoll or NP-lipopolisacharade (LPS) for type-I or type-II TI responses. In NP-KLH immunized Cbl-dko mice, the levels of anti-NP specific IgM responses were comparable to that produced by WT, c-Cbl−/−, and Cbl-b−/− mice; however, the production of IgG1 and IgG2b anti-NP antibodies in the mutant mice were moderately lower than that in control mice (Fig. 3a). Similarly, while Cbl-dko mice had a comparable level of IgM anti-NP antibodies to that produced by WT, c-Cbl−/−, and Cbl-b−/−mice after NP-Ficoll immunization, they produced significantly low amount of IgG3 anti-NP than did the control mice (Fig. 3b). The levels of IgM and IgG3 anti-NP in NP-LPS immunized mutant and WT mice were comparable (Fig. 3c), indicating that the type-II TI antibody response was not affected by the Cbl-dko mutation.
To determine whether the Cbl-dko mutation influenced B-cell activation and survival in vitro, we assessed the proliferation and apoptosis of purified splenic B cells upon anti-IgM stimulation. The proliferative response of Cbl-dko B cells to anti-IgM stimulation alone was severely impaired as compared to that of WT B cells; however, the proliferation of the mutant B cells was restored to normal in the presence of IL-4 or anti-CD40 antibody (Fig. 3d). The defective proliferation of mutant B cells in response to anti-IgM stimulation was not likely caused by a different ratio of immature vs mature B cells between WT and Cbl-dko B cell compartment, because a similar defect was also found in purified mature (B220+ AA4.1lo HSAlo) B cells (Supplementary Fig. 2). It was unlikely a result of impaired B-cell activation neither, because the mutant B cells up-regulated cell-surface markers CD69, CD86, and MHC-II as efficiently as did the WT B cells upon anti-IgM stimulation (Fig. 3e). To determine whether the Cbl-dko mutation promoted BCR activation-induced cell death, we analyzed apoptosis of Cbl-dko B cells after BCR stimulation in the presence or absence of anti-CD40 antibody or IL-4 (Fig. 3f). Cbl-dko B cells exhibited a much higher rate of apoptosis than did the WT cells upon stimulation with anti-IgM antibody alone; however, the difference of cell death between the mutant and WT B cells diminished significantly in the presence of anti-CD40 or IL-4.
BAFF signaling plays a critical role in B-cell survival and enhanced BAFF signaling has been linked to B cell-mediated autoimmune diseases (Lesley et al., 2004; Thien et al., 2004). To test whether the Cbl-dko mutation affected BAFF signaling, we cultured Cbl-dko and WT B cells for 4 days in the presence or absence of BAFF and then analyzed the rates of cell apoptosis by flow cytometry. We found that the addition of BAFF rescued both Cbl-dko and WT B cells from apoptosis as compared to the cultured cells without BAFF. In the immature B cell compartment, WT B cells exhibited a slightly better survival rate than Cbl-b-dko B cell. These results thus suggset that BAFF signaling is not enhanced in Cbl-dko B cells (Fig. 3g).
Taken together, we conclude that Cbl-dko B cells are neither hyperresponsive to BCR stimulation both in vivo and in vitro, nor are they resistant to anti-IgM induced apoptosis or exhibiting an enhanced BAFF signaling. These results also suggest that manifestation of the SLE-like disease in Cbl-dko mice is unlikely a consequence of generalized B-cell hyperactivation or improved survival mediated by BAFF signals.
While B-cell tolerance can be achieved through different mechanisms such as clonal deletion, BCR editing, and anergy of autoreactive B cells (Chen et al., 1995; Goodnow et al., 1988; Goodnow et al., 1995; Nemazee and Buerki, 1989), induction of anergy to self-antigen is the last safe-guard to prevent autoreactivity (Rajewsky, 1996). To determine whether the Cbl-dko mutation affected B-cell anergy, we crossed Cbl-dko mice to BCR transgenic mice that expressed membrane IgM specifically recognizing hen-lysozyme (HEL) (IgHEL) and soluble HEL (sHEL) transgenic mice. We found that approximately 50 % (3/6) of Cbl-dko IgHEL sHEL mice became sick and eventually died within three months of age. Further analyses revealed that Cbl-dko IgHEL B cells in Cbl-dko IgHEL sHEL mice were not anergic to the HEL antigen, as they expressed a higher level of IgMa (encoded by IgHEL transgene), exhibited a higher sHEL-binding activity, and entered the stage of mature (AA4.1lo HSAlo) B cells (Fig. 4a and 4b). Additionally, upon anti-IgM stimulation the mutant B cells efficiently up-regulated CD86 and MHC-II and elicited Ca2+ influx albeit at a slightly low level than that in WT cells (Fig. 4c). On the contrary, the WT IgHEL B cells from IgHEL sHEL double transgenic mice exhibited typical phenotypes of anergic B cells, as they significantly downmodulated cell surface IgMa and HEL-binding ability and failed to develop into follicular B cells (Fig. 4a and 4b). Anti-IgM stimulation could not induce CD86 and MHC-II expression, nor did the Ca2+ response in these anergic B cells (Fig. 4c). Based on these data, we conclude that B cell anergy is at least one of the major reasons that contribute to the manifestation of the SLE-like autoimmune disease in these mice.
The strength of BCR-proximal signaling plays a critical role in peripheral B-cell tolerance and activation (Monroe, 2004; Rajewsky, 1996). It may also dictate the development and differentiation of immature B cells into follicular, B1, and MZ B cells, as it has been shown that strong BCR signaling facilitates the development of MZ and B1 B cells whereas weak BCR signaling favors the differentiation of follicular B cells (Casola et al., 2004). Since the Cbl-dko mutation altered the ratios of follicular B cells to B1 and MZ B cells (Fig. 1c), we decided to assess whether BCR-proximal signaling was enhanced in Cbl-dko B cells after anti-IgM stimulation. We found that stimulation of Cbl-dko B cells elicited markedly enhanced and prolonged tyrosine phosphorylation of total cellular proteins as compared to that of WT B cells (Fig. 5a). In particular, the mutant B cells exhibited significantly protracted higher levels of tyrosine phosphorylation of Ig-α, Syk, PLCγ-2, and Vav, as well as Erk activities, than did the control cells (Fig. 5b). Additionally, a prolonged and elevated level of Ca2+ mobilization was also found in Cbl-dko B cells than in WT cells (Fig. 5c). Surprisingly, the level of tyrosine phosphorylation of BLNK was dramatically decreased in the mutant B cells as compared to that in WT B cells (Fig. 5b), indicating that the Cbl-dko mutation exerted a differential effect on BCR-proximal signaling pathways. A strong phosphorylated band of approximate 70–75 kD was reproducibly coimmunoprecipitated with BLNK, despite the identity of this protein remained unclear (Fig. 5b). Taken together, we conclude that Cbl proteins differentially control both the strength and duration of multiple BCR-proximal signaling pathways. Additionally, since most BCR-proximal signaling pathways are enhanced whereas BLNK phosphorylation is attenuated in Cbl-dko B cells, we propose that the loss of coordination between these BCR-proximal signaling pathways is likely responsible for the impaired B-cell tolerance in Cbl-dko mice.
Cbl proteins negatively regulate TCR signaling by promoting downmodulation of the TCR complex and ubiquitination of intracellular signaling components such as Lck and PI-3 kinase (p85) (Fang and Liu, 2001; Naramura et al., 2002; Rao et al., 2002b). Since the BCR delivers signals in a manner similar to the TCR, we investigated whether the Cbl-dko mutation affected BCR downmodulation and ubiquitination of BCR-downstream signaling components. To determine whether BCR downmodulation was blocked in the absence of Cbl proteins, we cross-linked cell-surface IgM of Cbl-dko mutant and WT B cells with biotin-anti-IgM (Fab’)2 for various periods of time and then monitored the remaining cell-surface IgM by staining the cells with fluorescent-coupled streptavidin (Fig. 6a). In the absence of cross-linking, the mutant B cells expressed slightly higher levels of cell-surface IgM than did the WT cells. While BCR cross-linking for 5 min already resulted in a significant loss of cell-surface IgM on the WT B cells, the same treatment induced little change with respect to the level of cell-surface IgM on the mutant B cells for at least 20 min, indicating that Cbl proteins indeed play a critical role in BCR downmodulation.
Cbl proteins form complexes with BCR-downstream signaling components, including Ig-α Syk, PLC-γ2, BLNK, PI-3 kinase (p85), and Vav. To explore whether Cbl protein-mediated ubiquitination was involved in BCR downmodulation and BCR signaling, we examined ubiquitination of these signaling components in WT and Cbl-dko B cells after anti-IgM stimulation. We found that Ig-α and Syk were heavily ubiquitinated in WT but not Cbl-dko B cells (Fig. 6b). By contrast, we could not detect any meaningful ubiquitination of PLC-γ2, BLNK, PI-3 kinase (p85), and Vav in either WT or Cbl-dko B cells (data not shown). Interestingly, despite the ubiquitination, the amounts of Ig-α and Syk in WT B cells did not seem to be affected even at 60 min after the stimulation (Fig. 6b), suggesting that Cbl proteins might regulate the signaling of these molecules through a non-degradation mechanism.
Taken together, our results indicate that Cbl proteins selectively promote ubiquitnation of the BCR-downstream signaling components including Ig-α and Syk during BCR activation. Since Ig-α is constitutively associated with membrane IgM and its ubiquitination state is closely correlated with the BCR downmodulation, it is therefore likely that Cbl proteins downmodulate the activated BCR, hence the BCR signaling by promoting Ig-α ubiquitination.
In this study, we show that the simultaneous ablation of E3 ubiquitin ligases c-Cbl and Cbl-b in B cells results in manifestation of SLE-like autoimmune disease, evidenced by the high levels of serum autoantibodies against the dsDNA and ANA, as well as pathological alterations in kidney and other major organs. The mutant B cells were not generally hyper-responsive to antigen stimulation in terms of antibody production and proliferation. However, BCR anergy to soluble protein antigen (sHEL) was impaired. Since in this mouse model the c-Cbl and Cbl-b are simultaneously inactivated only in B cells, we conclude that Cbl proteins control B cell-intrinsic checkpoint of tolerance induction.
B cell-mediated autoimmunity is frequently linked to the hyperactivation of B cells. In this regard, mice deficient in tyrosine kinase Lyn, tyrosine phosphatase SHP-1, or membrane receptor CD22 exhibit B-cell hyper-responsiveness upon BCR stimulation and manifest systemic autoimmune diseases (Cyster and Goodnow, 1995; Hibbs et al., 1995; O’Keefe et al., 1996). In addition to this mechanism, mutations that influence B-cell apoptosis and survival may also affect immune tolerance, as the germline deletion of PKC-δ or over-production of BAFF in mice, both of which promote the survival of B cells, cause severe autoimmune diseases (Lesley et al., 2004; Saijo et al., 2003; Thien et al., 2004). In contrast to these two mechanisms, we found that Cbl-dko B cells were not hyperactive to antigen stimulation both in vitro and in vivo. They were not resistant to BCR-induced apoptosis, nor did they exhibit any enhancement in BAFF signaling. These findings thus suggest that Cbl proteins control B-cell tolerance through a different mechanism. It is generally believed that B-cell tolerance may occur at immature B cell stage. In contrast to mature B cells which are activated when encountering an antigen, immature B cells usually become tolerized upon BCR triggering (Allman et al., 2001; Carsetti et al., 1995; Loder et al., 1999; Monroe, 2004; Rajewsky, 1996). Under physiological condition, immature B cells may take 3–4 days to become immune competent mature B cells (Allman et al., 1993; Rolink et al., 1998). This period of B-cell maturation naturally constitutes a time window when autoreactive B cells can be checked by various tolerance mechanisms such as clonal deletion, BCR editing, and anergy (Chen et al., 1995; Goodnow et al., 1995; Nemazee and Buerki, 1989). Our preliminary data suggest that the Cbl-dko mutation may expedite B-cell maturation (Supplementary Fig. 3). We therefore propose that this alteration could dramatically shorten the susceptible period of immune tolerance against autoreactive B cells, consequently breaking down the immune tolerance. This hypothetic model of B-cell tolerance is also supported by a recent observation showing that administration of female hormone prolactin may concomitantly facilitate B-cell maturation and development of SLE-like disease in anti-DNA antibody transgenic mice (Peeva et al., 2003). Since our results revealed that B-cell anergy to soluble HEL antigen was impaired in Cbl-dko mice, more experiments are needed to assess whether a shortened duration of B-cell maturation is directly responsible for the impaired B-cell anergy to autoantigen in Cbl-dko mice. It should be noted that we also found that while the mutant B cells still underwent BCR editing, the efficiency of the secondary rearrangement of the κ chains in the absence of Cbl proteins seemed to be less efficient (Supplementary Fig. 4b). It is therefore necessary to further investigate whether BCR editing is partially impaired by Cbl-dko mutation. Finally, since we found that Cbl-dko mice possessed significantly more B1 B cells, it remains to be determined whether the observed autoimmune symptom is linked to the abnormal development and function of B1 B cells.
Molecular mechanisms by which Cbl proteins regulate BCR signaling remain unclear. Previous experiments show that both c-Cbl and Cbl-b may function as E3 ubiquitin ligases in T cells (Joazeiro et al., 1999). c-Cbl and Cbl-b directly or indirectly form complexes with the Ig-α, Syk, PLC-γ2, PI-3 kinase (p85), Vav, and BLNK ((Bachmaier et al., 2000; Fang and Liu, 2001; Rao et al., 2002a; Sohn et al., 2003) and data not shown). However, we noted that in B cells the Cbl-dko mutation abolished BCR-induced Ig-α and Syk ubiquitination but it did not affect the ubiquitination states of PI-3 kinase, PLC-γ2, Vav, and BLNK, despite the fact that tyrosine phosphorylation of these molecules was markedly altered. Since Ig-α and Syk function at the top of the BCR signaling cascade, our results thus support the idea that Cbl proteins may negatively regulate BCR signaling cascade at the top of BCR-induced tyrosine phosphorylation cascade by promoting Ig-α and Syk ubiquitination. As we could not detect any meaningful degradation of Ig-α and Syk even after one hour of BCR stimulation, we believe that the ubiquitination of Ig-α and Syk by Cbl proteins might not direct them for degradation, but rather alter their transportation and/or association with other molecules during BCR signaling. A similar observation that PI-3 kinase (p85) is ubiquitinated but not degraded by Cbl-b has been reported in T cells (Fang and Liu, 2001). This conclusion of course cannot exclude the possibility that only a small fraction of Ig-α and Syk are ubiquitinated by Cbl proteins so that the degradation of these molecules is below the detectable level in our assay system.
It should be mentioned that T cells in our Cbl-dko mice were deficient in Cbl-b. Since our Cbl-b−/− mice are susceptible to autoimmune diseases (Chiang et al., 2000), it is possible that in Cbl-dko mice the Cbl-b−/− T cells also contribute to the development of SLE-like disease. However, since we did not find a similar autoimmune symptom in the littermate Cbl-b−/− (c-cblf/f cbl-b−/− CD19-Cre−) mice, we believe that B cell-intrinsic ablation of c-Cbl and Cbl-b are necessary for the development of the SLE-like diseases in our animal model. In supporting this idea, our results showed that Cbl-dko B cells indeed exhibited B cell-intrinsic alterations in terms of maturation and BCR signaling. Additionally, anergy of IgHEL B cells to sHEL was abolished, indicating that a B cell-intrinsic defect has developed in the absence of Cbl proteins. Further experiments are certainly required to assess to which extent B cell-intrinsic Cbl-dko mutation affects B-cell tolerance induction and whether Cbl-dko B cells are sufficient for the manifestation of the SLE-like diseases in the absence of T-cell help.
c-Cbl-floxed, Cbl-b deficient and CD19-Cre transgenic mice were described previously (Naramura et al., 2002). To generate Cbl-dko mice, c-Cbl floxed mice were crossed to Cbl-b−/− mice and then to CD19-Cre transgenic mice were kindly provided by A. Tarakhovsky and K. Rajewsky (Rickert et al., 1997). Mice used in this study were of a mixed C57BL/6 and 129 background. IgHEL and sHEL transgenic mice were originally generated by Goodnow et al. (Goodnow et al., 1988) and were kindly provided by A. Tarakhovsky. All the mice used in this work were maintained at The Twinbrook II Facility of NIAID and the Columbia University Hammer Health Science Center mouse facility under specific pathogen-free conditions according to institutional guidance.
Anti-CD5, CD21, CD23, CD24/HSA, B220, IgM, IgD, CD69, CD86, I-Ab, and AA4.1 antibodies were from BD PharMingen. The purified and biotinylated goat anti-mouse IgM F(ab’)2 antibody used for BCR cross-linking were from Jackson Immunoresearch Laboratory. Anti-Ig-α antibody was from Dr. Susan K. Pierce. Anti-BLNK/BASH antibody was from Dr. Andrew Chan and Dr. Daisuke Kitamura. Anti-Syk, PLCγ-2, Vav, ubiquitin, phospho-ERK, ERK1/2 were from Santa Cruz Biotechnology, Inc. Anti-phosphotyrosine antibody (4G10) was from Upstate. BAFF was from Apotech Biochemicals. Anti-CD40 and IL-4 were from BD PharMingen.
6–10 week-old mice were immunized i.p. with 50 μg of NP-KLH for TD immune response, NP-Ficoll for type-I TI immune response, or NP-LPS for type-II TI immune response. The antigens were precipitated in 100 μl of Imject-Alum adjuvant (Pierce). Immunized mice were bled from the tail vein on day 7 and 14 after primary immunization. The titers of anti-NP-specific antibodies of different Ig isotypes were determined by ELISA as described previously (Chiang et al., 2000).
B cells were purified using MACS column according to a B-cell enrichment protocol (Miltanyi Biotek) and were more than 95% pure based on cell surface CD19 staining. FACS purification of T1 and T2 (AA4.1hi HSAhi) and follicular (AA4.1lo HSAlo CD21hi CD23hi) B cells was performed after staining cells with anti-AA4.1, HSA, CD23, and CD21. Purified B cells (1×105 cells/well) were stimulated with 10 μg/ml anti-IgM F(ab’)2 antibody or 10 μg/ml of LPS for two days in a 96-well plate. For the anti-CD40 antibody and IL-4 culture, 50 μg/ml anti-CD40 antibody or 20 U/ml IL-4 were included in the culture. To determine the rates of cell proliferation, cultured cells were pulsed with [3H]-thymidine for 16 hrs. Cells were then harvested on a cell harvester and [3H]-thymidine incorporation was measured on a β-counter. To determine the dependence of cell survival on BAFF, purified immature or mature B cells were cultured for up to 4 days in the presence of recombinant BAFF (100 ng/ml). Apoptotic cells were quantified by staining the cells with FITC-conjugated Annexin V and PI. Up-regulation of cell surface CD69, CD86, and I-Ab was determined by flow cytometry.
Freshly purified splenic B cells were loaded with Fura-red and Fluo-4 (Molecular Probe) in HBSS buffer containing 1% FBS at 37°C for 30 min. After washing once with HBSS buffer, cells were stimulated with 5 μg/ml anti-IgM (Fab’)2 fragment. Increase of intracellular Ca2+ concentration in B220+ B cells was recorded in real time by flow cytometry for 300 seconds.
Purified B cells were stained with biotinylated anti-IgM (Fab’)2 at 4°C for 15 min. After washing with PBS, cells were incubated at 37°C pre-warmed HBSS buffer containing 1% FBS to allow internalization of BCR-anti-IgM (Fab’)2 complexes to occur. After various periods of incubation, cells were immediately transferred into cold HBSS buffer containing 0.1% azide to stop further internalization. Cell surface remaining anti-IgM (Fab’)2 were stained with streptavidin-PE and quantified on an LSR II.
Immunoprecipitation and Western-blot analyses were performed as previously described (Naramura et al., 2002). In brief, purified B cells were stimulated with anti-IgM (Fab’)2 for various periods and lysed in RIPA buffer containing a mixture of proteinase and phosphatase inhibitors (0.1 ng/ml Aprotinin, 0.01 ng/ml Leupeptin, 0.2 mM PMSF, 1 mM NaF, and 1 mM NaVO4). Ig-α, Syk, BLNK, Vav, and PLCγ-2 in the cell lysates were immunoprecipitated with the corresponding antibodies and subjected to electrophoresis and immunoblotting. The levels of tyrosine phosphorylation were determined using an anti-phosphotyrosine antibody. The levels of protein ubiquitination were determined using an anti-ubiquitin antibody (Santa Cruz Biotechnology, Inc).
Mouse tissues were harvested, snap frozen in liquid nitrogen, and embedded in OCT embedding medium (Sakura Finetek). 8-μM sections were air-dried and fixed with acetone. H/E staining was performed according to a standard protocol (Miyamoto et al., 2002). Immuno-fluorescent staining was performed using the following reagents: anti–IgM-FITC, anti–IgG-biotin (BD Biosciences), streptavidin–Alexa 568 (Molecular Probes). Anti-nuclear antibodies (ANA) were detected by intracellular staining of the Hep2 cells with mouse serum (1:100 dilution), followed by FITC-conjugated anti-mouse-IgG. Evans Blue staining was used to visualize cytoplasm.
Supplementary Figure 1. Generation of B cell-specific c-Cbl and Cbl-b dko (Cbl-dko) mice. (a) Restriction maps of c-Cbl WT, floxed (c-Cblf ) and deleted (c-Cbldel) alleles. Sc: Sac I digestion of the WT and c-Cbl mutant alleles. (b) Southern blot analysis of deletion of c-cblf/f gene in B cells. Genomic DNA from tail of c-Cblf/+ mice (f/+:tail), c-Cblf/+ Cbl-b−/− CD19-Cre (f/+: B) and c-Cblf/f Cbl-b−/− CD19-Cre (f/f: B) B cells were digested with Sac I. Solid black bar: probe. (c) Western blot analysis of c-Cbl expression in purified B cells. Shown are the expression levels of c-Cbl proteins in purified WT (f/+: B) and Cbl-dko (f/f: B) B cells. (d) Deletion efficiency of Cre-loxP mediated recombination in BM and splenic B-cell subsets. Shown are the histograms of YFP-expression in pro/pre (B220lo IgM−), immature (B220lo IgM+), and recirculating (B220hi IgM+) B cells in BM and immature (AA4.1hi HSAhi) and mature (AA4.1lo HSAlo) B cells in spleen. Percentages of YFP+ cells are indicated in the histograms. Data were from ROSA26-YFP and CD19-Cre double transgenic mice.
Supplementary Figure 2. Impaired proliferation of Cbl-dko B cells upon BCR stimulation. Immature (AA4.1hi HSAhi) and Mature (AA4.1lo HSAlo) B cells were purified from spleens of WT and Cbl-dko mice by FACS and stimulated with anti-IgM (10 μg/ml) or anti-IgM plus IL-4 (20 U/ml). Cell proliferation is determined by [3H]-thymidine incorporation. Shown are mean values and standard deviations of triplicates from one representative of three independent experiments.
Supplementary Figure 3. Expedited maturation of Cbl-dko B cells. (a) In vivo B cell maturation. Newly generated B cells in Cbl-dko and WT mice were labeled by BrdU. To do so, we labeled Cbl-dko and WT B cells with 5-bromo-2′-deoxyuridine (BrdU) with BrdU-containing water for 3, 5, and 7 days. Total splenic B cells were stained with anti-BrdU, anti-HSA and anti-IgD antibodies. Shown are the dot plots of HSA and IgD-expression profiles of the gated BrdU+ B cells from Cbl-dko and WT mice. HSAhi IgDlo/hi B cells represent newly generated follicular B cells, whereas HSAlo IgDhi cells are mature B cells. We found that after 3-day BrdU labeling, both WT and Cbl-dko mice possessed a similar minimal numbers of BrdU+ mature follicular (HSAlo IgDhi) B cells. At day 5, while BrdU+ B cells with the phenotype of follicular B cells were not altered in WT mice, there were 2-times more (19% vs 10%) BrdU+ Cbl-dko B cells became follicular B cells. By day 7, the number of BrdU+ follicular B cells in WT control mice increased to 17% of total BrdU+ B cells; however, more than 35% of BrdU+ Cbl-dko B cells maturated into follicular B cells. (b) Cell cycle analysis. To determine the fraction of splenic immature and mature B cells in active cell cycle, we stained DNA content of peripheral B cell with DAPI. Immature and mature B cells were identified based on anti-HSA and AA4.1 staining. BM B cells which contained a high percentage of dividing pro/pre B cells were used as positive control. Our results indicated that Cbl-dko and WT mice possessed a comparable number of dividing mature B cells. (c) B-cell maturation in vitro. To determine the speed of B-cell maturation in vitro, we purified AA4.1hi HSAhi immature B cells from the spleen of WT and Cbl-dko mice by FACS. Purified B cells were cultured for various days in the presence of a high concentration of BAFF (200 ng/ml). Percentages of mature (AA4.1lo HSAlo) B cells appeared in the culture was monitored by flow cytometry after staining with anti-AA4.1 anti-HSA. Shown at the top are dot plots of flow cytometric analysis of mature (AA4.1lo HSAlo) B cells. The percentages of mature B cells are indicated in the plots. The bottom curves are the statistics of the percentages of mature B cells after various days of culture. The results represent more than three independent experiments.
Supplementary Figure 4. BCR editing in Cbl-dko B cells. (a) RS Rearrangements in Cbl-dko immature and pre-B cells. A diagram shows the Ig κ locus indicating the relative locations of the primers (arrows) used to detect RS rearrangements. IRS and RS elements are represented by closed triangles and the Vκ RSS by an empty triangle. Agarose gel analysis of RS-Vκ, RS-IRS, and APRT PCR reactions was performed on the indicated samples. Negative images of ethidium bromide-stained gels are shown. (b) LM-PCR assays for V(D)J recombination reaction intermediates in Cbl-dko immature and pre-B cells. In a top diagram, locations of primers used to detect primary and secondary break intermediates are indicated on the κ locus with arrows. RSSs are shown as empty or shade triangles. Bottom diagrams show primary and secondary RSS break intermediates ligated to the linker. Linker-ligated DNA was analyzed by PCR for primary and secondary RSS breaks as described in upper diagram. Negative images of ethidium bromide stained agarose gel analyses of PCR reactions are shown. APRT is a non-rearranging control locus.
We thank Drs. K. Calame and Y. R. Zou for critical reading of the manuscript and members of H. Gu’s lab for discussion. We are thankful to Dr. A. Chan and D. Kitamura for antibodies, to A. Tarakhovsky and K. Rajewsky for CD19-cre mice. This work was supported by the NIH intramural research program, The Irene Diamond Fund/Professorship, and a grant from The NIH (AI 062931). Y. K. was partly supported by postdoctoral fellowships from The Uehara Memorial Foundation and The Charles Revson Foundation, and M. S. by a grant from The NIH (HL 48702).
Publisher's Disclaimer: This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final citable form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.