Cyclin D2 is essential for BCR-mediated proliferation and CD5 B cell development

Nanette Solvason1,7, Wei Wei Wu1, David Parry2, Daniel Mahony2, Eric W.-F. Lam3, Janet Glassford3, Gerry G. B. Klaus4, Piotr Sicinski5,6, Robert Weinberg6, Yong Jun Liu1, Maureen Howard1,7 and Emma Lees2

1 Department of Immunology and
2 Department of Cell Signaling, DNAX Research Institute, Palo Alto, CA 94304, USA
3 Ludwig Institute for Cancer Research and Virology and Cell Biology Section, Imperial College School of Medicine at St Mary's, Norfolk Place, London W2, UK
4 National Institute for Medical Research, Mill Hill, London NW7, UK
5 Department of Cancer Biology and Department of Pathology, Dana-Farber Cancer Institute, Harvard Medical School, Boston, MA 02115, USA
6 Whitehead Institute, MIT, Cambridge, MA 02142, USA

Correspondence to: E. Lees


    Abstract
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Progression into G1 in B lymphocytes is regulated by cyclins D2 and D3, components of the cell cycle machinery currently believed to have overlapping and potentially redundant roles in cell cycle control. To study the specific role of cyclin D2 in B lymphocyte proliferation, we examined B cells from cyclin D2–/– mice and demonstrate a specific requirement for cyclin D2 in BCR- but not CD40- or lipopolysaccharide-induced proliferation. Furthermore, conventional B cell development proceeds normally in the mutant mice; however, the CD5 B cell compartment is dramatically reduced, suggesting that cyclin D2 is important in CD5 B cell development as well as antigen-dependent B cell clonal expansion.

Keywords: anti-Ig, B1 B cells, B-CLL, cell cycle, chronic lymphocytic leukemia


    Introduction
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 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
The commitment to enter S phase and divide or remain in the G1 phase with an unduplicated genome takes place in mid-to-late G1 and is referred to as the restriction point (R) (13). Operationally, the restriction point separates the cell cycle into an initial mitogen-dependent period followed by a growth factor-independent stage (13). The D-type cyclins are the first components of the cell cycle machinery to become induced in response to mitogen stimulation (13), and their associated kinase activity has a critical role in the commitment to enter S phase by phosphorylating and thereby inactivating pRb, a negative regulator of cell cycle progression (3). Consistent with their major role in the positive regulation of G1, the D-type cyclins are required for S phase entry, and their overexpression both accelerates G1 and reduces the dependence on exogenous growth factors (4).

The three D-type cyclins, D1, D2 and D3, are expressed in proliferating cells in a lineage-specific manner with at least two different D-type cyclins being expressed in most cell types. Results from mice homologous for genetic disruptions in cyclins D1 or D2 suggest that D cyclin function is overlapping and potentially redundant since viable offspring were obtained in each case (5,6). More direct evidence for overlapping roles of D cyclins in proliferation is the ability of cyclins D1, D2 or D3 to restore a normal cycle progression time to a cyclin D1-deficient subline of DT40 (7). However, there are specific lines of evidence in certain cell types suggesting that an individual D-type cyclin has distinct biological roles during certain cell cycles (8,9) and that D-type cyclins may have specific roles in cellular processes other than proliferation (1013).

Our laboratory has demonstrated that proliferating B lymphocytes express both cyclin D2 and D3 but not D1 (14); however, it is the accumulation of cyclin D2 which has been suggested to be the critical event in passage through the restriction point (15). Consistent with this idea, stimulation of B cells with complete versus partial activation signals suggests that signals responsible for moving B cells through the restriction point impact at the level of cyclin D2 protein accumulation (14,16).

Further evidence suggesting discrete roles for D-type cyclins in B cells comes from studies of cyclin expression in B leukemia and lymphomas representative of different B cell subsets. Examination of human B cell-derived lines revealed specific phenotype-related differences in D-type cyclin expression (17,18). Furthermore, centrocytic lymphoma, representative of mantle zone B cells, is characterized by a t(11:14) translocation which results in the constitutive expression of cyclin D1, a cyclin which is not normally detected in proliferating mouse or human B cells (19,20). In contrast, cyclin D2 but not D1 or D3 was overexpressed in chronic lymphocytic leukemia (CLL), which is thought to be the transformed counterpart of the normal CD5-expressing B cells (21,22). These results are consistent with the idea that there may be specific roles for individual D-type cyclins during different B cell developmental stages in addition to specific roles in proliferation.

To sort out the individual role of cyclin D2 in B lymphoid cells we utilized an experimental system which differentially regulates D cyclin induction during G1 and S phase entry. The model system consists of stimulating normal resting B cells with a low dose of anti-Ig to mimic weak BCR cross-linking in the presence of IL-4. Although these two stimuli when used singly stimulate entry into G1 but not S, the combination of IL-4 plus low-dose anti-Ig promotes S phase entry and cell cycle completion. Consistent with previous reports, our results implicated the induction of cyclin D2 as the primary cyclin involved in the passage through R during BCR-mediated B cell proliferation. Furthermore, the requirement for cyclin D2 during S phase entry was confirmed by analysis of B cells from mice homologous for a genetic deletion of cyclin D2. Indeed, cyclin D2–/– B cells failed to proliferate in response to IL-4 plus low-dose anti-Ig. Additionally, analysis of B cell development in the mutant mice revealed a specific diminution in the CD5 B cell compartment while conventional B cell development appeared normal. These results demonstrate unique and non-redundant roles for cyclin D2 during proliferation and development in normal mouse B lymphoid cells.


    Methods
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Mice
BALB/c mice (6–8 weeks old) were obtained from Simonsen and were used as a source of B cells for initial stimulation with IL-4, low-dose anti-Ig and IL-4 plus low-dose anti-Ig (Fig. 1Go). The derivation of the cyclin D2–/– mice and the screening protocol for heterozygous identification have been previously described (5). Cyclin D2 mutation is carried on a mixed background of 129S/v and C57BL6. Mice used in experiments described here were fourth generation. Control mice used for all experiments (except Fig. 1Go) were the heterozygote littermate controls.



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Fig. 1. Induction of cyclin D2 and D3 in normal B lymphocytes. Time points were collected from normal BALB/c splenic B cells following culture with 100 U/ml IL-4, 1 µg/ml polyclonal F(ab')2 anti-IgM or the combination of 1 µg/ml polyclonal F(ab')2 anti-IgM plus IL-4 stimulation. Sequential screening of Western blots revealed expression levels of cyclin D2 (A) and D3 (B) in response to each experimental condition. Lysate from a mouse myeloma cell line, P3X, was used as a positive control. In (C) blots were screened with antibodies against ERK2 to demonstrate equal protein loading.

 
Reagents
Antibodies against cyclins D2, D3 and ERK2 were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). Horseradish peroxidase (HRP)-conjugated reagents were purchased from Amersham (Arlington Heights, IL). Fluorescent-conjugated antibodies to IgM, B220, IgD, CD5, CD21, CD19 and CD23 were purchased from PharMingen (San Diego, CA). HRP-conjugated anti-rat, anti-rabbit and anti-mouse secondary reagents were purchased from Amersham. The affinity-purified F(ab')2 goat anti-mouse IgM used to stimulated B cells was purchased from Cappel (Durham, NC). The rat anti-mouse monoclonal anti-CD40 antibody was described previously.

FACS analysis of CD5 B cell population
The criteria for identifying CD5-expressing B cells was as follows. CD5 B cells were identified in the normal control littermates by first staining with the combination of anti-IgM–FITC plus a phycoerythrin (PE)-conjugated isotype control. A second aliquot of cells was stained with anti-IgM–FITC plus anti-CD5–PE. The isotype control profile was used to set the horizontal bar for the calculation of the percentage of CD5-expressing cells in the second aliquot of cells. Peritoneal cells from the mutant mice were stained using the same combination of antibodies and the settings for CD5 positivity were again obtained from the results from the isotype control antibody.

Purification of splenic B cells
To isolate splenic B cells, single-cell suspension were prepared from spleens of unprimed mice. Red blood cells were lysed using red blood cell lysis buffer (Sigma, Irvine, CA) according to the manufacturer's instructions. To deplete T cells, splenocytes were stained using anti-Thy-1 (NEN Research Products, Wilmington, DE) and anti-CD4 (RL172), followed by incubation in rabbit complement (Cedarlane, Hornby, Ontario, Canada) at 37°C for 45 min. These cells were consistently >90% B220+IgM+, and included both the low- and high-density B cells. Mice referred to as wild-type in the text are the heterozygous littermate control mice (–/+).

In vitro stimulations and whole-cell lysate preparation
The expression of cell cycle regulatory proteins in stimulated B cell populations was evaluated using procedures described previously (23). Briefly, 50 ml cultures were set up at 106 B cells/ml in 75 cm2 flasks (Becton Dickinson, Lincoln Park, NJ) and stimulated with 100 U/ml IL-4, 1 µg/ml F(ab')2 anti-IgM (low-dose anti-Ig) or IL-4 plus 1 µg/ml F(ab')2 anti-IgM (low-dose anti-Ig). Cells were collected at times indicated post-stimulation. All stimulations were carried out in supplemented RPMI containing 10% FCS (JR Scientific, Woodland, CA), 5x10–5 M 2-mercaptoethanol (Polysciences, Warrington, PA), 2 mM glutamate (JR Scientific), 10 mM HEPES buffer (Irvine Scientific, Santa Ana, CA), 100 U/ml penicillin and 100 µg/ml streptomycin (Irvine Scientific). Following stimulation, cells were recovered, washed 3 times in cold PBS, divided into three pellets, and stored at —80°C for later use in Western blotting and in vitro kinase assays. When all time points were collected, one pellet from each time point was lysed in NP-40 lysis buffer (1% NP-40, 250 mM NaCl, 1 mM HEPES, pH 7.5 and 1 mM DTT; USB, Cleveland, OH) with protease inhibitors added [final concentration: 5 µg/ml aprotinin (Sigma), 125 µg/ml Pefabloc (Boehringer Mannheim, Minneapolis, MN) and 5 µg/ml pepstatin (Sigma)]. Protein concentrations were calculated using a BioRad protein assay kit (BioRad, Hercules, CA) according to the manufacturer's procedures. An equal volume of 2xSDS sample buffer (Novex, San Diego, CA) with 2-mercaptoethanol was added to the total cell lysates. Then 125 µg of lysate was run per time point on 12% Novex reducing gels according to the manufacturer's suggested protocol and transferred onto Immobilon PVDF membranes (Millipore, Bedford, MA) at 30 V overnight in 4°C.

Thymidine incorporation
Aliquots of 200 µl (2x105 total cells ) of stimulated cells were placed in 96-well flat-bottom plates at times indicated and pulsed for 4 h in 1 µCi [3H]thymidine (Amersham). Cells were harvested and incorporated c.p.m. counted using a PHD cell harvester (Cambridge Technology, Cambridge, MA).

Testing of serum IgM and IgG3 levels by ELISA
Sera was collected from the mutant and heterozygote littermate control mice. ELISA was used to quantitate the levels of individual isotypes levels in the mice according to protocols described elsewhere (24).

Purification of peritoneal CD5 B cells and conventional splenic B cells
CD5 B cells were purified from the peritoneal cavity of heterozygous littermate control (+/–) mice by sorting with monomeric F(ab') anti-IgM (Southern Biotechnology Associates, Birmingham, AL) in combination with anti-Mac1. The sorted peritoneal cell population expressed IgMhiMac1lo which are hallmarks characteristics of the CD5 B cell population. In parallel, total splenic B cells were purified by the same method and the IgM+Mac1 cells were collected for comparison. To ensure that the monomeric anti-Ig could not induce cross-linking and subsequent activation of peritoneal B lymphoid cells during the sorting procedure, the monomeric preparation of anti-IgM was tested for its ability to induce splenic B cell proliferation in vitro, a property which is known to require cross-linking of the BCR. The F(ab') anti-IgM did not induce splenic B cell proliferation (not shown) nor did it stimulate the up-regulation of either cyclin D2 or D3 induction in splenic B cells (see Fig. 5Go), confirming that there was no signal to cell cycle machinery generated at the BCR by the sorting reagent.



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Fig. 5. Expression of cyclin D2 and D3 in peritoneal CD5 B cells and conventional splenic B cells. Lysates were prepared from (1) mouse myeloma, P3X, (2) unstimulated peritoneal CD5-expressing B cells (prepared as described in Methods) (3) CD5 B cells stimulated for 24 h with 1 µg/ml polyclonal F(ab')2 anti-IgM plus 100 U/ml IL-4 or (4) unstimulated splenic B cells. Total protein (100 µg) from each sample was used for the Western blotting. Blots were sequentially probed with antibodies against cyclin D2 and D3 as described in Fig. 1Go.

 

    Results
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Up-regulation of cyclin D2 in splenic B cells after activation with IL-4 and low-dose anti-Ig
To study the induction of D cyclins during cell cycle progression in B lymphocytes, we stimulated resting splenic B cells with either IL-4 alone, weak BCR cross-linking (mimicked by a low dose of anti-Ig, 1 µg/ml) alone or the mitogenic combination of IL-4 plus low-dose anti-Ig. Time points were collected at 0, 6, 12, 24, 36 and 48 h post-stimulation. Lysates were prepared and 100 µg of total protein from each time point was loaded on a gel. Western blotting was performed as described previously (23) and the blots were sequentially screened with mAb to cyclin D2, D3 and D1. Results shown in (Fig. 1aGo, top) demonstrate that IL-4 stimulation resulted in a dramatic induction of cyclin D2 6 h after stimulation but rapidly diminished thereafter. Cyclin D2 was induced at very low levels in response to low-dose anti-Ig stimulation and this level of induction was maintained throughout the time course (Fig. 1aGo, middle). In contrast, stimulation of B cells with the mitogenic combination of IL-4 plus low-dose anti-Ig resulted in the dramatic and sustained induction of cyclin D2 (Fig. 1aGo, bottom). Cyclin D3 induction was similar between the three stimuli (Fig. 1bGo). To confirm that an equal amount of protein was loaded onto each well, the blots were probed with an antibody to ERK2 which is constitutively expressed under each of the stimulation conditions. Figure 1(c)Go confirms that equal protein was loaded in each well. Furthermore, consistent with previous reports (23,25), cyclin D1 was not detected under any experimental conditions (not shown).

B cells from cyclin D2–/– mice have an impaired proliferation response to IL-4 plus anti-Ig, anti-IgM or anti-IgD
Since the difference in D cyclin induction between all three stimulants was primarily at the level of cyclin D2, we examined B cell development and activation in cyclin D2–/ mice (5). B cells from the heterozygous littermate control (+/–) mice proliferated in response to low-dose anti-Ig plus IL-4 which peaked at 48 h post-stimulation (Fig. 2aGo, squares). However, B cells from the mutant mice did not proliferate (Fig. 2aGo, circles). Interestingly, stimulation of mutant B cells through the BCR did not result in an increase in cell death, but rather a specific failure to enter the cell cycle (data not shown). To study the proliferation defect further, we purified B cells from the mutant and wild-type mice, and stimulated them in vitro with different anti-Ig reagents and monitored the entry into S phase with [3H]thymidine incorporation. A dramatic unresponsiveness was demonstrated using agonistic mAb directed against IgM (B7-6) (26) or IgD to cross-link the BCR. Using these reagents, the mutant B cell proliferative response was depressed by 90 and 80% respectively when compared to wild-type B cells (Fig. 2bGo). When mutant B cells were stimulated across a broad range of concentrations of polyclonal anti-Ig, we detected a proliferative response which approached 30% of the wild-type response at high concentrations of anti-Ig (Fig. 2cGo). We performed Western blots on B cells stimulated with a high dose of anti-Ig and confirm that the proliferation occurred in the absence of cyclin D2 induction (Fig. 2dGo) and furthermore that the induction of cyclin D3 was similar between the mutant wild-type B cells (not shown).



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Fig. 2. Proliferation defects in cyclin D2–/– mice. (A) B cells from heterozygous littermate control (+/–) ({blacksquare}) or mutant (–/–) ({circ}) mice were treated with 1 µg/ml polyclonal F(ab')2 anti-IgM plus IL-4 at concentrations described in Fig. 1Go. [3H]Thymidine incorporation was used to monitor S phase entry at 48 h post-stimulation. (B) B cells from wild-type (black bar) and mutant (dotted bar) mice were stimulated with 10 µg/ml of monoclonal anti-IgM or 10 µg/ml monoclonal anti-IgD (C). B cells from wild-type ({square}) or mutant (•) mice were treated with polyclonal F(ab')2 anti-IgM across a broad concentration range in the absence of IL-4. [3H]Thymidine incorporation was used to monitor S phase entry at 48 h post-stimulation. (D) Cyclin D2 expression was analyzed in wild-type and mutant B cells stimulated with 10 µg/ml anti-IgM. (E) B cells from wild-type (black bar) and mutant (dotted bar) mice were stimulated with 50 µg/ml LPS or 10 µg/ml monoclonal anti-CD40. Results are representative of three separate experiments.

 
Cyclin D2–/– B cells respond normally to lipopolysaccharide (LPS) and antibody to CD40
We next tested non-BCR mitogens and demonstrated that the mutant B cell proliferative response to LPS (Fig. 2eGo), a non-specific polyclonal B cell mitogen, was only minimally affected by the loss of cyclin D2. Similarly, the proliferative response in the mutant B cells following stimulation through CD40, a signal known to induce long-term proliferation in B lymphocytes in vitro, was intact, although diminished by 35% (Fig. 2eGo).

Cyclin D2–/– mice have decreased serum IgG3 and IgA
Since B lymphocytes with mutations in cyclin D2 exhibit defects in proliferation in vitro, we analyzed the serum from the mutant mice to determine if there was evidence of functional defects in the peripheral B cell pool. Serum was collected from six mutant (Fig. 3Go, closed circles) and six heterozygous littermate control (Fig. 3Go, open boxes) mice and compared by ELISA to determine the level of each Ig isotype in an unstimulated animal. Mutant mice expressed normal levels of serum IgM, IgG1, IgG2a, IgG2b and IgE; however, there was a statistically significant decrease in the levels of IgG3 (P = 0.016) and a less dramatic decrease in the levels of IgA (P = 0.066).



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Fig. 3. Serum Ig levels in cyclin D2—/— mice. Serum isotype levels were determined in the heterozygote littermate control ({square}) and mutant (•) mice by ELISA (23). Statistical significance was determined by Mann–Whitney test (*P = 0.016; **P = 0.066).

 
Cyclin D2–/– mice have reduced CD5+ B cell compartment
Although there have been no previously reported immune deficiencies in the cyclin D2–/– mice (5), we were interested to determine if B cell development was affected by a loss of cyclin D2. Since the CD5 B cell population is known to be a major source of both IgG3 and IgA and we detected substantial deficits in these serum isotypes, we examined the mutant mice to determine if this B cell subset was present. Examination of the lymphoid compartment in the peritoneal cavity revealed a dramatic reduction in CD5 B cells in the mutant mice. Figure 4Go shows FACS profiles of heterozygote littermate (+/–) and mutant (–/–) peritoneal cells stained with a panel of mAb which identify the CD5 B cell population. In Fig. 4(a)Go, the CD5-expressing B cells are evident in the wild-type (top, see box), but severely diminished in the mutant mouse (bottom, box). In Fig. 4(b)Go, the characteristic IgMhiIgDlo population which contains the CD5 B cell subset is again present in wild-type but diminished in the mutant mouse. Indeed, our analysis revealed a statistically significant decrease in both the proportion (P = 0.002) and total number (P = 0.005) of CD5-expressing B cells detected in the mutant mice (Table 1Go).



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Fig. 4. Flow cytometric analysis of CD5 B cells in cyclin D2–/– mice. Peritoneal cells were stained with (A) PE–anti-CD5 plus FITC–anti-IgM or (B) PE–anti IgD plus FITC–anti-IgM. In (A) top and bottom, the box indicates the position of the CD5-expressing B cells. In (B) top and bottom, the box indicates the position of the IgMhiIgDlo-expressing B cells which contain both the CD5 B cell population and the CD5 `sister' population. The arrow shows position of conventional B cells (IgMloIgDhi). The compilation of statistical analysis of numbers and proportions of B cells in the individual B cell subsets is shown in Table 1Go.

 

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Table 1. Number and phenotype of B lymphoid cells in peripheral lymphoid organs of mutant and wild-type mice
 
Interestingly, we noted that the conventional B cell compartment (IgMloIgDhi) was intact in the peritoneal cavity of both the wild-type and mutant mouse (Fig. 4bGo, arrow) suggesting that B cell development in the bone marrow was not affected by loss of cyclin D2. Indeed, we detected no significant difference in the proportion or total number of B cell precursors in the bone marrow of mutant and wild-type mice (Table 1Go). Furthermore, there was no difference in the ratio of pre-B:B cells or in the level of expression of B220 or sIgM (Table 1Go). Additionally, the number and phenotype of B cells in the spleen including the marginal zone and follicular B cells were comparable between the mutant and wild-type mice (Table 1Go). The failure of the CD5 B cell subset to develop while the conventional B cell compartment is unaffected by the loss of cyclin D2 highlights a specific requirement for cyclin D2 during the genesis of this specific B cell subset.

To further evaluate the unique requirement for cyclin D2 in development of CD5-expressing B cells but not conventional B cells, we isolated CD5 B cells from the peritoneal cavity and conventional B cells from spleen of wild-type mice. Western blots were prepared from lysates of unstimulated CD5 B cells, CD5 B cells stimulated with IL-4 plus low-dose anti-Ig or unstimulated total splenic B cells (Fig. 5Go). Cyclin D2 was expressed at high levels in the unstimulated CD5 B cell population and, interestingly, the level of cyclin D2 expression did not increase following anti-Ig plus IL-4 stimulation in this B cell subset. Cyclin D2 was weakly detectable in the unstimulated total splenic B cells. The analysis of cyclin D3 revealed that the slower migrating D3 form is detected in both activated and non-activated CD5 B cell populations while both forms of D3 are detectable in the total splenic B cell population. Neither the functional significance nor biochemical modifications associated with the different migrating forms of cyclin D3 are known. Taken together these results confirm a differential expression of cyclin D2 and D3 in the normal peritoneal CD5 population. The unexpected expression of high levels of the G1-promoting cyclin, D2, in resting CD5 B cells is consistent with the suggestion that CD5 B cells, in contrast to splenic B cells, are `partially activated' and, furthermore, appear to be arrested in the G1 cell cycle phase (27).


    Discussion
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
IL-4 or low-dose anti-Ig can promote B cell cycle progression to G1; however, the combination of the two promotes B cell cycle progression into S phase. Our results demonstrate that IL-4 plus anti-Ig, IL-4 alone or anti-Ig alone induce comparable levels of cyclin D3. However, IL-4 plus anti-Ig but not IL-4 alone or anti-Ig alone induce cyclin D2, suggesting that in this model system, cyclin D2 is the S phase-promoting factor. Direct evidence for a non-redundant role for cyclin D2 in G1/S in our system is demonstrated by the dramatic loss of proliferative capacity of B cells from cyclin D2–/– mice in response to IL-4 plus low-dose anti-Ig. These results are consistent with a specific role for cyclin D2 in the commitment to enter S phase while the ability to enter G1 may be independent of cyclin D2.

The role for cyclin D2 in commitment to enter S phase is, however, most evident in BCR-induced proliferation. Indeed, the B cell proliferative response induced by LPS or CD40 are only minimally affected by loss of cyclin D2. This suggests that the cyclin D2 dependency of BCR-mediated B cell proliferation may be a unique characteristic of antigen receptor signaling rather than a strict role of cyclin D2 in S phase entry in B lymphoid cells. A functional unresponsiveness of the BCR signaling pathway was demonstrated in three experimental systems: (i) low-dose anti-Ig plus IL-4, (ii) optimal concentrations of a monoclonal anti-IgM antibody preparation and (iii) optimal concentrations of a monoclonal anti-IgD antibody preparation. It is important to note, however, that stimulation with a high dose of the polyclonal anti-Ig can induce some proliferation in the mutant B cells.

The BCR dependency on cyclin D2 appears restricted to the mature B cell stage. Indeed, B cell development in the bone marrow is thought to require signaling through the pre-BCR which leads to expansion of B cell precursors during certain developmental stages (28). Given this, we were surprised that B cell development proceeded normally in the mutant mice. Our demonstration that the pre-B cell compartment in the mutant mice is comparable to that in the wild-type suggests that pre-BCR-induced proliferation is not dependent on cyclin D2, but may instead require induction of cyclin D3. Indeed, in our analysis of B cell lines from different developmental stages, the D3-expressing B cell lines were clustered in the pro- and pre-B cell stages where the pre-BCR would be expressed (data not shown). Furthermore, it has been demonstrated that IL-7, a potent growth factor which promotes the development of precursor B cells, induces active kinase complexes with cyclin D3 and not D2 (29). Additionally, cyclin D3 but not D2 has been detected in hematopoietic cells of the fetal liver, a major site of B cell development in the fetus. Together these results suggest that expansion of B cell precursors during conventional B cell development is independent of cyclin D2 and may instead require cyclin D3.

Despite the apparently normal development of the conventional B cell compartment, there was a dramatic decrease in the number and proportions of CD5 B cell compartment in the peritoneal cavity. The co-existence of a defect in BCR signaling and CD5 B cell development has previously been detected in mutant mice in which either natural or genetically engineered defects in various components of the BCR signaling pathways have been disrupted (3033). These data argue for an important role for BCR signaling in the genesis of CD5 B cells. The developmental origins of CD5 B cells are controversial (27,34,35), but our data are the most consistent with the hypothesis put forward by Wortis et al. (36). In an in vitro system developed in his laboratory, CD5 B cells could be generated from conventional splenic B cells by stimulation through the antigen receptor with anti-Ig reagents. This experimental manipulation causes the cells to undergo a single round of cell cycle division prior to CD5 expression. Intriguingly, stimulation through CD40 or LPS, both of which cause B cell proliferation even in the absence of cyclin D2, was incapable of inducing the expression of CD5 on conventional B cells. Thus, our data further the observations by Wortis by demonstrating a dependence on cyclin D2 for this unique function of the BCR.

In vivo, CD5 B cells have a dual role in clearing certain bacterial pathogens (37,38). These B cells switch to and secrete antibodies of the IgG3 subtype which is known to be a major component of the response to bacterial polysaccharide antigens (39). Additionally, CD5 B cells contain precursors of IgA-secreting plasma cells in the gut which also contribute dramatically to the host defense against bacterial pathogens (40). Indeed, in the cyclin D2–/– mice we noted a specific immunodeficiency in IgG3 as well as IgA levels. We believe these immunodeficiencies are secondary to the defect in CD5 B cell development rather than the result of a defect in the general ability of B cells to switch to or secrete these specific Ig isotypes.

It is of interest that CLL, the most prevalent leukemia in the western hemisphere, is considered the transformed counterpart of normal CD5 B cells (41,42). Significantly, examination of these leukemias for D cyclin expression has demonstrated overexpression of cyclin D2 but not D3 or D1. Extending these reports, we demonstrate here that freshly isolated CD5 B cells express high levels of cyclin D2 even in the absence of mitogenic stimulation. The constitutive expression of this G1/S phase-promoting cyclin in CD5 B cells may contribute to the unusual characteristics of these cells. Indeed, normal CD5 B cells are larger than conventional B cells and appear to be in the G1 phase rather than G0. Furthermore, the well-documented self-replenishing ability of CD5 B cells may depend on long-term expression of cyclin D2. The specific requirement for cyclin D2 during CD5 B cell development may lead, in turn, to an increased susceptibility of these B cells to oncogenic transformation in response to deregulated expression of this cyclin.


    Acknowledgments
 
DNAX Research Institute is fully funded by Schering Plough.


    Abbreviations
 
CLL chronic lymphocytic leukemia
HRP horseradish peroxidase
LPS lipopolysaccharide
PE phycoerythrin

    Notes
 
7 Present address: Corixa Corp., Redwood City, CA 94063, USA Back

Transmitting editor: J. Kearney

Received 5 November 1999, accepted 18 January 1999.


    References
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 Abstract
 Introduction
 Methods
 Results
 Discussion
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