Activation-Induced Cytidine Deaminase-Dependent DNA Breaks in Class Switch Recombination Occur during G₁ Phase of the Cell Cycle and Depend upon Mismatch Repair¹

Antibody class switch recombination (CSR)³ involves the replacement of the IgM C region gene (Cμ) by a downstream C region gene, e.g., Cγ, Cα, or Cε, and can improve the efficacy of the immune response. The recombination occurs via the formation of double-strand breaks (DSBs) in switch (S) region DNA located upstream of the C region genes, and the joining of two different S regions by an end-joining mechanism, resulting in deletion of the intervening DNA from the genome (1). CSR requires activation induced cytidine deaminase (AID), which converts cytosines in DNA to uracils (2, 3, 4, 5, 6, 7). The target for AID is ssDNA, which can be generated during transcription, and only transcriptionally active S regions undergo CSR (1, 4, 8, 9, 10). S regions consist of tandem repeats (TR) that are unique to each isotype, although all contain numerous targets for AID: the hot-spot motif WRC/GYW, where W = A or T, R = G or A, and Y = C or T (6, 11, 12). The underlined C is determinated by AID.

Repair of the dU residues resulting from AID activity leads to DNA breaks necessary for recombination. Uracil in DNA is removed by uracil DNA glycosylase (UNG) and CSR is severely reduced in mice that lack UNG and in patients with mutations in UNG (13, 14). UNG activity generates abasic sites that could be recognized and cleaved by AP endonucleases (APE) to create single-strand breaks (SSBs), and recent data indicate that APE is important for CSR (53). AID- and UNG-dependent DSBs have been detected in S regions, and the breaks occur preferentially at G:C bp in AID hot-spot motifs, which is consistent with cleavage by APE at the nucleotide deaminated by AID (14, 15, 16, 17). The current data support the conclusion that repair of the dU residues generates SSBs, but SSBs must be converted to DSBs to excise the DNA segment intervening between Sμ and the downstream S region.

Currently, nothing is known about the mechanism by which SSBs in S regions become DSBs. In this study, we test three hypotheses regarding how DSBs are created during CSR. SSBs initiated by AID activity could be converted into DSBs during replication, when the elongating DNA strand reaches a nick on the template strand. However, γH2AX and Nbs-1, two proteins involved in CSR, are found associated with the IgH locus during G₁/early S phase, but not during S/G₂M phase in splenic B cells undergoing CSR (18). To address this issue, we analyzed the cell cycle regulation of DSBs in S regions, with emphasis on separation of G₁ phase from S phase cells.

Nucleotide excision repair (NER) could recognize a variety of damage intermediates resulting from AID activity and could introduce DNA breaks with its associated endonucleases Ercc1-Xeroderma pigmentosum F (XPF) and XPG (19). A minor role for Ercc1 in CSR has been described (20), but it is not known whether Ercc1-XPF acts during CSR in conjunction with the NER pathway or independently as a structure-specific endonuclease. We tested whether XPA, which is essential for NER, is involved in CSR.

Nicks sufficiently close (<5 bp apart) on opposite strands can form DSBs spontaneously, as shown by experiments in which the restriction enzyme I-Sce1 can produce a chromosomal DSB (21). However, distal SSBs will not lead to DSBs, due to stability of the DNA duplex, unless the DNA ends are processed. AID is known to deaminate dC nucleotides on both the transcribed and nontranscribed strands (22, 23), and the density of AID hot spots occurring in the S region TRs could lead to nicks on opposite strands in close proximity to each other. However, deletion of the SμTR region results in only a modest (2-fold) reduction in CSR (24). CSR can occur upstream of, as well as within, the SμTR region (25, 26), although recombination outside of the SμTRs requires the DNA mismatch repair (MMR) pathway (26, 27). In fact, CSR is nearly ablated in SμTR^−/− B cells that also lack the MMR proteins Msh2 or Mlh1 (28) (J. Eccleston, C. E. Schrader, J. Stavnezer, and E. Selsing, manuscript in preparation). Because MMR is required for recombination outside the SμTR region where AID hot spots are relatively infrequent, we have proposed that MMR is needed to convert SSBs into DSBs when the single-strand nicks are too far apart to form spontaneous DSBs (29). This hypothesis is consistent with the finding that CSR does not require MMR, but is reduced 2- to 5-fold in MMR-deficient B cells, i.e., at least 50% of CSR events require MMR, depending on the isotype (27, 30, 31, 32, 33, 34). To test this hypothesis, we analyzed Sμ DSBs in B cells from mice deficient in MMR proteins that have the WT Sμ region or the SμTR deletion. Altogether, the data support the hypothesis that distal SSBs are converted by MMR into DSBs, constituting an important step in CSR.

AID-deficient mice were obtained from T. Honjo (Kyoto University, Kyoto, Japan). Msh2-deficient mice were obtained from T. Mak (University of Toronto, Toronto, Canada). Mlh1- and Pms2-deficient mice were obtained from R. M. Liskay (Oregon Health Sciences University, Portland, OR). XPA-deficient mice were obtained from H. van Steeg (National Institute of Health, Bilthoven, The Netherlands) (35). These lines have all been extensively backcrossed to C57BL/6. xpa^+/− mice were bred to ung^+/− mice (obtained from D. Barnes and T. Lindahl, London Research Institute, London, U.K.) to generate doubly heterozygous mice, the breeding of which generated wild-type (WT), xpa^−/−, ung^−/−, and xpa^−/− ung^−/− littermates used for the experiment shown in Fig. 3. SμTR^+/− mice were bred to mlh1^+/− and to msh2^+/− mice to generate double knockouts. WT (+/+) littermates from MMR-deficient mice were used as controls for all experiments (except that shown in Fig. 3) and the results were identical for all WT littermates. All mice were housed in the same room of the Institutional Animal Care and Use Committee-approved specific pathogen-free facility at University of Massachusetts Medical School and were bred and used under guidelines formulated by the University of Massachusetts Animal Care and Use Committee.

Spleen cells were dispersed and RBCs lysed in Gey’s solution followed by T cell depletion with a mixture of anti-T cell Abs, as described previously (30). Cells were cultured at 1 × 10⁵/ml in 6-well plates and activated to induce CSR. All cultures contained LPS (50 μg/ml; Sigma-Aldrich) and human BLyS (100 ng/ml), Human Genome Sciences. IL-4 (800 U/ml) was added to induce switching to IgG1; IFN-γ (10 U/ml) for IgG2a; dextran sulfate (30 μg/ml; Amersham Biosciences) for IgG2b; anti-δ-dextran (0.3 ng/ml) for IgG3, and to induce IgA switching, TGF-β (2 ng/ml), IL-4 (800 U/ml), IL-5 (1.5 ng/ml; BD Biosciences), and anti-δ-dextran (0.3 ng/ml) were added. Fixing of cells and staining of cell surface Abs were performed as described previously (30), except the results were analyzed by FlowJo software.

Splenic B cells were cultured for 2 days as described above, after which cell density was adjusted to 1 × 10⁶ cells/ml and stimulated B cells were incubated for 90 min at 37°C with 3.5 μg/ml Hoechst 33342 (Invitrogen Life Technologies) in HBSS containing 1% FCS. Cells were finally resuspended in 1 ml of HBSS with 1% FCS; 7-aminoactinomycin D was added (0.6 μg/ml) and cells were sorted by flow cytometry based on DNA content using a UV laser-equipped FACSVantage SE (BD Biosciences).

To prepare nuclear extracts, cells were resuspended in hypotonic buffer (10 mM HEPES (pH 8), 1 mM EDTA, 10 mM KCl, 0.1 mM EGTA, 1 mM DTT, 2 μM pepstatin, and complete protease inhibitor mixture); after a 15-min incubation on ice, cells were lysed by addition of Nonidet P-40 to a final concentration of 0.625%. After centrifugation at 12,000 rpm in a microfuge for 10 min, the pelleted nuclei were washed once with hypotonic buffer and resuspended in hypertonic buffer (20 mM HEPES (pH 8), 1 mM EDTA, 400 mM NaCl, 1 mM EGTA, 1 mM DTT, 2 μM pepstatin, and complete protease inhibitor mixture). Protein content of nuclear extracts was determined using the Bradford assay (Bio-Rad). Proteins were electrophoresed on 10% SDS-polyacrylamide gels or 4–20% gradient SDS-polyacrylamide gels (Bio-Rad), and blotted onto Immobilon-P polyvinylidene fluoride membranes (Millipore). Immunoblotting was performed using rabbit polyclonal Abs against AID and UNG (rabbit anti-peptide amino acids 280–295 from mouse UNG), APE1 (36), MSH2 (sc-494), MSH6 (sc-10798), MLH1(sc-582), and GAPDH (Santa Cruz Biotechnology), and monoclonal mouse anti-TBP1 (Abcam) followed by goat-anti-rabbit or donkey anti-mouse-HRP (Santa Cruz Biotechnology) and ECL substrate (Pierce).

Whole cell extracts were prepared from splenic B cells activated for 2 days, washed in ice-cold PBS, and lysed in buffer I (10 mM Tris-HCl (pH 7.8), 200 mM KCl with Complete protease inhibitors (Roche)). An equal volume of buffer II was added (buffer I with 2 mM EDTA, 40% glycerol, 0.2% Nonidet P-40, and 2 mM DTT added) before tumbling for 1 h at 4°C. Supernatants were stored at −80°C. Nuclear extracts were made from sorted cells by lysis in buffer A (10 mM HEPES (pH 8), 10 mM KCl, 0.1 mM EDTA, 0.1 mM EGTA, 1 mM DTT, 0.5 mM PMSF, 2 μM pepstatin, and Complete protease inhibitor) with 0.625% Nonidet P-40. Nuclei were pelleted and extracted with buffer C (20 mM HEPES (pH 8), 400 mM NaCl, 1 mM EDTA, 1 mM EGTA, 1 mM DTT, 40% glycerol, 0.5 μM pepstatin, and Complete protease inhibitor). Extracts were incubated with a [³²P]5′ end-labeled uracil-containing oligonucleotide (5′-GATTCCCCATCTCCTCAGTTTCACT/ideoxyU/CTGCACCGCATG-3′; IDT) in BER buffer (40 mM HEPES-KOH (pH 7.8), 5 mM MgCl₂, 0.5 mM DTT, 0.1 mM EDTA, 2 mM NaVO₄, 50 mM NaF, 25 mM β-glycerophosphate, 2 mM sodium pyrophosphate) at 37°C. After 1 h, NaOH was added to 0.1 M and reactions were heated to 95°C for 7 min and electrophoresed on an 8 M urea, 15% polyacrylamide bis (19:1) gel.

After culture for 2 days, viable cells were isolated by flotation on Ficoll/Hypaque gradients (ρ = 1.09), or lympholyte (Cedarlane Laboratories), cells were imbedded in low-melt agarose, and DNA was isolated and used for LM-PCR as described (15). Briefly, DNA was ligated overnight to linker, which was prepared by annealing 5 nM each of LMPCR.1 (5′-GCGGTGACCCGGGAGATCTGAATTC-3′) and LMPCR.2 (5′-GAATTCAGATC-3′) in 300 μl of 1× ligase buffer, resulting in a double-stranded oligo with a 14-nt single-strand overhang that ligates unidirectionally. Ligated DNA samples were assayed for GAPDH by PCR to adjust DNA input before LM-PCR. The primer 5′ Sμ (5′-GCAGAAAATTTAGATAAAATGGATACCTCAGTGG-3′) was then used in conjunction with linker primer (LMPCR.1) to amplify DNA breaks. Three-fold dilutions of input DNA were amplified by HotStar Taq (Qiagen). PCR products were run on 1.25% agarose gels and vacuum blotted (VacuGene XL; Pharmacia) onto nylon membranes (GeneScreen Plus; PerkinElmer). Blots were hybridized at 37°C overnight with an Sμ -specific oligonucleotide probe (μ probe, 5′: AGGGACCCAGGCTAAGAAGGCAAT) end labeled with [γ-³²P]ATP and washed at 55°C with 2× SSC/0.1% SDS.

LM-PCR products were cloned into the vector pCR4-TOPO (Invitrogen Life Technologies) and sequenced by Macrogen using T3 and T7 primers. Cloned breaks in Sμ were aligned with germline Sμ sequenced from C57BL/6 chromosome 12 (GenBank Accession number AC073553) with numbering starting at nt 136,645. This is the 5′ Sμ primer-binding site and ∼800 nt upstream of the beginning of the TRs.

A possible mechanism for conversion of SSBs into DSBs is DNA replication, and many of the proteins involved in CSR are associated with DNA surveillance and repair during replication, e.g., MMR proteins and UNG (37). To determine when AID-dependent DSBs are made, we examined the cell cycle regulation of Sμ DSBs. DSBs dependent on AID and UNG are induced in the S regions of B cells activated in vitro to undergo CSR and can be detected by linker LM-PCR (15). We activated splenic B cells for 2 days with LPS and IL-4 to induce IgG1 CSR or LPS and anti-δ-dextran (a-δ-dex) to induce IgG3 CSR, stained with Hoechst 33342, and then sorted cells into G₁ and S/G₂/M fractions on the basis of DNA content (Fig. 1,A). Dead cells were excluded by 7-aminoactinomycin D staining. At 48 h, there are no undivided cells in these cultures, as determined by CFSE staining (our unpublished data). As shown in Fig. 1 B, AID-dependent DSBs are detected almost exclusively in the G₁ fraction in cells activated under either condition. There are a few breaks detected in S/G₂/M phase, but there are about as many in AID-deficient B cells. These data are therefore inconsistent with the hypothesis that DSBs detected in Sμ in switching B cells are due to replication across SSBs. Although a few DSBs are detected in aid^−/− cells, they do not specifically occur at the G:C base pair in the AID target hot spots, unlike DSBs in WT cells (15). They might be due to mechanically induced breaks or apoptosis occurring during the procedure, or perhaps due to replication.

We then asked whether the levels of AID, UNG, APE1, and MMR proteins are regulated by the cell cycle. B cells activated with LPS and a-δ-dex were sorted into G₁, S, and G_2/M fractions (Fig. 2,A) and extracts were prepared for Western blotting (Fig. 2, B–D). AID protein is difficult to detect in nuclear extracts, but is expressed in the cytosol in all phases of the cell cycle (Fig. 2,C). UNG is abundant both in the nucleus (Fig. 2,B) and cytoplasm (Fig. 2,C) of G₁- and S-phase cells. The multiple UNG bands observed in nuclear extracts represent differently phosphorylated species (38). The Ab should also detect the mitochondrial form of UNG in the cytosol. We observed a decrease in UNG in the cytosol as the cells progress through cell cycle, consistent with a report that UNG is down-regulated in late S phase (39). APE1 and the MMR proteins Msh2, Msh6, and Mlh1 are all expressed in the nucleus throughout the cell cycle (Fig. 2 D). We conclude that proteins important for CSR are present in all phases of the cell cycle, and specifically in G₁ phase when AID-dependent DSBs are detectable.

UNG is not present in resting (G₀) cells (14) and has been proposed to act in S phase at replication forks during CSR and somatic hypermutation (40). G₁-phase B cells have not been examined for UNG activity. We prepared nuclear extracts from G₁- and S-phase B cells activated for 2 days and assayed UNG activity by an oligonucleotide cleavage assay (Fig. 2 E). Activity is clearly detectable in as little as 10 ng of nuclear extract from G₁-phase cells, which has as much activity as extracts from S-phase cells. As this substrate is single stranded, the predominant activity detected here is likely to be due to UNG and not SMUG-1, as SMUG-1 has 800-fold lower activity on ssDNA than UNG (k_cat/K_m) (41). From these combined results, we conclude that AID-dependent DSBs in Sμ are made and resolved in Sμ in G₁ phase and are not due to replication across SSBs.

We next asked whether NER might contribute to the generation of DSBs during CSR. We have previously shown a minor role for Ercc1 in CSR (20). The heterodimer Ercc1-XPF is an essential component of the NER pathway, but Ercc1-XPF can also act as a structure-specific endonuclease in the absence of the rest of the NER pathway, cutting at the junction of ssDNA and dsDNA (19). Here, we analyzed CSR in vitro in B cells from mice lacking XPA, a protein essential for NER. We found no reduction in CSR in xpa^−/− B cells relative to cells from WT littermates (Fig. 3,A). In the absence of UNG, it seemed possible that the accumulation of uracils might result in distortion of the DNA helix sufficient to create a substrate for NER, so we also analyzed CSR in ung^−/−xpa^−/− B cells CSR is severely reduced in B cells deficient in UNG, as reported by others (13, 14), but still detectable relative to AID-deficient B cells. However, CSR is not further reduced in ung^−/−xpa^−/− B cells relative to B cells from ung^−/− littermates (Fig. 3, B and C). In this experiment, there was a reduction in IgG1 and IgG2a CSR in xpa^−/− cells, but this was not typical as can be seen from the compiled data in Fig. 3 A. We conclude that the minor role played by Ercc1-XPF in CSR most likely involves its ability to cut at the junction of ssDNA and dsDNA (discussed below and in Ref. 29), and does not involve other components of the NER pathway.

The hypothesis that SSBs are converted to DSBs in S regions by MMR predicts fewer DSBs in Sμ in MMR-deficient B cells relative to WT cells. To test this prediction, LM-PCR was performed on genomic DNA isolated from B cells 2 days after induction of CSR. A representative Southern blot of LM-PCR fragments amplified with a 5′ Sμ-specific primer from WT, msh2^−/−, mlh1^−/−, and pms2^−/− B cells is shown in Fig. 4,A. Blunt DSBs in MMR-deficient cells are reduced compared with WT cells analyzed in parallel. Quantitation by densitometry scanning of the autoradiographs from several experiments demonstrates that MMR deficiency results in ∼20–50% of the number of Sμ DSBs in WT cells (Fig. 4 B). Thus, blunt DSBs induced in Sμ during CSR are reduced in MMR-deficient cells to a degree consistent with the reduction observed in CSR.

As described in the introduction, CSR in B cells lacking the Ig Sμ TR region (SμTR^−/−) is severely reduced (90–95%) in the absence of Msh2 or Mlh1. We used LM-PCR to determine whether this might be due to a decrease in DSBs. As shown in Fig. 5, DSBs in SμTR^−/− B cells are detected at only a slightly lower frequency than in WT cells (65% of WT), consistent with the 2-fold reduction in CSR in these B cells (24), but when the cells also lack Msh2 or Mlh1, DSBs are reduced to only 20% of that seen in WT cells. The number of DSBs detected in Msh2/SμTR and Mlh1/SμTR double knockout cells is almost as low as in AID-deficient cells.

Relative to the WT Sμ region, the SμTR^−/− intron has a 2-fold lower incidence of AID hot spots (WRC/GYW, where the underlined nucleotide is the target site), and the strongly preferred hot-spot GCT (11, 33) is 3.6-fold less frequent than in WT Sμ (Fig. 6). We asked whether the DSBs in SμTR^−/− cells occur preferentially at these remaining hot spots by cloning and sequencing LM-PCR products from WT and SμTR^−/− B cells. Table I presents the percent of DSBs occurring at the G:C or A:T base pair, at all AID hot spots (GYW), and at the four different GYW motifs (GCT, GCA, GTT, and GTA), as well as the frequency of occurrence of these motifs in the Sμ-region sequence in WT and SμTR^−/− cells. The percentage of DSBs located at GYW sequences is increased in SμTR^−/− B cells (51.6% compared with 40.5% in WT), supporting the importance of this sequence for AID targeting.

Further analysis of the breakpoints used in the SμTR^−/− intron support previous conclusions that GCT/AGC is the strongest AID hot spot. The preference for breaks at GCT in WT Sμ is 3.3-fold over random (35.4% of DSBs at GCT vs 10.6% frequency of GCT in the sequence, p < 0.001). However, GCT occurs in the WT Sμ sequence at a much higher frequency than the other GYW hot spots (Table I). In the SμTR^−/− intron, the frequency at which GCT occurs relative to the other GYW motifs is lower; interestingly, DSBs at GCT occur 10-fold more often than predicted by the sequence (29% of DSBs are at GCT compared with 2.9% frequency in the sequence, p < 0.001). These results confirm that the preference of AID for GCT hot spots previously found in vitro with purified AID and from analyses of somatic hypermutation (6, 11, 12, 42, 43) is also true during CSR, and is even more apparent when these motifs are less abundant. We hypothesize that the reduction in AID hot spots in SμTR^−/− B cells results in SSBs that are farther apart, and thus less likely to be sufficiently close to form a DSB spontaneously. DSB formation and CSR are therefore more dependent on MMR in these mice.

The results from our studies support the hypothesis that distal SSBs are converted into DSBs by MMR recognition and repair of dU:dG mismatches introduced by AID. We obtained data arguing against an alternative model for conversion of SSBs to DSBs, i.e., replication across a SSB, as we detected very few DSBs in S/G₂/M phase cells. Instead, the AID-dependent DSBs occur in the G₁ phase. These results are in agreement with previous reports showing that AID-dependent γH2AX/Nbs1 foci colocalize with IgH loci during the G₁/early S phase in splenic B cells activated to switch (18). They are also consistent with the finding that switch recombination is not accompanied by sister-chromatid exchange (44), and that a specific protein complex binds near the transcriptional initiation site for germline γ1 transcription during G₁/early S phase, but not during G₂/M phase in splenic B cells activated to undergo CSR (45). We also found that AID, UNG, and MMR proteins are present in the G₁ phase of the cell cycle, and that extracts from G₁-phase B cells possess UNG activity.

MMR proteins could convert SSBs into DSBs in the course of normal MMR activity if nicks are present on both strands. Both AID and UNG prefer ssDNA substrates and appear to act on both DNA strands during transcription (1, 12, 22, 46). We hypothesize that the DNA duplex reforms before all of the dUs can be removed by UNG, and MMR would then compete more effectively than UNG for repair of dUs in duplex DNA. AID is a highly processive enzyme and although UNG has a very high catalytic rate, it appears to be unable to remove all the dUs introduced by AID into S regions (6, 12, 22, 23, 41). UNG is likely, however, to create numerous abasic sites, which could then be nicked by APE to generate entry sites for excision by Exo1 in the 5′ to 3′ direction. If excision continued until a SSB on the opposite strand is reached, a DSB would be formed. A similar model has been proposed for Escherichia coli dam cells exposed to the methylating agent MNNG, in which DSBs arise when MMR excision on one strand encounters a nick generated by APE on the other (47). Depending on the orientation of the SSBs relative to each other, some DSBs created by MMR activity will be blunt and some will have 5′ or 3′ overhangs. 5′ overhangs can be filled in by polymerase to become blunt, and we have proposed that Ercc1-XPF can remove 3′ overhangs (20, 29). Ercc1-XPF cuts 3′ overhangs at the junction of ssDNA and dsDNA without requiring additional NER proteins (48). Consistent with Ercc1-XPF acting independently of NER in CSR, we found that XPA, an essential NER protein, is not involved in CSR even in the absence of UNG, where helix-distorting lesions recognizable by NER might form.

If the SSBs are the results of AID-instigated lesions, the resulting blunt DSBs would occur preferentially at the G:C base pair in AID hot spots, which is consistent with the location of DSBs in Sμ determined by LM-PCR (Table I) (15, 29). No significant differences were observed between the sites of Sμ DSBs in WT and MMR-deficient B cells (Table II). In the absence of MMR, we predict that the only DSBs formed would do so spontaneously from closely spaced nicks. The staggered DSBs produced in the absence of MMR could then be end processed as described above to form the blunt DSBs at the G:C base pair in AID hot spots detected by LM-PCR. The staggered ends can be detected by LM-PCR by pretreatment with T4 DNA polymerase before ligation (15, 17). We find a 3-fold increase in DSBs after T4 treatment of DNA from both WT and MMR-deficient cells (data not shown).

It has been reported that 94% of Sμ-Sγ3 junctions in Msh2-deficient cells occur within the Sμ TRs, which contain numerous closely spaced GAGCT sequences, and only 5% occur 5′ to the TRs, whereas in WT cells a greater proportion (17%) of the junctions occur 5′ to the repeats (26, 27). These results predict that blunt DSBs in MMR-deficient B cells would localize preferentially to the Sμ TRs. However, we found a similar fraction of DSBs occur 5′ to the Sμ TRs in WT and msh2^−/− cells, 42 and 40%, respectively. This difference from the previous reports might be due to differences in the analysis methods. We used a PCR primer located at the 5′ side of Sμ, and extension would terminate at the first break, favoring detection of breaks located upstream of Sμ.

We consistently find fewer blunt DSBs in B cells from mice lacking Mlh1 or Pms2 than in those lacking Msh2. This might be explained by the recent finding that Mlh1 decreases Exo1 processivity (49). If Exo1 excises farther in the absence of Mlh1-Pms2, perhaps past the nick on the opposite strand, long single-strand tails and fewer blunt DSBs would be generated. Long single-strand overhangs might also explain the increase in microhomology found at CSR junctions from mlh1^−/− and pms2^−/− mice. In addition, it has recently been shown that the Mlh1-Pms2 heterodimer has endonuclease activity that introduces nicks on either side of the mismatch (50). This activity appears to be restricted to the previously nicked strand (50), and so by itself would not result in DSBs during CSR. Furthermore, as the Mlh1-Pms2 endonuclease does not appear to be restricted to a specific DNA sequence, and Sμ DSBs strongly favor the G:C base pair, this activity is unlikely to contribute significantly to S-region breaks.

Our results are consistent with the model that MMR is more important for CSR in situations when SSBs are limiting, as when AID targets are scarce. Thus, loss of both MMR and the SμTR region results in a dramatic reduction in switching and a reduction in Sμ DSBs. Although we cannot determine the sites of the SSBs that gave rise to any given DSB, we were able to confirm that DSBs occur preferentially at AID hot spots even when the frequency of hot spots is low. We propose that only SSBs on opposite strands within a few nucleotides of each other can form a DSB in the absence of MMR. It is possible that MMR may play a significant role in DSB formation at other loci at which AID might initiate DSB formation, such as bcl6 and c-myc (51, 52). AID targets in these genes are infrequent and therefore, DSB formation, which can lead to translocation and tumor development, might be more dependent on MMR.

We thank Dr. Xiaoming Wu for unpublished data and the University of Massachusetts Flow Cytometry Facility for cell sorting and FACS analysis.

The authors have no financial conflict of interest.