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The Journal of Immunology, 2007, 179, 7544 -7552
Copyright © 2007 by The American Association of Immunologists, Inc.
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Transitional B Cells Lose Their Ability to Receptor Edit but Retain Their Potential for Positive and Negative Selection1

Hongsheng Wang2,*, Jianxun Feng*, Chen-Feng Qi*, Zhaoyang Li*, Herbert C. Morse, III* and Stephen H. Clarke2,{dagger}

* Laboratory of Immunopathology, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Rockville, MD 20852; and {dagger} Department of Microbiology and Immunology, University of North Carolina at Chapel Hill, Chapel Hill, NC 27599


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Ligation of B cell receptors on immature bone marrow B cells, either by an endogenous Ag or by an anti-B cell receptor Ab induces secondary V(D)J gene rearrangements, termed receptor editing. Whether the same signal induces receptor editing in transitional B cells is not clear. In this study, we examined the responses of immature and transitional B cells from VH12V{kappa}1A Ig transgenic mice to stimulation with an anti-Igβ Ab. Our results demonstrated that immature B cells stimulated with a low concentration of anti-Igβ Ab, mimicking Ag stimulation, underwent receptor editing both in vivo and in vitro, as evidenced by the detection of dsDNA breaks at J{kappa} recombination signal sequences, whereas transitional B cells did not. The lack of dsDNA breaks in transitional B cells contrasts with their increased expression of RAG1 and RAG2, suggesting a novel mechanism that may prevent rearrangements. Furthermore, treatment of transitional B cells with high concentrations of anti-Igβ Abs induced apoptosis, whereas low concentrations induced differentiation. Our results support the idea that transitional B cells lose the capacity to edit, but are sensitive to positive and negative selection.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
B cell development in bone marrow (BM)3 follows ordered cellular stages and proceeds through sequential rearrangement of Ig H and L chain genes. The recombinases, RAG1 and RAG2, first mediate H chain gene rearrangement at the pro-B cell stage to produce a functional µH chain and then mediate L chain gene rearrangement at the small pre-BII cell stage to produce a {kappa} or {lambda} L chain. H and L chains assemble with the Ig{alpha}-Igβ heterodimer to form a B cell Ag receptor (BCR) complex on the cell surface. BCR+ immature B cells are immediately subject to Ag-mediated negative selection events including apoptosis (central deletion), anergy, or secondary gene rearrangement (receptor editing) to eliminate harmful autoreactive clones. Recent studies (1, 2, 3) suggest that receptor editing is the predominant mechanism of negative selection. For cells expressing autoreactive BCRs, encounter with Ag triggers a BCR signal leading to a 2-h delay in development and increased expression of Rag1 and Rag2 genes (1). Subsequent cleavage at the recombination signal sequence (RSS) of L chain loci results in rearrangement of a new L chain, which assembles with the H chain to form a new BCR with reduced autoreactivity. Only those immature B cells that succeed in passing these tests of negative selection exit the BM and continue to differentiate in the periphery.

In the periphery, the newly immigrated immature B cells, termed transitional 1 (T1) cells, pass through phenotypically distinct transitional stages before they are selected into one of the three mature B cell pools: follicular (FO) B2, marginal zone (MZ), and B1 B subsets (4, 5). A ligand-independent BCR tonic signal is essential for B cell survival and maturation from the transitional to the FO stage (6). Tonic signals are affected by the level of BCR expression, the activity of accessory molecules, such as CD19 and CD22, and secondary messengers of the BCR signaling pathway (revewed in Ref. 7). Changes in the expression or function of these molecules are found to affect the development of mature B cells (7). In contrast to FO B2 cell differentiation, MZ, and B1 B cell differentiation requires a ligand-dependent signal (8). The strength of this signal is critical, because strong ligand-mediated BCR signals can induce negative selection such as anergy or deletion (9, 10). Whether receptor editing in transitional B cells normally occurs is uncertain. Although receptor editing in mature and germinal center B cells is well documented, these studies were conducted under special circumstances, such as immunization, ongoing autoimmune responses, or in cells with mutant BCR signaling molecules (11, 12, 13, 14, 15). This type of peripheral receptor editing, also known as "receptor revision", is purported to generate high-affinity Abs rather than to silence autoreactive B cell clones.

We have generated VH12V{kappa}1A H and L chain transgenic (Tg) mice (designated 6–1/V{kappa}1A) to study Ag-mediated positive and negative selection mechanisms (16, 17). B cells expressing the 6–1/V{kappa}1A BCR are developmentally blocked at the transitional 2 (T2) cell stage with no evidence of Ag-imposed negative regulation. These cells are short lived, express high levels of BCRs, proliferate, and secrete Abs normally in response to a variety of stimuli, and most importantly, differentiate into B2-like cells following treatment with a low dose anti-Igβ Ab in vivo (17). Furthermore, 6–1/V{kappa}1A transitional B cells are sensitive to cell death induced by a high-dose anti-Igβ Ab in vivo. We attributed this developmental arrest to a lack of a ligand-mediated positive selection signal.

In this report, we used low doses of anti-Igβ Ab to mimic Ag stimulation and compared receptor editing in immature and transitional B cells from 6–1/V{kappa}1A Tg and normal non-Tg mice. Our results demonstrated that in response to anti-Igβ stimulation, immature B cells underwent RAG-mediated receptor editing but transitional B cells did not, even though they increased RAG expression. Instead, transitional B cells underwent differentiation to a more mature B cell.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Mice

VH12 Tg (6–1), 6–1/V{kappa}1A, 6–1/V{kappa}1A/RAG1–/–, and 6–1/V{kappa}4 Tg mice have been previously described (17, 18). CB17 mice were purchased from Taconic Farms. All mice were kept under specific pathogen-free conditions. All animal protocols were approved by the University of North Carolina Institutional Animal Care and Use Committee and the National Institute of Allergy and Infectious Diseases Animal Care and Use Committee.

Abs and other reagents

mAbs for flow cytometry were purchased from BD Biosciences. Fluorescence-encapsulated liposomes was made previously (18). PMA was purchased from Sigma-Aldrich. Ionomycin was obtained from Molecular Probes (Invitrogen Life Technologies). Anti-Igβ Ab (clone HM79) was described previously (17).

Flow cytometry, cell purification, and treatment protocols

Bone marrow and splenic cells were stained with fluorochrome-labeled Abs and analyzed on a FACSCalibur (17) or Cyan ADP. Cells were sorted on a MoFlo high speed sorter (DakoCytomation) according to their phenotypes: pre-BII cells as B220lowIgM–CD43lo/–; immature B cells as B220lowIgM+; T1 cells as B220+IgMhighCD23–CD21–; T2 cells as B220+IgMhighCD23+CD21–; B2 cells as B220+IgMlowCD23+CD21int; pre-MZ cells as B220+IgMhighCD23+CD21high; MZ B cells as B220+IgMhighCD23–CD21high; B1int as B220+CD5+CD23+; B1 as B220+CD5+CD23– (5, 17, 19). The purity of sorted cells was >90%.

For in vitro culture experiments, BM and splenic B cells were purified by negative selection using a Mouse B cell Recovery Column kit (Cedarlane Laboratories) or magnetic-activated Dynal beads (Invitrogen Life Technologies). Cells were cultured with RPMI 1640 medium supplemented with 10% FBS and antibiotics. Stimulants, including anti-Igβ mAb (0.1–10 µg/ml), PMA (2 ng/ml), and ionomycin (2 ng/ml), were added to cultures for 24 h. For cell cycle analysis, the cells were fixed with 70% ethanol overnight, treated with RNase (100 µg/ml) plus propidium iodide (50 µg/ml) and analyzed by flow cytometry.

For in vivo treatment, 6–1/V{kappa}1A mice were injected i.p. with a single dose of 10 µg of anti-Igβ Ab or normal hamster IgG for 5 days. BM and spleen cells were prepared for cell sorting as described above.

Ligation-mediated (LM)-PCR

DNA was purified from cells using a DNeasy Tissue kit (Qiagen) according to the manufacturer’s instructions. The linker ligation and PCR conditions have been described previously (20, 21, 22). In brief, 2 µg of genomic DNA was ligated with the BW linker and 100 ng of ligated DNA was amplified for 12 cycles with the linker primer BW-1 and a locus-specific primer V{kappa}S, followed by an additional 27 cycles with BW-1 and a nested locus-specific primer. The PCR products were analyzed by electrophoresis in 1.2% agarose gels and transferred to a nylon membrane (Bio-Rad). The blots were hybridized with 32P-labeled locus-specific internal oligonucleotides and analyzed with the Storm Phosphorimager using ImageQuant software (Molecular Dynamics). To control for the amount of amplifiable DNA in each sample, 5 µl of ligated DNA was amplified for 27 cycles under the identical conditions with the primers CD14L and CD14R to amplify the CD14 gene.

Analysis of RAG expression

Quantification of Rag1 and Rag2 transcripts was performed by real time quantitative PCR (qPCR). Total RNA was isolated using a RNeasy Mini kit (Qiagen). Approximately 200 ng of total RNA in 20 µl was reverse-transcribed with SuperScript II reverse transcriptase and random hexamer primers (Invitrogen Life Technologies). The PCR was performed in an ABI Prism 7000 and 9000 Sequence Detection Systems with the SYBR Green PCR Master Mix reagents (Applied Biosystems). The cDNAs were serially diluted (50/10/5/1) to check the efficiency of the PCR to amplify Rag1, Rag2, and the house keeping gene hypoxanthine phosphoribosyltransferase (Hprt). PCR primer sequences were: Rag1 forward, 5'-GAAATTCAACACCCACAAATCAAA-3'; Rag1 reverse, 5'-TTCTAGGTAAGGTTTCCCCTCTGA-3'; Rag2 forward, 5'-CAGTCTTGCCAGGAGGAATCTC-3'; Rag2 reverse, 5'-ACAAGGCTGCAGACCATCCTT-3'; Hprt forward, 5'-GTTCTTTGCTGACCTGCTGGAT-3'; Hprt reverse, 5'-GTCCCCC-GTTGACTGATCAT-3'. Quantification was performed relative to the expression levels of non-Tg pre-B cells.

The expression of RAG2 proteins in anti-Igβ-stimulated B cells was detected by immunoblotting. In brief, proteins extracted from equal amount of cells were subjected for SDS-PAGE and transferred to PVDF membranes using standard procedures. The membranes were blocked with 4% BSA and probed with Abs against RAG2 (Santa Cruz Biotechnology), followed by HRP-conjugated secondary Ab. Immunoreactive proteins were detected by using the ECL kit (Amersham Biosciences). The films were scanned and the density of protein bands were quantitated by using Image J software (http://rsb.info.nih.gov/ij).

Chromatin immunoprecipitation (ChIP) assay

ChIP assays were performed using a ChIP assay kit purchased from Millipore. In brief, BM cells from 6–1/V{kappa}1A mice were cultured with IL-7 for 5 days followed by 18 h without IL-7. The cells were then harvested and cultured with and without 1 µg/ml anti-Igβ Ab for 18 h. Splenic cells from 6–1/V{kappa}1A mice were similarly cultured with 1 µg/ml anti-Igβ Ab for 18 h. The cells were then stained with Abs reacting with B220-PE plus IgMa-FITC or B220-PE plus liposome for BM and spleen cultures, respectively, and sorted for B220+IgM+ and B220+ liposome– B cells, respectively. The cells were cross-linked by addition of 1% formaldehyde for 15 min at room temperature. Cells were lysed in SDS lysis buffer and sonicated using an optimized condition to shear DNA to lengths between 200 and 1000 bp. The lysates were precleared with 80 µl of salmon sperm DNA/protein A agarose-50% slurry followed by incubation with anti-acetyl-histone H3 or anti-trimethy-histone H3-K4 Abs (Millipore). The immune complexes were recovered by salmon sperm DNA/protein A agarose and washed extensively. The DNA was recovered and real-time PCR was performed to amplify Jk5 and Tcra. The primers were: Jk5, forward, 5'-GGATCGGAGAATAAGCATGAGTAGTTA-3', reverse, 5'-TTCCTCATCCCCT-CCAAATCT-3'; Tcra constant region, forward, 5'-CGACTGCTGTTTGCCAAGAC-3', reverse, 5'-AGGGTGAAGCTGGGAATATGAA-3'.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
6–1/V{kappa}1A B cells arrest at a transitional B cell stage

In the BM of 6–1/V{kappa}1A Tg mice, almost all IgM+ B cells are B220low, a phenotype consistent with that of immature B cells (Fig. 1). In the spleen, ~10% of B cells were specific for phosphatidyl choline (PtC), detected using liposomes with membrane incorporated PtC. These cells expressed the B1 cell markers CD5 and CD43 ((16) and our unpublished data), suggesting that they are B1 cells. These PtC+ B cells expressed endogenous L chains, predominantly the V{kappa}4/5H gene required for PtC binding (16). Consistent with our previous report (17), the majority of the PtC– B cells were CD21–, CD23–/low, and IgMhigh, resembling a transitional B cell phenotype (Fig. 1). On a RAG1-deficient background, the splenic B cells of 6–1/V{kappa}1A mice were exclusively T1 and T2 cells (17). The lack of mature B cells in the periphery may account for the absence of mature recirculating B cells in the BM (Fig. 1). Thus, 6–1/V{kappa}1A B cells were unable to develop into mature B2 cells, but could become B1 cells through expression of an endogenous L chain.


Figure 1
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  		FIGURE 1. 6–1/V{kappa}1A mice lack mature B cells. Numbers indicate the percentage of B cells falling within each gate. Data shown are representative of more than eight independent analyses.

 
Expression of endogenous L chains by some B cells of 6–1/V{kappa}1A mice is indicative of incomplete L chain allelic exclusion. Similar "leakiness" of allelic exclusion has been observed in other L chain-targeted mice (23). To rule out Ag-induced secondary L chain gene rearrangement in 6–1/V{kappa}1A mice, we determined the levels of Rag1 and Rag2 transcripts and the generation of dsDNA break (DSB) at RSS of J{kappa} loci. As shown in Fig. 2A, DSBs were very rare in immature B cells of 6–1/V{kappa}1A mice when compared with immature B cells from VH12 H chain only Tg mice (designated 6–1) (Fig. 2A) or non-Tg mice (Fig. 2B). These results suggested that only a small number of immature B cells were undergoing endogenous V{kappa} rearrangement. Consistent with this, the levels of Rag transcripts in 6–1/V{kappa}1A immature B cells were not up-regulated compared with similar cells isolated from 6–1, 6–1/V{kappa}4, and non-Tg mice as controls (Fig. 2C). 6–1/V{kappa}4 B cells are PtC specific and are positively selected into the B-1 subset and used here as a negative control for endogenous L chain rearrangement and Rag gene expression. Taken together, these results suggested that the arrest of 6–1/V{kappa}1A B cell differentiation at the transitional stage was unlikely due to negative selection.


Figure 2
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  		FIGURE 2. Detection of DSB and RAG expression in B cell subpopulations of 6–1/V{kappa}1A and control mice. A, Evaluation of DSB at J{kappa} RSS in indicated populations by LM-PCR. CD14 levels were evaluated as a loading control. Data shown are representative of two independent experiments with sorted cells from three to five mice per group. Pre-BII and immature B cells were sorted from BM cells; PtC–CD23–, PtC–CD23+, T1, T2, B1int, and B1 cells were sorted from splenic cells. Similar results with unsorted cells were observed in four separate experiments. *, Prolonged exposure of J{kappa}2. B, Detection of DSB at J{kappa} RSS sites in B cell subsets from normal non-Tg mice. Pre-B, IgMhigh, and IgMlow cells were sorted from BM cells; T1, T2, pre-MZ, B2, and MZ cells were sorted from splenic cells. C, RAG1 and RAG2 expression in indicated populations was quantitated by qPCR. The sorted cells as in A were used to prepare RNA. Representative data from three separate experiments with multiple mice is shown.

 
Immature but not transitional B cells of 6–1/V{kappa}1A mice undergo receptor editing in response to low dose anti-Igβ stimulation

To determine whether transitional B cells can edit in response to a BCR signal, we stimulated immature and transitional B cells of 6–1/V{kappa}1A Tg mice with anti-Igβ Ab or PMA plus ionomycin (P.I.) for 24 h in vitro. PMA activates protein kinase C leading to activation of the MAP kinase cascade, a major signaling pathway downstream of the BCR, whereas ionomycin triggers a calcium signal that bypasses the BCR to induce receptor editing in immature B cells (24). Because RAG proteins are sensitive to cell cycling, low concentrations of PMA and ionomycin (1–5 ng/ml) were used to minimize their effect on cell cycle. Under these conditions, cell survival and growth were not changed (Fig. 3). Anti-Igβ Ab was also used at a low concentration (1 µg/ml) to maintain normal viability. Under these culture conditions, the viability of the cells was ~80% as determined by staining with propidium iodide (Fig. 3) or Annexin V (data not shown). LM-PCR analyses of 6–1/V{kappa}1A immature B cells revealed that anti-Igβ treatment induced an increase in DSB at J{kappa}2 and J{kappa}5, whereas P.I. induced an increase in J{kappa}5 DSB and a modest increase in J{kappa}2 DSB (Fig. 4A). Neither treatment had an effect on transitional B cells (Fig. 4A). Note that, although levels varied from mouse to mouse, a longer film exposure revealed that there were more DSB in untreated immature than in transitional B cells (Figs. 2A and 4A). We also observed that DSB were induced in immature B cells with as little as 0.1 µg/ml anti-Igβ Ab (data not shown). This result supports the idea that the signaling threshold for receptor editing is set very low in immature B cells. Surprisingly, anti-Igβ treatment of 6–1/V{kappa}1A immature B cells failed to stimulate RAG expression (Fig. 4, B and C). By contrast, both RAG2 mRNA and protein were slightly increased in anti-Igβ-treated transitional B cells (Fig. 4, B and C). Despite the absence of significant changes in RAG expression, RAG activity in anti-Igβ treated immature B cells was significantly increased as assessed by the production of RSS breaks. Although treatment with P.I. did not alter RAG expression in immature B cells, it induced a significant down-regulation of RAG expression in transitional B cells (Fig. 4B), indicating differential responses of RAG expression to P.I. stimulation between the cells of these two subsets. The up-regulation of RAG expression in 6–1/V{kappa}1A transitional B cells was not due to contamination with a small number of PtC+ B cells, because anti-Igβ stimulation of PtC+ B cells from VH12 H chain only Tg mice down-regulated RAG expression (data not shown). Thus, failure of transitional B cells to initiate RAG-mediated L chain gene rearrangement following stimulation with either anti-Igβ or P.I. suggests that they have lost the ability to undergo receptor editing.


Figure 3
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  		FIGURE 3. Cell cycle analysis of stimulated immature and transitional B cells. Purified BM immature and splenic transitional B cells from 6–1/V{kappa}1A mice were cultured with the indicated stimuli for 24 h. The cells were stained with propidium iodide to quantitate DNA content. The numbers are percentages of cells falling in each region used to define apoptotic (A), G0/G1, and S/G2/M cells. Data is representative of two independent experiments.

 

Figure 4
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  		FIGURE 4. Receptor editing is induced in BM immature B cells but not in splenic transitional B cells. A, Analysis of DSB in whole BM and purified splenic B cells from 6–1/V{kappa}1A mice. Where indicated, cells were stimulated with anti-Igβ Ab (1 µg/ml) or PMA plus ionomycin (P.I., 2 ng/ml each) for 24 h. Purified BM immature B cells exhibited similar results. B, Cells from the same experiment as in A were quantitated for RAG mRNA expression by qPCR. C, Western blot analysis of RAG2 protein expression in extracts from control and anti-Igβ-stimulated cells for 24 h. Purified BM and splenic B cells from 6–1/V{kappa}1A mice were used. β-actin levels were evaluated as a loading control. The bar chart summarizes protein densities of RAG2. The mean values are arbitrary units normalized by actin and the error bars are SD of two independent experiments. D, Detection of DSB at J{kappa} RSS in sorted B cell subpopulations from 6–1/V{kappa}1A mice treated with 10 µg of anti-Igβ Ab or a control Ab for 5 days. E, Analysis of J{kappa} RSS DSB in purified non-Tg transitional B cells stimulated with anti-Igβ Abs or P.I. as in B. F, Anti-Igβ induced RAG transcripts in transitional B cells of non-Tg mice. The cells were treated as in E. ND, Not done. All data are representative of two to six independent experiments with two to five mice per group.

 
To verify this result in vivo, we injected 6–1/V{kappa}1A mice i.p. with 10 µg of anti-Igβ Ab and tested for evidence of receptor editing in BM and splenic B cells. This dosage was previously shown to induce differentiation of transitional B cells to FO B cells in 6–1/V{kappa}1A/RAG1–/– mice (17). Five days after anti-Igβ treatment, a novel PtC–IgMlow CD23+ B cell population was detected in the spleen, in addition to the two native B cell populations, PtC–IgMhigh and PtC+IgMint (Fig. 5). This unique subset not only down-regulated surface IgM, but also gained characteristics of B2-like cells; they acquired a FO B cell phenotype (CD23+, CD21low, AA4.1–, CD24low, and CD40high (Fig. 5 and Ref. 17) and survived longer than control cells in vitro (17). For BM immature B cells, treatment with anti-Igβ induced a slight down-regulation of surface IgM and a reduction in the number of IgM+ cells (Fig. 5), which could be explained by a delayed generation of immature B cells, deletion in immature B cells, or migration of immature B cells to the spleen. The observation that treatment with anti-Igβ Ab at the higher concentration of 10 µg/ml did not induce apoptosis in vitro (Fig. 3) argues against induced deletion. However, 100 µg anti-Igβ Ab did induce deletion (17).


Figure 5
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  		FIGURE 5. B cell phenotypic changes following anti-Igβ treatment in vivo. 6–1/V{kappa}1A mice were injected with 10 µg of anti-Igβ Ab or a normal hamster IgG isotype control Ab for 5 days. BM and splenic cells were stained with liposomes and Abs against IgM, CD19, and CD23 and analyzed by flow cytometry. Numbers in top panel indicate the percentages of cells falling in each gate. The gates are also used for sorting. Data are representative of three mice in each group. Lip, Liposomes. MFI, mean fluorescence intensity.

 
To determine whether anti-Igβ induces DSB in transitional B cells, we sorted PtC–IgMlowCD23+ and PtC–IgMhigh B cells from the spleen and BM immature B cells, and measured DSB at J{kappa} RSS sites. As shown in Fig. 4D, anti-Igβ treatment induced DSB in immature B cells, but not in either subset of transitional B cells. We conclude that 10 µg of anti-Igβ Ab given in vivo induced transitional B cells to differentiate to the FO B cell stage, rather than to undergo receptor editing. Moreover, similar results with immature and transitional B cells were observed following treatment with a low affinity anti-Ig{kappa} Ab both in vivo and in vitro (our unpublished data). Thus, anti-Igβ simulated Ag stimulation induced receptor editing only in BM immature B cells.

Transitional B cells of non-Tg normal mice fail to edit in response to low dose anti-Igβ stimulation

To determine whether normal transitional B cells can be induced to edit, we purified and stimulated transitional B cells from non-Tg mice that had received sublethal irradiation (500 rads) 14 days earlier. Spleens of these mice are highly enriched for T1 and T2 cells (17, 25). Consistent with the findings with 6–1/V{kappa}1A transitional B cells, treatment of non-Tg transitional B cells with anti-Igβ Ab or P.I. did not induce an increase in DSB at J{kappa} RSS sites, although they did induce increased RAG expression (Fig. 4, E and F). We, therefore, conclude that both 6–1/V{kappa}1A and non-Tg transitional B cells are resistant to anti-Igβ-induced receptor editing, even using concentrations of anti-Igβ 10-fold higher than that required to induce receptor editing in immature B cells.

BM stromal cells do not support receptor editing in transitional B cells

Monroe and colleagues have shown that a Thy1dullDX5+ stromal BM cell population can redirect Ag-induced B cell apoptosis to receptor editing (26, 27). To determine whether BM stromal cells can promote anti-Igβ-induced receptor editing in 6–1/V{kappa}1A transitional B cells, we cocultured purified transitional B cells with BM cells isolated from RAG1–/– mice. Under these conditions, treatment with anti-Igβ Ab failed to induce DSB in transitional B cells, although it did induce DSB in immature B cells (Fig. 6A). However, the levels of RAG transcripts increased in both cell types (Fig. 6B). We obtained the same result with purified transitional B cells from sublethally irradiated normal mice (Fig. 6, C and D). The failure to induce editing in 6–1/V{kappa}1A and normal transitional B cells could not be ascribed to a loss of the Thy1dullDX5+ cell population, because it was readily identified by flow cytometry (Fig. 6E). The increased RAG expression in normal anti-Igβ-treated transitional B cells was consistent with previous reports using anti-IgM Abs (26, 27). However, unlike anti-Igβ, anti-IgM did not induce DSB unless cells were prevented from undergoing apoptosis (26). Based on propidium iodide staining, a concentration of 1 µg/ml (used in this study) or 10 µg/ml anti-Igβ Ab did not cause apoptosis (Fig. 3), whereas 10 µg/ml anti-IgM Ab readily induced apoptosis (Fig. 3). We conclude that anti-Igβ-stimulated transitional B cells are unable to undergo efficient L chain receptor editing, even in the presence of BM stromal cells.


Figure 6
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  		FIGURE 6. BM stromal cells were unable to promote receptor editing in anti-Igβ-treated transitional B cells. A, Detection of DSB at J{kappa} RSS sites in BM stromal cell cocultured B cells. Purified immature and transitional B cells from 6–1/V{kappa}1A mice were cocultured with Rag1–/– BM cells at a ratio of 1:3 (B cell to BM). The cells were stimulated with anti-Igβ or control Abs for 24 h and analyzed for DSB. J{kappa}5 was not detectable in both immature and transitional B cells. B, RAG expression in B cells of 6–1/V{kappa}1A mice cocultured with Rag1–/– BM stromal cells. The cells from the same source as in A were used to measure the levels of RAG1 and RAG2 transcripts by qPCR. C, RAG expression in non-Tg transitional B cells cocultured with Rag1–/– BM stromal cells. The culture condition was same as in A. D, Evaluation of DSB at J{kappa} RSS sites in non-Tg transitional B cells. The cells were from the same source as in C. All data represent three independent experiments. E, Flow cytometry analysis showed the existence of Thy1dullDX5+ stromal cells in Rag1–/– BM. The numbers are percentages of cells falling in the gate (n = 4).

 
The levels of histone H3 acetylation and trimethylation on lysine4 (K4) are comparable between immature and transitional B cells

Chromatin remodeling regulates V(D)J rearrangements through progressive stages of early B cell development (28). High levels of histone H3 acetylation and methylation on K4 are associated with an open chromatin structure for the recombination machinery (29, 30). To determine the level of acetyl-H3 and methyl-H3-K4 at J{kappa} gene segments, we performed ChIP assays using cross-linked DNA from 6–1/V{kappa}1A immature and transitional B cells and Abs against acetylated H3 and trimethylated H3-K4. After cross-linking was reversed, the J{kappa}5 sequence was amplified by real time PCR. As shown in Fig. 7, the levels of acetyl-H3 at J{kappa}5, calculated as the percentage of input DNA, were comparable between resting transitional B cells (27.7%) and immature B cells (15.5%). The levels of trimethyl-H3-K4 at J{kappa}5 were also similar between the two subsets (transitional B cells, 7.5%, vs immature B cells, 7.8%). These levels were equivalent to those in pre-BII cells in which an open chromatin structure at J{kappa} loci is expected. After stimulation with anti-Igβ Ab, the levels of acetyl-H3 and trimethyl-H3-K4 at J{kappa}5 increased in both subpopulations. Transitional B cells appear to have higher levels of acetyl-H3 (59.2%) than immature B cells (32.5%) even though the difference was statistically insignificant. As a negative control, the levels of acetyl-H3 and trimethyl-H3-K4 at the TCR{alpha} locus were very low (≤0.1%, Fig. 7), consistent with a "closed" locus. We conclude from these data that the J{kappa} gene loci of transitional B cells are likely to be accessible for RAG recombination. This parallels the open chromatin structure seen at Ig VH loci in mature B cells (31).


Figure 7
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  		FIGURE 7. BM immature B and splenic transitional B cells show equal levels of histone H3 acetylation and trimethylation on Lysine4. 6–1/V{kappa}1A BM immature B cells were obtained from cultured BM cells with IL-7 for 5 days followed by 18 h without IL-7. BM immature B cells and whole splenic cells from 6–1/Vk1A mice were stimulated with anti-Igβ Abs for 18 h and were sorted for IgM+PtC– cells for use in ChIP analyses. The small pre-BII cells sorted from non-Tg mice were used as a control. Data represents one of two separate experiments with similar results.

 
Immature and transitional B cells exhibit distinct phenotypic changes following BCR ligation

Treatment with PMA (2 ng/ml) stimulated increased expression of CD23 on transitional but not on immature B cells (Fig. 8). In addition, PMA-activated transitional cells exhibited a decrease in IgM and HSA expression as well as an increase in CD21 expression (Fig. 8). Fewer immature B cells underwent these changes. These phenotypic changes were consistent with alterations seen after in vivo anti-Igβ injection (Fig. 5 and Ref. 17). These findings further support the conclusion that immature and transitional B cells are programmed to activate innately different responses to signals induced by BCR ligation.


Figure 8
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  		FIGURE 8. BM immature and splenic transitional B cells exhibited different phenotypic changes following stimulation with PMA. BM cells from 6–1/V{kappa}1A/RAG1–/– mice were cultured with 10 ng/ml IL-7 for 5 days followed by 18 h without IL-7 to enrich immature B cells (>95% pure). Splenic B cells from 6–1/V{kappa}1A/RAG1–/– mice were purified by negative selection. The cells were then cultured in parallel with and without 2 ng/ml PMA for 24 h and analyzed by flow cytometry. Numbers indicate the percentage of cells falling in each gate. For overlays, filled line, medium controls; open line, PMA treated. Representative data from more than four separate experiments is shown.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
These experiments demonstrate that transitional B cells lose the capacity for receptor editing in response to a BCR signal. This differs from immature B cells that undergo BCR ligation-triggered secondary L chain gene rearrangements in response to the same treatment. Interestingly, the capacity for receptor editing is not correlated with RAG expression levels; whereas there was no significant change in RAG expression, either at the RNA or protein levels in anti-Igβ-treated immature B cells, the expression levels of RAGs were increased considerably in anti-Igβ-treated transitional B cells. Thus, caution should be used in interpreting increases in RAG levels.

The 6–1/V{kappa}1A B cell development is blocked at the T2 cell stage (17). Phenotypical and functional analyses have failed to reveal any difference between 6–1/V{kappa}1A and normal transitional B cells (17). The presence of B cells expressing endogenous L chains in BM immature B cells of 6–1/V{kappa}1A mice is most likely due to a "premature" expression of {kappa} L chains in early (CD43+) B cells (32, 33) than to self-Ag-induced receptor editing. We are currently unable to rule out the possibility that the 6–1/V{kappa}1A BCR is unable to generate sufficient tonic signaling to complete L chain allelic exclusion and mature B cell differentiation. However, several findings are inconsistent with weak tonic signaling. First, 6–1/V{kappa}1A B cells express normal levels of surface BCR and elicit normal levels of protein tyrosine phosphorylation following BCR cross-linking (17). Second, insufficient tonic signaling would most likely result in frequent endogenous L chain rearrangement in pre-BII/immature B cells, rather than in just a few cells (Fig. 2). Third, Shivtiel et al. (34) have demonstrated that the loss of CD19 induces an increase in receptor editing due to the failure of the BCR to provide an effective tonic signal. This predicts that if the defect in 6–1/V{kappa}1A B cells is a lack of tonic signaling, then reducing the level of CD19 should have no effect on DSB and RAG expression. However, the BM immature B cells of 6–1/V{kappa}1A/CD19+/– mice with the expression level of CD19 reduced by 50% exhibited a 5-fold increase in RAG expression and more abundant DSB at J{kappa}2 and J{kappa}4 loci compared with 6–1/V{kappa}1A/CD19+/+ mice (data not shown). CD19 deficiency also failed to promote B cell maturation beyond the T2 cell stage (data not shown). Thus, the 6–1/V{kappa}1A BCR provides a tonic signal that is sufficient to block receptor editing in the bone marrow.

Receptor editing plays a critical role in shaping the Ab repertoire. It is estimated that 25–50% of developing immature BM B cells undergo receptor editing (1, 3, 35, 36). Earlier studies with mice bearing conventional transgenes to MHC class I show that receptor editing occurs in IgMlow but not in IgMhigh immature B cell populations (24). However, studies using more physiologically relevant Ig gene knockin mice suggest that editing occurs in pre-BII cells rather than in immature B cells (37). This observation has been extended by Casellas et al. (1) showing that editing at the pre-BII cell stage causes 2-h delay in the transition from pre-BII to immature B. In contrast to these findings, Tze et al. (38) demonstrated that both IgMlow and IgMhigh immature B cells undergo receptor editing when stimulated with soluble HEL. Thus, when receptor editing occurs during B cell differentiation is in dispute. It may be dependent on the nature of the Ag and the location of Ag encounter. Even though our study did not distinguish between IgMlow and IgMhigh immature B cells, our data are consistent with the conclusion that receptor editing can occur in immature BM B cells.

In the spleen, transitional B cells are either positively selected into the long-lived mature B cell pool or die by negative selection mechanisms including deletion (apoptosis) and negligence (lack of positive selection). Evidence for peripheral deletion comes from studies using Tg mice expressing BCRs for a liver-expressed autoantigen with high affinity (9). In this setting, cross-linking BCRs on transitional B cells results in apoptosis in vitro (10, 39). These studies indicate that transitional B cells are as sensitive as BM immature B cells to BCR-mediated apoptosis. Our recent studies have shown that the development of 6–1/V{kappa}1A B cells is blocked at a T2 cell stage without evidence of Ag-imposed negative selection (17). Interestingly, injection of 6–1/V{kappa}1A/Rag1–/– mice with a high dose of anti-Igβ Ab induced apoptosis while a low dose of anti-Igβ induced differentiation as assessed by the expression of maturation markers and longer survival of treated B cells in vitro (17). This model argues that "negligence" or lack of a ligand-mediated positive selection signal may account for the death of 6–1/V{kappa}1A B cells at the T2 cell stage. Studies by others also support a role of Ag-mediated positive selection in the development of mature B cells (reviewed in Ref. 8). Thus, transitional B cells appear to be hypersensitive to both negative and positive signals and the choice of positive vs negative selection is highly dependent on the BCR signaling strength. That the same anti-Igβ signal induces receptor editing in immature cells and promotes transitional to FO B cell differentiation suggests that the threshold for deletion/editing vs positive selection shifts during differentiation.

The role of receptor editing in development of transitional B cells is poorly understood. The levels of RAG1 and RAG2 expression drop drastically when immature B cells become transitional B cells (Fig. 2 and Ref. 40, 41). This is consistent with an inability of transitional B cells to edit. It is surprising that expression of RAG can be reinduced when transitional B cells receive a BCR signal (Fig. 4 and Ref. 26), yet DNA rearrangement does not occur (Fig. 4). Thus, the mechanism preventing editing in transitional B cells does not involve regulation of RAG protein levels. Control of editing could be due to multiple mechanisms. First, transitional B cells may be unable to survive stimulation strong enough to induce editing. This possibility is supported by a previous study in which an anti-apoptotic agent was used to facilitate anti-IgM-induced receptor editing (26). Second, the comparable levels of histone H3 acetylation and trimethylation at Lys4 on J{kappa}5 between immature and transitional B cells argue against a "closed" chromatin structure in transitional B cells. However, if RAG proteins are functional, there may be other levels of histone modification that prevent RAG access to J{kappa} genes. In addition, our results suggested that the differing response of immature and transitional B cells to BCR stimulation could be due to distinct signaling networks. Such differences also exist even among transitional B cells (42). The expression level of mitogen- and stress-activated kinase 2 (MSK2) is >6-fold higher in transitional B cells than in immature B cells (data not shown). MSK2 is involved in phosphorylation of histone H3 at Ser (10) and Ser (28) (43). The increased expression of MSK2 and possibly other signaling molecules in transitional B cells may induce secondary modification of histones leading to decreased accessibility of Ig loci.

Third, the activity of RAG proteins may be differently regulated by phosphorylation and/or ubiquitinylation in immature and transitional B cells. The discrete domains outside the recombinase-core region of RAGs are potential regulatory structures that can be phosphorylated (44) and ubiquitinylated (45). Because the levels of RAG transcripts and proteins were comparable between immature and transitional B cells (Fig. 2 and Fig. 4), modification of RAG proteins by either of these two mechanisms would reduce recombinase activity and prevent secondary gene rearrangements. Future studies are warranted to clarify these issues.

The suggestion that transitional B cells are unable to edit does not conflict with other studies in which receptor editing was found in periphery lymphoid organs of mice undergoing immunization or during an autoimmune response (11, 12, 13, 14). Under those conditions, non-BCR signals, such as cytokines, may override the recombination restraining mechanism found in transitional B cells and activate V(D)J recombination fully. Because stimulation of circulating B cells (including transitional and mature B cells) with a bacterial Ag, Staphylococcus aureus Cowan I, plus IL-2 induces expression of RAG and L chain gene rearrangement (46), it is also possible that receptor editing in peripheral B cells could occur through a BCR-independent mechanism. This may account for receptor editing found in germinal center B cells (11, 21).

Why do transitional B cells develop such a restriction mechanism to prevent recombination? We speculate that this may be of biological importance for preserving genetic stability and reducing susceptibility to RAG-mediated lymphomagenesis. More importantly, this mechanism may prevent emergence of dual L chain-expressing B cells. We reason that if receptor editing is permissible in transitional B cells, dual receptor-expressing B cells would be more frequently produced due to the increased sensitivity of transitional B cells to Ag-mediated positive selection. In view of our previous findings that low concentrations of anti-Igβ Ab induced differentiation of transitional B cells and high concentrations induced apoptosis (17), we conclude that although transitional B cells are limited in receptor editing, they remain sensitive to other positive and negative selection mechanisms.


   Acknowledgments
 
We thank Dr. Larry Arnold at University of North Carolina at Chapel Hill (Chapel Hill, North Carolina) for assistance in cell sorting.


   Disclosures
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
The authors have no financial conflict of interest.


   Footnotes
 
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1 This work was supported in part by the Intramural Research Program of the National Institutes of Health, National Institute of Allergy and Infectious Diseases, and National Institutes of Health Grants AI29576 and AI43587 (to S.H.C.) Back

2 Address correspondence and reprint requests to Dr. Hongsheng Wang, Twinbrook 1, Room 1518, Laboratory of Immunopathology, National Institute of Allergy and Infectious Diseases, National Institutes of Health, 5640 Fishers Lane, Rockville, MD 20852 and Dr. Stephen H. Clarke, Department of Microbiology and Immunology, University of North Carolina at Chapel Hill, Chapel Hill, NC 27599. E-mail addresses: wanghongs{at}niaid.nih.gov and shl{at}med.unc.edu Back

3 Abbreviations used in this paper: BM, bone marrow; BCR, B cell receptor; RSS, recombination signal sequence; FO, follicular; MZ, marginal zone; Tg, transgenic; ChIP, chromatin immunoprecipitation; PtC, phosphatidyl choline; DSB, dsDNA break; P.I., PMA plus ionomycin; LM, ligation mediated; int, intermediate; K4, lysine4; MSK2, mitogen- and stress-activated kinase 2. Back

Received for publication February 1, 2007. Accepted for publication September 26, 2007.


   References
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 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 

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