Transitional B Cell Fate Is Associated with Developmental Stage-Specific Regulation of Diacylglycerol and Calcium Signaling upon B Cell Receptor Engagement¹

The BCR plays a fundamental role in B lymphocyte development (1, 2). Surface IgM-expressing immature B cells that do not recognize self-Ag migrate from the bone marrow to the spleen as immature or transitional B cells. In the spleen, they then develop into mature follicular (Fo)⁴ and marginal zone (MZ) B cells (3). The transitional B cell population is heterogeneous and can be divided at least into three subsets based on variable expression of the cell surface markers (3, 4, 5). Cells of the earliest transitional (T1) stage display an IgM^highHSA^highIgD⁻CD21⁻CD23⁻ phenotype (3, 4). The late transitional (T2) B cells can be classified into two subsets; the T2 subset, which serves as a precursor for mature FoB cells (IgM^highIgD^highCD21⁺CD23⁺) originally defined by Allman et al. (4) and Loder et al. (3), and the newly defined T2-pre-MZB (IgM^highIgD^highCD21^highCD23⁺), which serves as a precursor for mature MZB cells (5, 6, 7). The Loder et al. (3) T2 B cell population, which includes T2-pre-MZ B cells, expresses AA4.1 and high levels of heat-stable Ag (HSA) (4, 8, 9). Thus, T1 B cells may give rise to T2 B cells, which either serve directly as a precursor for FoB cells or may transit through a T2-pre-MZ stage before becoming mature MZB cells. The molecular mechanisms that regulate the developmental transitions between immature transitional and mature B cells remain unclear.

According to the current model for FoB cell development, self-reactive T1 B cells are eliminated in response to strong BCR-induced signals (negative selection), whereas non-self-reactive T1 cells develop into T2 B cells via either a default pathway or tonic signaling, and then become resistant to BCR-induced apoptosis and ultimately require BCR-induced signals for survival and further differentiation into mature FoB cells (positive selection) (8, 9, 10, 11, 12, 13, 14, 15). Analysis of the peripheral B cell repertoire, in both mice and humans, supports the hypothesis that peripheral immature to mature FoB cell differentiation results from positive selection (10, 16). Although the functional properties of transitional B cell subsets in vitro remain controversial (3, 4, 14, 15, 17), experimental evidence for the disparate biological outcomes of negative and positive selection, as well as the molecular mechanisms that underlie differential responses to BCR engagement within transitional B cell subsets, is now emerging. Recently, we and others have demonstrated that T1 and T2 B cell subsets exhibit distinct biological responses upon BCR ligation in vitro: T1 B cells die, whereas T2 B cells are resistant to apoptosis and display phenotypic characteristics of mature FoB cells (14, 15). However, the understanding of biochemical mechanisms that underlie the disparate biological responses of T1 compared with T2 B cells remains elusive.

BCR engagement leads to recruitment of multiple enzymes including Bruton’s tyrosine kinase (BTK) to the plasma membrane, forming a large multicomponent signaling complex termed the BCR signalosome (1, 18). We and others have previously demonstrated that BTK deficiency leads to a severe mature FoB cell deficiency, despite intact T2 and mature MZ B cell subsets, termed Xid (xid) in mice (19, 20, 21, 22). An xid-like phenotype is also observed in animals with gene-targeted deletion of PLC-γ2, which is a direct target of BTK, and B cell linker protein (BLNK), an adaptor which facilitates BTK and PLC-γ2 interaction (23, 24, 25, 26, 27). These observations support an active role for BCR signaling, particularly the BTK/PLC-γ2 signaling axis, in the differentiation of T2 into mature FoB cells (3, 4, 15, 25, 26, 28).

Biological significance of the BTK/PLC-γ2 signaling axis in B cell survival and/or differentiation is further supported by studies suggesting that one of the molecular defects in BTK- and PLC-γ2-deficient B cells is their inability to activate IκB kinase and NF-κB (4, 26, 28, 29, 30). Consistently, deletion of IκB kinase results in a reduction of the mature FoB cell compartment (31, 32). We have also shown that the transcription factor NF-κB up-regulates the antiapoptotic protein Bcl-x_L in response to BCR cross-linking in a BTK/PLC-γ2-dependent manner (33). Moreover, BCR-induced NF-κB activation proceeds via protein kinase Cβ (PKCβ) (34, 35, 36). Together, these studies suggest that the BTK/phospholipase C (PLC)-γ2 signaling axis regulates B cell survival by activation of PKC/NF-κB signaling pathways.

Phosphatidylinositol-4,5-bisphosphate (PIP₂) is at a critical nexus in BCR signaling because it serves as a substrate for both PLC-γ2 and PI3K, which act on distinct targets to modulate B cell fate (37, 38, 39, 40). PIP₂ is synthesized in response to BCR via BTK-dependent recruitment of PI5K (40). PLC-γ2 catalyzes the hydrolysis of PIP₂ to produce two lipid second messengers, DAG and inositol 1,4,5-triphosphate (IP₃) (41). DAG activates various signaling pathways, including those of PKC and Ras, whereas IP₃ binds to its receptors on the endoplasmic reticulum triggering intracellular calcium (Ca²⁺) release (34, 42, 43, 44). High levels of IP₃ facilitate emptying of the intracellular Ca²⁺ stores, which signals sustained influx of extracellular Ca²⁺ via calcium release activated channels (45). Additionally, L-type and L-type-like channels have been implicated in both initial and sustained mobilization of calcium in B cells and B cell lines (46, 47). Using DT40 chicken B cells, we recently provided genetic evidence that BTK and PLC-γ2 are necessary for optimal production of DAG upon BCR ligation (34, 48). Prior studies have demonstrated that BCR-induced proliferation of mature FoB cells correlated with PIP₂ hydrolysis, intracellular Ca²⁺ increase, and PKC activation, whereas BCR-induced apoptosis of immature B cells correlated with an increase in intracellular Ca²⁺ in the relative absence of IP₃ production (49, 50).

Here, we provide evidence supporting the hypothesis that T2 to mature FoB cell differentiation requires BCR-mediated DAG and calcium signals via a BTK- and PLC-γ2-dependent mechanism. In contrast, calcium signals without significant DAG production in T1 cells may result in tolerance following BCR engagement. These studies indicate that developmental stage-specific DAG and calcium signaling may regulate the fate of developing splenic B cells.

The generation of Btk-deficient (btk^−/−), PLC-γ2-deficient (plc-γ2^−/−) and Notch2 haploinsufficient (notch2^+/−) mice has been previously described (19, 25, 51). For wild-type controls, 129/Sv × C57BL/6 or C57BL/6 mice (The Jackson Laboratory) were used. All mice were ∼6 wk of age. Mice were treated humanely in accordance with federal and state government guidelines and their use was approved by the institutional animal committee.

Primary B lymphocytes were isolated using MiniMACS magnetic sorting by negative selection (CD43 depletion) to avoid inadvertent activation of B cells as previously described (14). The purity of B cells isolated in this manner was ∼90–95% as verified by flow cytometry (FCM) analysis using anti-B220 and anti-IgM Abs (BD Pharmingen). All purifications were performed at 4°C, and cells were used immediately. Splenic B cell subsets were purified based on a combination of the schemes described previously (3, 4, 5) (Table I). To avoid use of anti-IgM Abs, which could influence BCR signaling under study, HSA and CD21 were used for cell sorting and to exclude T2-pre-MZ B from T2 cells. T1, T2, and mature FoB cell subsets for DAG studies were FACS purified using Abs directed against HSA, CD21, and CD23, and the purity of each subset was between 85 and 95% as previously described (14). In some experiments, DAG production in T1 B cells was compared with a pool of T2/mature FoB cells, and T1 B cells were purified using a series of MACS depletions and positive selections, which included CD43 depletion followed by CD23 depletion and AA4.1-positive selection. The purity of T1 (CD43⁻CD23⁻AA4.1⁺) and T2/mature FoB cells (CD43⁻CD23⁺) was >95% as determined by FCM. T1 and T2 B cells for use in IP₃ studies and phospho-PKC immunoblotting were obtained by MACS enrichment for AA4.1⁺ transitional B cells, followed by FACS purification using Abs directed against HSA and CD23 to sort AA4.1^highHSA^highCD23⁻ (T1) and AA4.1^midHSA^highCD23⁺ (T2) B cells. The purity of the T1 subset was ∼95%, and the purity of the T2 subset was ∼75–90% as determined by FCM. Mature FoB cells were MACS-enriched from the AA4.1-negative fraction by CD43 depletion followed by CD9 depletion (to remove mature and precursor MZ B cells). The purity of the mature FoB subset was ∼85% as determined by FCM.

Purified B cells or B cell subsets were labeled for 6 h with palmitic acid (2.5 μCi/ml), as a fatty acid precursor. Lipid synthesis was monitored in wt, btk^−/−, and plc-γ2^−/− B cells, and the production of DAG was quantified as previously described (34, 48). Labeled B cells were washed twice with PBS and stimulated through the BCR with 20 μg/ml goat anti-mouse IgM F(ab′)₂ for the indicated times. The reactions were stopped by the addition of cold PBS. Cells were centrifuged, and the pellet was used for lipid extraction following suspension in 2 ml of chloroform-methanol (2:1 v/v) and addition of 0.4 ml of 0.5 N HCl. The mixture was shaken vigorously, centrifuged, and the upper (aqueous) phase was discarded while the lower (organic) phase was washed twice with theoretical upper phase (52). The radioactivity of lipids was determined on an aliquot fraction by scintillation counting. Dipalmitin carrier standard was added, the lipid phase evaporated gently under N₂ flux, and the residue dissolved in a minimum volume of chloroform-methanol (2:1 v/v). DAG was separated by TLC on silica gel G plates using a double development system and was detected under UV light after spraying the plates with primulin as described previously (53). The area containing DAG was scraped, and the radioactivity contents were determined. The data represent the mean of three experiments performed in triplicate and analyzed using Student’s t test.

The soluble immunoprecipitation (IP) profile for B cells was determined by adapting a described previously protocol (54). Briefly, FACS-purified B cell subsets (∼5 × 10⁶) were cultured in 0.5 ml of inositol-free DMEM containing 30 μCi of [³H]inositol (PerkinElmer) for 4 h. Cells were harvested, washed twice with 1 ml of PBS, and suspended in 200 μl of PBS. One hundred microliters of cells were treated with 25 μl of PBS with or without 20 μg/ml goat anti-mouse IgM F(ab′)₂. Treatments were stopped by adding 6.6 μl of 10 N HCl and chilling cells on ice. Soluble IPs were extracted by adding 465 μl of chloroform/methanol (1:2 v/v). The mixture was vortexed for 2 min at maximum speed, followed by addition of 156 μl each of chloroform and 2 M KCl and another 2 min of vortexing. The lysate was spun at 13,000 × g for 5 min, and the supernatant was recovered. Samples were analyzed by HPLC, with the IPs resolved by a Whatman partisphere SAX strong-anion exchange column (4.6 × 125 mm) and a linear gradient from 10 mM to 1.7 M ammonium phosphate (pH 3.5) for 25 min, followed by elution with 1.7 M ammonium phosphate for 20 min.

Total splenocytes or MACS-enriched B cells were stimulated with 20 μg/ml goat anti-mouse IgM F(ab′)₂ Abs for the indicated times, and then immediately fixed with 2% paraformaldehyde. After centrifugation, cells were permeabilized (BD Cytofix/Cytoperm), and were intracellularly labeled with a biotinylated Ab raised against PI(4,5)P₂ (Z-B045; Echelon Biosciences), or a rabbit polyclonal Ab raised against phosphorylated PKCδ (9374S; Cell Signaling). Cells were labeled with Abs directed against CD19, HSA, and CD21 for the identification of splenic B cell subsets. PIP₂ was visualized by PerCP-Cy5.5-conjugated streptavidin. Phospho-PKCδ was visualized by anti-rabbit Alexa 488. Data were acquired on a BD Biosciences FACSCalibur flow cytometer. Results are expressed as mean fluorescence intensity and are representative of multiple experiments.

FACS-sorted T1 and T2 B cells and MACS-enriched mature FoB cells were rested on ice in complete RPMI 1640 for 3 h, at room temperature for 15 min, and then warmed to 37°C for 15 min before stimulation with anti-IgM F(ab′)₂ for 2, and 5 min, or left nonstimulated. For immunoblot analysis of phospho-pan-PKC and phospho-PKCδ (Cell Signaling), cytoplasmic extracts were prepared in radioimmunoprecipitation assay buffer containing protease inhibitor (Roche), and the cytoplasmic fractions were cleared by centrifugation at 14,000 rpm for 20 min. Protein was quantified by Bradford assay and 20 μg of total protein per point was denatured in Laemmli reducing buffer by boiling at 100°C for 5 min. Proteins were separated by SDS-PAGE and were electrotransferred onto polyvinylidene difluoride membrane (Immobilon; Millipore). The membranes were sequentially probed with Abs specific for p-cPKC(pan), p-PKCδ (Cell Signaling), and actin (Santa Cruz). Bound Abs were revealed by HRP-conjugated goat anti-rabbit IgG Ab (Pierce), followed by ECL detection (Pierce) on autoradiography film.

Purified B cells (10 × 10⁶ cells/ml in HBSS) were loaded with 5 μg/ml indo-1-AM for 30 min at 37°C. Cells were then washed, placed on ice, and labeled with PE-conjugated anti-HSA, biotinylated anti-CD23 (visualized by PerCP-Cy5.5-conjugated streptavidin), and allophycocyanin-conjugated anti-C1qRp (clone AA4.1) for B cell subset identification. Abs raised against CD19 and CD21 were not used to avoid inadvertent stimulation of B cells. Before acquisition, cells were resuspended at 1 × 10⁶/ml in HBSS containing 1 mM calcium, warmed to 37°C for 15 min, and applied to a BD LSR II cytometer. Baseline, nonstimulated measurements were taken for 90 s, followed by the addition of 10 μg/ml goat anti-mouse IgM F(ab′)₂ Abs for the indicated time. In some experiments, cells were preincubated with 100 μM 2-aminoethoxyphenylborate (2-APB) (Calbiochem 100065) or 100 μM verapamil (Calbiochem 676777) for 15 min before application to the cytometer.

For our studies, B cell subsets were gated using a combination of Loder et al. (3) and Allman and colleagues (4, 5) schemes to classify five major splenic B cell subsets: immature (AA4.1⁺) T1 and T2 B cells, mature (AA4.1⁻) FoB and MZB cells, and the recently described MZ precursors (AA4.1^low/−CD23⁺CD21^high) (Table I) (55). B cell subsets were visualized using HSA and CD21 (Fig. 1,A), allowing for clear visual separation of T2 B cells from MZ and MZ precursors (T2-pre-MZ), which both express high levels of CD21. MZ and T2-pre-MZ can be further distinguished from each other by expression of CD23 and AA4.1 (Fig. 1,A, right panel). The HSA-CD21 MZ B cell gate was based on low reactivity to CD23; however, it is important to note that, although >80% of the cells in the MZ gate are AA4.1-negative, a significant proportion (∼50%) of these cells do retain reactivity to CD23. To ascertain that the T2 cells we are analyzing do indeed belong to the immature splenic B cell compartment, we investigated the expression of CD93/C1qRp (recognized by the mAb AA4.1), which is exclusively expressed on immature B cells (4, 56). FCM analysis revealed that neither mature FoB nor MZ B cells exhibit significant levels of reactivity to AA4.1 (Fig. 1,B). In contrast, reactivity to AA4.1 is clearly detected in T1 as well as T2 B cell subsets. We observe that T1 B cells react higher to AA4.1 than T2 (5.0- ± 0.8-fold higher and 3.6- ± 0.5-fold higher relative to mature B cells, respectively), and that AA4.1 is detected on the T2-pre-MZ subset (2.8- ± 0.7-fold higher relative to mature B cells). Additionally, all of the transitional B cell populations express high levels of HSA. Thus, all transitional B cells demonstrate characteristics of immature B cells. Additionally, disparate biological outcomes between T1 and T2 B cells are not due to differences in the cell surface expression levels of surface IgM (Fig. 1,C). We have also found that there is comparable up-regulation of the activation marker CD69 in both T1 and T2 B cells following anti-IgM F(ab′)₂ stimulation (Fig. 1 D), revealing that both T1 and T2 B cell subsets are capable of transducing BCR-mediated signals.

Using FACS-purified T1, T2, and mature FoB cell populations (Fig. 2,A), we measured [³H]DAG production in response to stimulation with anti-IgM F(ab′)₂ Abs (Fig. 2,B). Our studies reveal that BCR-induced DAG production occurs in T2 and mature B cells following both 5 min (2.1- ± 0.1-fold increase, and 5.1- ± 0.3-fold increase, respectively, compared with the nonstimulated control) and 30 min (2.8- ± 0.3-fold increase, and 4.8- ± 0.4-fold increase, respectively, compared with the nonstimulated control) stimulation with anti-IgM. In sharp contrast, T1 B cells do not produce detectable levels of DAG under similar stimulatory conditions. To rule out the possibility of an early transient production of DAG in T1 B cells, purified T1 and a mixture of T2/mature FoB cells were stimulated through the BCR for 0.5 and 1 min (Fig. 2,C). DAG production in response to BCR cross-linking was not detectable in T1 B cells at either of the early time points, whereas the mixture of T2/mature FoB cells exhibits DAG production upon BCR engagement. The kinetic profile of DAG production that we observed is comparable to previous studies performed with total B cells that were stimulated through the BCR (57). In addition to DAG, PIP₂ hydrolysis by PLC-γ2 produces IP₃ in primary wt B cells (57). Therefore, using FACS-sorted T1 and T2 B cells (Fig. 2,D), we also investigated the production of IP₃ in response to BCR stimulation (Fig. 2,E). Consistent with DAG generation, we found that IP₃ is preferentially produced in T2 relative to T1 B cells following anti-IgM stimulation (3.0- ± 0.2-fold increase, and 1.3- ± 0.5-fold increase, respectively; n = 4). Because both T1 and T2 B cells up-regulate the activation marker CD69 following anti-IgM stimulation (Fig. 1 D), reduced DAG/IP₃ production is not likely due to a general inability of T1 B cells to signal via the BCR. We estimate that the T2 cells used for the analysis of DAG may contain ∼14% T2-pre-MZ B cells, whereas <9% of the T2 cells used in the IP₃ assay were identified as potential T2-pre-MZ B cells. Because DAG and IP₃ are the products of PIP₂ hydrolysis, and we obtained similar profiles for DAG and IP₃ production, we believe that T2-pre-MZ B cells did not contribute significantly to the levels of DAG in T2 B cells, although we cannot formally exclude this possibility. Together, these data indicate that distinct stage-specific signaling mechanisms are likely to be responsible for differential lipid metabolism in the T1 and T2 B cell subsets.

To understand the mechanisms underlying differences in BCR-induced DAG production within the T1 and T2 B cell subsets, we first examined the levels of PIP₂, which serves as a substrate for PLC-γ2 in the production of DAG (58). To determine both basal and BCR-induced levels of PIP₂, we performed FCM analysis using an anti-PI(4,5)P₂ Ab. Our results demonstrate that basal levels of PIP₂ are lower in T1 relative to T2 B cells (Fig. 3,A). Although the kinetics of PIP₂ upon BCR cross-linking are similar in T1 and T2 B cells (Fig. 3,A), the magnitude of change in PIP₂ was again lower in T1 relative to T2 B cell subset. It is also noteworthy that both basal and BCR-induced levels of PIP₂ were higher in T1 B cells than in mature FoB cells. Additionally, the T2-pre-MZ cells possessed the highest basal and BCR-induced level of PIP₂ (Fig. 3,A). The higher PIP₂ level in T2 B cells is unlikely to result from contaminating T2-pre-MZ cells because similar results were obtained with B cells depleted of CD9⁺ cells, which should eliminate the majority of MZ and pre-MZ B cells (Fig. 3,B) (59, 60), as well as with B cells isolated from notch2^+/− mice, which are severely impaired in MZ and pre-MZ B cell development (Fig. 3 C) (7, 51, 61).

The generation of IP₃ induces Ca²⁺ mobilization in primary wt B cells (57). The duration and magnitude of calcium flux is likely to modulate B cell survival and differentiation by the activation of specific transcription factors including NFAT and NF-κB (62). To precisely define stage-specific Ca²⁺ mobilization within transitional B cell subsets, we stimulated purified wt B cells with anti-mouse IgM F(ab′)₂ Abs and determined Ca²⁺ flux in each splenic B cell subset by FCM. Our results demonstrate that despite differential IP₃ production, the initial Ca²⁺ peak is similar in T1 and T2 B cells following BCR engagement (Fig. 4, top panel). However, T1 B cells display higher sustained levels of Ca²⁺ relative to T2 and mature FoB cells. The initial Ca²⁺ peak appears to depend, at least in part, on IP₃Rs because pretreatment of the cells with 2-APB, a inhibitor of IP₃R-mediated Ca²⁺ release, reduced the initial Ca²⁺ peak in all B cell subsets (Fig. 4, middle panel). However, the initial Ca²⁺ peak in T1 B cells appears to be more sensitive to pretreatment with 2-APB than T2 or mature FoB cells. In contrast, sustained Ca²⁺ influx in T1 B cells appears to be less sensitive to 2-APB treatment than in T2 and mature FoB cells. Additionally, inhibition of L-type channels with verapamil led to a more severe reduction in calcium influx in T1 relative to T2 and mature FoB cells (Fig. 4, bottom panel).

DAG targets multiple signaling pathways including PKCs, chimaerins, and RasGRPs (42, 44). Our results demonstrate a reduction in BCR-mediated DAG production in T1 compared with T2 B cells. A consequence of reduced DAG production should be reduced activation of PKC in T1 relative to T2 B cells upon BCR engagement. We therefore monitored the magnitude of BCR-induced phosphorylation of DAG-responsive PKCs in transitional B cell subsets using immunoblotting and FCM. Surprisingly, BCR-induced phosphorylation of conventional PKCs and nonconventional PKCδ was detected in T1 B cells, albeit at a modestly reduced level compared with T2 as well as FoB cells (Fig. 5,A). Because sufficient numbers of FACS-sorted T2-pre-MZ and MZ B cells for immunoblotting could not be obtained, BCR-induced phosphorylation of PKCδ in T2-pre-MZ and MZ B cells was examined using FCM. Again, an increase in PKCδ phosphorylation was clearly detectible in all B cell subsets (Fig. 5,B); however, both basal and BCR-induced phosphorylation of PKCδ was higher in T2-pre-MZ and MZ B cells relative to T1, T2, and mature FoB cells. Overall, the differences in PKC phosphorylation between T1 and T2 B cells are modest when compared with the observed differences in the production of DAG (Fig. 2), suggesting that very small amounts of DAG, which may be undetectable in our assay system, are sufficient to induce PKC phosphorylation. Together, these results suggest that phosphorylation of PKCs is not the major DAG-dependent mechanism responsible for differential responses of T1, T2, and mature B cells to BCR stimulation.

Although the BTK/PLC-γ2 signaling axis appears to be dispensable in the differentiation of T1 to T2 B cells, functional integrity of this signaling axis is essential for the transition from T2 to mature B cells (14, 15, 25, 26, 28, 63). To test the hypothesis that BTK and PLC-γ2 control this process by mediating DAG production, wt, btk^−/−, and plc-γ2^−/− B cells were stimulated with anti-IgM F(ab′)₂ Abs for the indicated times and DAG production was monitored (Fig. 6). Consistent with previous reports (57), an increase in DAG production in wt B cells is observed in response to BCR cross-linking (5.1- ± 0.9-fold increase at 5 min, and 6.8- ± 1.0-fold increase at 30 min compared with the nonstimulated B cells). In contrast, BCR stimulation did not induce DAG in either btk^−/− or plc-γ2^−/− B cells. These results demonstrate that BTK and PLC-γ2 are required for BCR-induced production of DAG. They also extend our previous findings, which indicate a requirement for BTK and PLC-γ2 in BCR-directed DAG production in DT40 B cells (34, 48).

Because DAG is a strong inducer of cell survival and differentiation (34, 64), our finding that DAG is preferentially produced in T2 relative to T1 B cells (Fig. 2, B and C) correlates with the observed BCR-induced survival and differentiation of T2 B cells but not T1 B cells (9, 14, 15). It is noteworthy that similar BCR-induced DAG production is observed in T2 and mature FoB cells, suggesting that DAG plays an important role in the survival of both T2 and mature FoB cells. We also observed that T2 B cells display higher constitutive levels of DAG and IP₃, as well as PIP₂ and phosphorylated PKC, compared with T1 cells (Figs. 2, A and C; 3; and 5). These data suggest an active BCR signaling process, which is consistent with T2 B cell survival and positive selection into the mature FoB cell pool. We also demonstrated that both btk^−/− and plc-γ2^−/− B cells, which are impaired in the development of mature FoB cells, are unable to generate DAG following BCR engagement (Fig. 6). Coupled with previous genetic evidence demonstrating that the BTK/PLC-γ2 signaling axis is critical for T2 to mature FoB cell development, our results support the hypothesis that DAG plays an important role in T2 B cell survival and differentiation into mature FoB cells. It is unlikely that BTK/PLC-γ2-dependent DAG production plays a significant role in T2-pre-MZ to MZ B cell maturation because MZ B cells develop normally in BTK- and PLC-γ2-deficient mice (3, 26, 65). Thus, the BTK/PLC-γ2/DAG signaling axis may promote the differentiation of T2 into mature FoB cells; however, this pathway appears dispensable for the differentiation of T2-pre-MZ into MZ B cells, and may even inhibit the development of MZ B cells as previously proposed (65).

Although DAG has been implicated in cell survival and differentiation, the significance of PIP₂ in these processes is unknown. Despite having the lowest basal and BCR-induced levels of PIP₂, we found that mature FoB cells produce substantial levels of DAG upon BCR cross-linking. These results suggest that differences in PIP₂ levels between T1 and T2 B cells may not be responsible for the disparate DAG production observed upon BCR stimulation. Moreover, BCR signal-induced changes in PIP₂ levels may not reflect the generation of lipid second messengers (DAG and IP₃). Thus, higher levels of DAG in mature FoB relative to transitional B cells may reflect differentiation stage-specific metabolism of DAG rather than the levels of available substrate (PIP₂).

It was recently proposed that DAG-dependent activation of conventional PKCβ initiates a survival, but not a differentiation, program in response to BCR cross-linking through the activation of NF-κB and induction of Bcl-x_L (33, 36). Thus, DAG at least in part may regulate T2 B cell survival via activation of the PKC pathway. In this model, survival of T2 cells may be regulated by PKCβ, and their differentiation by nonconventional PKCs. Multiple nonconventional PKC isoforms are expressed in B cells and are activated in response to BCR cross-linking, but their specific roles in splenic B cell maturation remain largely unresolved (66). However, our data showing only minor differences in the phosphorylation of both conventional and nonconventional PKCs in T1 and T2 B cells (Fig. 5) argues against a central role for PKC-mediated signaling in transitional B cell differentiation processes. Consistently, mice deficient for either conventional and nonconventional PKCs display a milder B cell deficiency than that observed in BTK- or PLC-γ2-deficient mice (9, 19, 36, 67, 68), suggesting that other DAG-dependent and PKC-independent pathways likely contribute to T2 B cell survival and differentiation. Alternatively, DAG may regulate T2 B cell survival and differentiation via activation of the Ras-MAPK pathway. In this regard, we have previously shown that ERK, a downstream effector of Ras, is preferentially activated in T2 compared with T1 B cells (14). The Ras/ERK pathway has also previously been shown to control early B cell development, indicating its role in B cell differentiation (69, 70). Thus, DAG may activate survival via the PKCβ/NF-κB/Bcl-x_L signaling axis and differentiation via the Ras pathway (34).

Despite differential IP₃ production (Fig. 2,E), both T1 and T2 B cells mobilize Ca²⁺ in response to BCR cross-linking (Fig. 4, top panel). This observation is in agreement with the study by Yellen et al. (50), which demonstrated that intracellular Ca²⁺ is mobilized similarly in both immature and mature B cell populations despite the lack of IP₃ production in immature B cells. Together, these data suggest that the Ca²⁺ mobilization observed in T1 B cells may occur independently of PIP₂ hydrolysis, which leads to DAG and IP₃ generation. Alternatively, T1 B cells may be more sensitive to IP₃, and may thus be able to respond to lower levels of IP₃ than T2 B cells. In this regard, we observed that the initial Ca²⁺ peak in T1 B cells is more sensitive to treatment with 2-APB, an inhibitor of IP₃R-mediated calcium release, whereas sustained calcium influx in T1 B cells is less sensitive to 2-APB than in T2 and mature FoB cells (Fig. 4, middle panel). Sustained Ca²⁺ influx via calcium release activated channels has been attributed to sustained high levels of IP₃ (45); however, we have demonstrated that T1 B cells produce significantly less IP₃ compared with T2 B cells. This suggests that T1 cells may use other channels for sustained Ca²⁺ influx, such as L-type or L-type-like channels, as has been reported for B cells and B cell lines (46, 47). Consistent with this hypothesis, inhibition of L-type channels with verapamil led to a more severe reduction in calcium mobilization in T1 relative to T2 and mature FoB cells (Fig. 4, bottom panel). Taken together, these results suggest that the mechanisms by which both intracellular calcium release and sustained calcium influx occur may be different in each transitional B cell subset. Additionally, we have previously reported that verapamil inhibits the induction of NFAT, whereas 2-APB is an inhibitor of NF-κB activation (34). Differential responsiveness of transitional B cell subsets to these inhibitors (Fig. 4) suggests that following BCR engagement, T1 B cells may preferentially use NFAT, whereas T2 B cells may use both NFAT and NF-κB for the transcription of genes necessary for survival and/or differentiation. Furthermore, activation of NFAT in the absence of its binding partner, AP-1 (which is DAG/PKC dependent), has been shown to induce tolerance in T lymphocytes, and apoptosis in B cell lines (71, 72). Thus, it is possible that induction of NFAT following BCR engagement may promote apoptosis specifically in T1 B cells.

In contrast to T2 B cells, T1 B cell survival and differentiation into T2 B cells likely occurs independently of DAG. Consistent with this hypothesis, the atypical DAG/Ca²⁺-insensitive PKCλ in pre-B cells mediates NF-κB activity through Src family protein tyrosine kinases including Fyn, Lyn, and Blk independently of Btk, PLC-γ2, Vav, or CD19 (73). Moreover, T1 B cell death may occur by an active process that involves production of ceramide, which triggers a proapoptotic program as observed in the immature B cell line WEHI 231 (74). Additionally, a recent report indicated that cholesterol levels are similar in T1 relative to T2 immature B cells, but lower than that of mature B cells (75). This difference correlates with developmental differences in surface BCR compartmentalization and signaling, underscoring the importance of the quality and quantity of lipid species in splenic B cell subsets.

Regardless of the specific lipid-mediated mechanisms that control T1 vs T2 B cell fate, we postulate that mobilization of calcium without the balancing DAG-mediated signals may promote apoptosis in T1 B cells, whereas balanced calcium and DAG-mediated signals may promote T2 B cell survival and differentiation into mature FoB cells (Fig. 7).

We thank Dr. Yoshio Hamada (Tissue and Cell Culture Laboratory, National Institute for Basic Biology, Japan) for the generous gift of notch2^+/− mice (provided by James Thomas, Vanderbilt University, Nashville, TN), and the Vanderbilt University Medical Center Flow Cytometry Core (Jim Higginbotham and Michele Nadef) for assistance with FACS sorting and calcium mobilization experiments.

The authors have no financial conflict of interest.