R-loop editing by DNA cytosine deaminase APOBEC3B modulates the activity of oestrogen receptor enhancers

Domain: chemistry
Published: 2026-02-18
Editorial outcome: minor_revision

Abstract
Oestrogen receptor (ER) activation leads to the formation of DNA double strand breaks (DSB), promoting genomic instability and tumour heterogeneity. The single-stranded DNA cytosine deaminase APOBEC3B (A3B) serves as a co-activator of ER and is implicated in inducing DSBs at transcriptional enhancers regulated by ER. Using whole-genome sequencing in an engineered cell model lacking base excision repair (BER) function, we demonstrate that A3B preferentially targets transcriptionally active regulatory regions in an R-loop-dependent manner. Strand-specific DNA:RNA immunoprecipitation sequencing (ssDRIP-seq) and ssDNA-associated protein immunoprecipitation sequencing (SPI-seq) confirm that A3B binds to and deaminates ssDNA within R-loops, a process facilitated by ER transactivation. Furthermore, BER-mediated processing of A3B-induced uracil bases contributes to the formation of R-loop-associated DSBs, which are essential for ER-regulated gene activation. These findings establish a role for A3B in R-loop homeostasis and transcriptional regulation, with implications for understanding ER-driven genomic instability and potential therapeutic targeting of A3B.

Introduction
Oestrogen (17-βE2; E2) plays a crucial role in the development and maintenance of the human reproductive system and secondary sexual characteristics. Prolonged exposure to E2 has been implicated in the pathogenesis of breast, ovarian, and uterine cancers1. E2 exerts its effects by binding and activating the oestrogen receptor (ER). Upon binding E2, ER translocates to the nucleus and acts as a transcription factor, recognising E2-responsive elements (EREs) in genomic DNA. This event triggers altered expression of numerous downstream genes2. The set of genes regulated by ER activation encompasses those encoding essential reproductive functions, including cell proliferation and survival, and dysregulated ER activity has been associated with cancer development3.

In addition to its role in gene regulation, ER has been implicated in promoting cancer development by inducing DNA damage4. Importantly, short-term E2 stimulation in breast epithelial cells expressing ER leads to the formation of DNA double-strand breaks (DSBs)5,6, contributing to chromosome instability and aneuploidy7. ER-positive breast cancers exhibit increased genomic damage, characterised by mutations, alterations in gene copy number, and recombination8,9. ER-induced DNA damage is considered to be crucial for the activity of ER, since elements of the DNA repair pathway, such as transcription-coupled nucleotide excision repair (TC-NER) promote and participate in transcription activation10,11, The action of TC-NER also creates a favourable environment for transcription12 by facilitating DNA demethylation13, chromatin remodelling14, and enhancer-associated RNA (eRNA) synthesis15.

Various models have been proposed to elucidate the origin of oestrogen-induced DNA double-strand breaks (DSBs). These models include the involvement of DNA topoisomerase IIβ16, the formation and processing of co-transcriptional R-loops6, and DNA damage triggered and edited following deamination by apolipoprotein B mRNA-editing enzyme catalytic polypeptide-like 3B (APOBEC3B, A3B)5. A3B belongs to the APOBEC3 enzyme family, which comprises seven closely related DNA deaminases responsible for catalysing cytosine-to-uracil (C>U) editing of single-stranded DNA (ssDNA)17 or RNA. Apart from their innate functions as immune defences against DNA viruses and transposons18,19, several family members, including A3B, have been implicated in driving genetic heterogeneity in cancers by inducing C>T transitions and C>G transversions at 5’-TCW motifs (W=A or T)8,20. Elevated expression of A3B has been observed in more than 50% of primary breast cancers20. In addition, as the sole constitutively nuclear-localised member of the APOBEC3 family, A3B has emerged as one of the primary candidates underlying the APOBEC3 mutational signature identified in various cancers20,21.

Furthermore, the activation of A3B has been shown to induce DNA damage repair (DDR)9,22,23, where A3B directly interacts with ERα and edits ER binding sites, which are subsequently processed into DSBs by DNA base excision repair (BER) and non-homologous end joining (NHEJ) mechanisms5. The appropriate generation of these DSBs, along with subsequent epigenetic modifications, plays a critical role in determining the transactivation activity of ER5. Functional loss of A3B abolishes ER-driven cell proliferation and extends the therapeutic efficacy of tamoxifen in ER-positive xenograft models24. These findings underscore the role of A3B as an important regulator of ER activity.

Elevated levels of transcriptional activity result in the formation of chromatin-associated R-loop structures, characterised by the formation of a three-stranded structure comprising a DNA:RNA hybrid and a stretch of displaced ssDNA25. Previous research has indicated that the clearance of co-transcriptionally formed R-loops by the DDR machinery contributes to the generation of DSBs mediated by ER activation6. Given that the displaced ssDNA within R-loops could potentially serve as a substrate for A3B, it has been postulated that A3B may participate in R-loop-associated functions. This hypothesis gains support from studies utilising the APOBEC-Cas9 engineered system to edit cytosines in R-loop DNA26, a recent investigation revealing the influence of A3B on R-loop homoeostasis and consequent R-loop-associated mutagenesis27. However, despite these endeavours, it remains unknown whether the editing of R-loops by A3B is directly associated with its role as an ER coactivator in cells.

To comprehensively investigate the impact of A3B-mediated R-loop editing on the co-activation of ER-regulated transcription, we used genome-wide profiling of A3B editing sites in ER-dependent models. This approach offers insights into potential hotspots of A3B editing, shedding light on the underlying mechanism governing A3B-dependent DNA deamination and its regulatory role in ER activity. Previous large-scale whole-genome sequencing (WGS) studies conducted on cancer cell lines and primary human tumour samples have successfully identified APOBEC editing sites9,28. However, these investigations are potentially influenced by a survivorship bias, as the detection of A3B-edited sites is restricted to those that evade repair by BER, which rectifies U:G mispairing resulting from A3B editing29,30. This limitation will also impair studies employing A3B-overexpressing cell models31–34. To overcome this challenge, we employed an engineered BER-deficient breast cancer cell model to sample A3B editing sites and adopted a multi-omic approach to elucidate the intricate role of A3B in the molecular mechanism governing ER-regulated gene expression.

Results
Detection of A3B DNA deamination sites using a BER-deficient cancer cell model

To identify A3B deamination sites with potential functional impacts on protein coding or gene regulation, we conducted WGS of ER-positive T47D human breast cancer cells expressing doxycycline-inducible A3B. As a control, we adopted an enzymatically inactive A3BE68Q/E255Q double mutant (A3B**), which was confirmed to lack detectable enzymatic activity in multiple cell line models (Fig. 1a and Supplementary Fig. 1). Bicistronic expression of the humanised bacteriophage PBS2 uracil glycosylase inhibitor (hUGI) peptide was employed to inhibit BER-associated uracil-DNA glycosylase (UDG/UNG)20,30. We showed that lentiviral hUGI expression prevents the removal of A3B-induced uracil bases, allows more effective capture of A3B editing events, and it is sufficient to block the majority of UDG/UNG activity within cells (Supplementary Fig. 2). The selection of the T47D cell line was based on its TP53 loss-of-function mutation, which is known to confer protection against synthetic lethality induced by ectopic A3B expression35.

Doxycycline-induced expression of A3B/hUGI and effective uridine incorporation were confirmed by immunoblotting and slot blotting using Mycobacterium smegmatis UdgX as a uridine DNA sensor (Fig. 1a and Supplementary Fig. 3a, b). The cells were then exposed to doxycycline for five days, and genomic DNA samples were sequenced from five individual T47D clones expressing A3B-hUGI (A+ colonies, Fig. 1b). For control purposes, sequencing results from three uninduced colonies (A- colonies) were included to represent single-nucleotide variants (SNVs) arising from isolating individual clones. In addition, three T47D colonies co-expressing hUGI and the enzymatically inactive A3B** (M+ colonies) were also included. A3B** neither interfered with the activity of endogenous A3B (Supplementary Figs. 1, 2) nor triggered an unfolded protein response (Supplementary Fig. 4), indicating that it has no functional impact on the endogenous A3B enzyme. The M+ colonies were therefore expected to capture base-editing events mediated by endogenous APOBEC/AID family enzymes in T47D cells, predominantly A3B5. Slot blots with UdgX uridine DNA sensor confirmed that both the endogenous and induced wild-type A3B in the individual colonies retained full enzymatic activity (Supplementary Fig. 3c, d).

Joint mutation calling for the separated colonies was performed using a parental T47D reference sample. Analysis of the WGS data confirmed that the integrated lentiviral construct, including the A3B open‑reading frame and flanking vector sequence, was sequence‑intact, with no deep or non-synonymous mutations detected following A3B induction (Supplementary Table 1). Subsequent variant analysis revealed a significantly higher proportion of non-CpG C>T mutations in the A+ colonies compared to the M+ and A- colonies (Fig. 1c). To better characterise the nature and spatial distribution of these mutations, we analysed whether they occurred individually or as clusters. As indicated by previous studies, mutations can occur as clusters with various patterns, including large foci of close-together mutations also described as kataegis (thunderstorm) or alternatively, clusters of 3-4 mutations that occur in short distances known as omikli (fog)22. Studies using BER-functional models have detected prevalent kataegis-like clusters of mutations associated with increased A3B expression31,33. Here, we analysed intermutational distance for the detected mutations (Fig. 1d and Supplementary Fig. 5), and employed the HyperClust algorithm for the detection of localised somatic hypermutations22. Figure 1e clearly shows that the A+ colonies exhibit a greater number of non-recurrent and diffused clusters of mutations consistent with omikli rather than the previously reported kataegis. This observation suggests that the inhibition of BER in our cell model captures a distinct pattern of A3B-mediated mutations in alignment with deficient DNA mismatch repair frequently observed in human cancer genomes22.

Analysis of the resulting mutations against predefined COSMIC single-base substitution (SBS) signatures using non-negative matrix factorisation (NMF)28 identified enrichment of the SBS2 signature, attributed to the activities of the AID/APOBEC family of cytidine deaminases (Fig. 1f). De novo NMF signature extraction revealed two prevalent signatures among the sequenced control colonies: signature A, more prevalent in the control A- or M+ colonies, potentially accounting for the variation among T47D subclones, and signature B, primarily enriched in A+ colonies, characterised by the A3B-preferred 3’-TCW trinucleotide motif resembling the mutational process associated with A3B editing (Fig. 1g, h). In addition, analysis of the M+ colonies revealed a measurable but weaker enrichment of C>T substitutions at the A3B-preferred 3′-TCW trinucleotide motif and partial retention of SBS2 and Signature B. This effect, although attenuated compared with wild-type A3B induction, reflects deamination by the endogenous A3B enzyme whose activity becomes unmasked when BER is suppressed by the bi-cistronically expressed hUGI peptide. These data demonstrated that the BER-deficient breast cancer model captures A3B activity on genomic DNA in the absence of interference by downstream DNA repair mechanisms.

A3B preferentially deaminates genomic regulatory regions

We investigated the functional consequences of the captured A3B-editing sites using the Ensembl Variant Effect Predictor (VEP)36. Compared with random genomic locations, we observed a significant enrichment of TC>TT substitutions in A+ colonies within regulatory and transcription factor (TF) binding regions, but not in protein-coding regions. Intrinsic APOBEC-mediated TC>TT mutations in M+ colonies followed a similar but less significant trend, while TC>TT mutations arising from colony separation (A- colonies) displayed no obvious preference. Also, this enrichment was not observed for control VC>VT or ND>NN substitutions (Fig. 2a). To better understand the functional roles of these transcriptional regulatory regions, we constructed an 18-chromatin state ChromHMM model for the parent T47D cells, utilising previously published ChIP-seq data (Supplementary Fig. 6). We then examined the overlap between the captured mutations and specific chromatin states derived from this model using methods described previously37 (Fig. 2b). In the A+ colonies, we identified a significant overrepresentation of TC>TT mutations at transcription start sites (TSS) and enhancers, rather than main body (UTR, exon or intron) of the gene, particularly in the flanking regions of TSS and active enhancers. A similar pattern, although to a lesser extent, was observed for intrinsic APOBEC-induced mutations in M+ colonies. In addition, we analysed DNA replication timing using parent T47D Repli-seq data from the ENCODE dataset (ENCFF440QFG), which showed that the mapped TC>TT substitutions in A+ colonies, either as SNV or omikli events, were predominantly associated with early DNA replication sites (Fig. 2c). These findings align with previous observations indicating that A3B targets actively transcribed genomic regions and those involved in early replication22, and underscore the potential functional significance of A3B-associated modifications22. Overall, our data provide evidence of a direct association between A3B-mediated base-editing and sites of transcriptional regulation. These results are consistent with the previously established role of A3B5, and argue against a primary role for A3B’s DNA deamination activity in altering protein-coding sequences.

A3B binds and deaminates ssDNA in R-loops

Next, we sought to understand the sequence context flanking A3B-mediated mutations to determine features that may facilitate substrate recognition by A3B. In A+ colonies, we observed that TC>TT substitutions are accompanied by a pronounced GC-skewness in the flanking DNA sequences, indicating an overrepresentation of guanine bases in the same DNA strand as the TC>TT editing event (Fig. 3a). Furthermore, there was an enrichment of G-quadruplex-forming sequences within a ±500bp region surrounding A3B-editing sites (Fig. 3a). A similar, albeit weaker, effect was observed around TC>TT substitutions in M+ colonies, which represents the deamination by intrinsic A3B. In contrast, no such patterns were observed for TC>TT or other di-nucleotide substitutions derived from A- colonies (Fig. 3a and Supplementary Fig. 7). Both GC-skewness and G-quadruplex-forming sequences are known to facilitate the formation or stabilisation of R-loops38. Therefore, we explored the potential role of R-loops as substrates for A3B activity by first verifying their presence at A3B editing sites. We employed strand-specific DNA:RNA immunoprecipitation sequencing (ssDRIP-seq) combined with RNaseH-dependency to corroborate mapping DNA:RNA hybrids present in R-loops, GC-skewness, and MNase-seq analyses in parent T47D cells (Supplementary Fig. 8). ssDRIP-seq specifically captures and maps DNA:RNA hybrid structures, enabling genome-wide identification of R-loops39. The results identified significantly elevated R-loop signals near TC>TT substitutions in A+ colonies. Strand-specific sequencing revealed that the TT>TC substitutions occurred on the displaced single-stranded DNA (ssDNA) strand of the R-loop, consistent with A3B’s substrate preference for ssDNA (Fig. 3b). A similar but less pronounced enrichment of R-loop signals was found at TC>TT substitutions in M+ colonies, while no such signal was found in other di-nucleotide substitutions (Supplementary Fig. 9), confirming a similar site preference for intrinsic A3B base editing.

To further investigate the correlation between R-loops and A3B-mediated editing, we conducted immunoprecipitation experiments using the R-loop-binding S9.6 antibody, a well-established method for isolating R-loops and their associated proteins40. Similar to Top1, a validated R-loop binding protein40, A3B was found to co-immunoprecipitate with R-loops. This interaction was disrupted upon exogenous expression of RNaseH1, an enzyme that degrades RNA:DNA hybrids and removes R-loop structures (Fig. 3c).

To map the genomic occupancy of flag-tagged A3B and test its dependency on R-loops, we employed ssDNA-associated protein immunoprecipitation sequencing (SPI-seq) to capture A3B-bound ssDNA41 in Flag-A3B-expressing T47D cells, with or without co-expression of RNaseH1 to eliminate R-loop formation. We selected SPI-seq as this method would capture A3B-bound ssDNA in addition to dsDNA, the latter typically identified by conventional ChIP-seq. Following quality control analysis and validation of SPI-seq specificity by qPCR using T47D samples with or without A3B depletion (Supplementary Fig. 10a–c), we found the overexpression of RNaseH1 greatly reduced the number of Flag-A3B peaks captured and hindered Flag-A3B’s binding to both promoter and enhancer regions (Supplementary Fig. 10b–f). A close inspection of the Flag-A3B peaks eliminated by RNaseH1 co-expression revealed enrichment of R-loop sequences and G-quadruplex motifs relative to the unaffected peaks (Fig. 3d). Notably, SPI-seq captured A3B-bound ssDNA fragments were found to be localised within the body of R-loops, the signal abundance of which can be affected by RNaseH1 expression (Fig. 3e). We also observed a more pronounced decrease in A3B SPI-seq peaks signal in the neighbourhood of R-loops upon RNaseH1 expression (Fig. 3f), indicating that A3B binding sites proximal to R-loops are more susceptible to R-loop clearance, thus demonstrating a dependency of A3B’s ssDNA binding on R-loop formation. In addition, enrichment of the A3B binding signal can be observed in previously identified A3B-driven TC>TT substitutions, the strength of which was attenuated upon R-loop clearance by RNaseH1 overexpression, suggesting C>T DNA editing by A3B can also be affected by R-loop availability in cells (Supplementary Fig. 11). Collectively, our ssDRIP-seq and SPI-seq data support the notion that R-loop formation facilitates A3B binding and cytidine deamination of genomic DNA.

ER-induced R-loop formation facilitates A3B binding

Given that the activation of ER triggers the co-transcriptional formation of R-loops, which can influence gene expression6, we postulated that increased levels of genomic R-loops would enhance the availability of substrates for A3B and subsequently facilitate A3B binding to ER-activating enhancer regions. To investigate this, we employed SPI-seq and ssDRIP-seq to map the binding sites of flag-tagged A3B and R-loops responsive to E2 stimulation in T47D cells, enabling the examination of their spatial-temporal relationship. Here, we employed quantitative SPI-seq (with n=2 replicates) to identify genomic regions bound by Flag-tagged A3B and quantitative ssDRIP-seq (with n=3 replicates) to map R-loop formation in T47D cells treated with or without E2 (Fig. 4a and Supplementary Fig. 12), allowing us to analyse their spatial-temporal relationship. Consistent with previous findings5,6, we observed a global increase in A3B binding (23,711 peaks; 53% of all peaks) and R-loop formation (18,567 peaks; 16% of all peaks) after two hours of E2 exposure (Fig. 4b). A closer inspection found that A3B binding sites were enriched for promoter and enhancer sites irrespective of E2 treatment (Supplementary Fig. 13). In contrast, only R-loops induced by E2 treatment showed a trend of enrichment near TSS regions, whereas R-loops formed independently of E2 did not (Supplementary Fig. 14). Remarkably, a substantial portion of newly-formed E2-dependent A3B binding sites were found in proximity to R-loops following E2 exposure (p<10−5, χ2 test), and correspondingly, R-loops near A3B binding sites exhibited increased sensitivity to E2 induction (Fig. 4b). A substantial overlap was seen between E2-induced A3B binding sites and R-loops (Fig. 4c), as well as a significant enrichment of E2-induced R-loops in proximity to E2-induced A3B binding sites (p<10−5, χ2 test, Fig. 4d). These observations strongly suggest a link between R-loop dynamics and A3B’s substrate binding following E2 treatment.

Our data suggest a potential causal relationship underlying R-loop formation and A3B binding at sites of E2-induced gene expression. However, depletion of A3B using siRNA did not impact the overall levels of R-loop signal or the increase in R-loops upon E2 induction determined by S9.6 antibody-based slot blotting (Supplementary Fig. 15). Furthermore, quantitative ssDRIP-seq analysis revealed an increase in R-loops proximal to A3B binding sites despite A3B knockdown (Supplementary Fig. 16a). By employing the DESeq2 quantification method featuring two-factor interaction analysis42, we quantitatively examined the combinatorial effects of A3B siRNA treatment and E2 induction on R-loop formation (Supplementary Fig. 16b), which showed minimal interaction between these two factors, indicating that A3B depletion is unlikely to directly affect E2-induced R-loop formation. On the other hand, more than half of the E2-induced A3B binding sites were found either located near (<2kb) E2-induced R-loops (34%) or pre-existing R-loops prior to E2 treatment (28%, Fig. 4e), with R-loop-proximal A3B peaks displaying a higher degree of induction upon E2 treatment compared to R-loop-distal A3B peaks (Supplementary Fig. 16c).

These findings suggest that elevated A3B binding is unlikely to drive the induction or formation of R-loops, but instead, the availability of R-loops may dictate E2-induced A3B binding and editing chromatin in or adjacent to the R-loop. To further examine this, we investigated the impact of RNaseH1 overexpression on A3B binding in seven loci characterised by well-defined ER binding and R-loop formation (Fig. 4f). In contrast to the two A3B binding sites lacking R-loops at promoters of TNFSF8 and RMST, clearance of R-loops by RNaseH1 overexpression impaired E2-induced A3B binding (Fig. 4g and Supplementary Fig. 17) in T47D and MCF7 cells. These results confirm that the binding of A3B to specific regions induced by E2 exposure involves R-loop serving as A3B substrates.

Our findings suggest a model where the formation of R-loops remodels the genomic landscape surrounding EREs, facilitating the recruitment of A3B. Using our ChromHMM model, we identified that E2-induced A3B binding sites overlapping with R-loops exhibit high enrichment scores in regions proximal to transcription start sites (TSS), TSS flanking regions and transcribed enhancers (Fig. 4h). Importantly, we found that E2-induced A3B binding sites showed a distinct preference toward transcriptional enhancer sites undergoing active transcription, which was not observed at other A3B binding sites. This coincides with the finding that E2-induced R-loops display the highest enrichment in actively transcribed enhancer regions. In addition, analysis of transcription factor (TF) binding DNA motifs surrounding E2-associated A3B binding sites reveals a distinct set of TFs that may be enriched in E2-regulated R-loop proximal peaks, particularly characterised by G-rich sequence motifs, in contrast to R-loop-distal A3B peaks (Supplementary Fig. 16d). This observation further supports the notion that formation of R-loops, facilitated by G-rich motifs, may actively reshape the localisation of the A3B binding site. Overall, our data strongly suggest that the increased availability of R-loops induced by ER transactivation could regulate A3B binding.

A3B promotes DSB formation at E2-induced R-loops

Several investigations have demonstrated that the formation of double-strand breaks (DSBs) induced by ER transactivation can result from the processing of co-transcriptionally formed R-loops or A3B editing sites through DDR mechanisms5,6. However, whether these two processes are functionally linked remains unclear. To explore the interconnection between these processes, we employed DSBCapture-seq to accurately map genomic DSB sites43 and investigate the dependence of E2-induced DSBs on A3B binding and editing and/or R-loop formation. T47D control and A3B-depleted cells were exposed to DMSO or E2 for a duration of two hours, followed by DSBCapture-seq analysis (Fig. 5a). Differential peak analysis of the sequencing outcomes using DiffBind44 revealed two categories of DSBs: E2-induced DSBs and A3B-modified DSBs (Fig. 5a). Our results corroborate the previously reported extensive DNA damage triggered by ER activation4–6,45 and demonstrate that the 15,259 E2-induced DSB peaks account for 30% of all detectable DSBs in the basal parent T47D cell line genome. 5308 DSB peaks were modified by A3B depletion, with the majority significantly (88%, p<105 by Χ2 test) overlapping with the E2-induced DSBs (Fig. 5b). This enrichment of A3B-modified DSBs within the E2-induced DSB set aligns with previous reports5. Support for the involvement of A3B in these DSBs came from our evidence that nearly all (98.6%) of the A3B-modified DSB sites we detected overlapped with the E2-induced A3B binding sites we identified through quantitative SPI-seq (Fig. 5c). However, A3B binding alone was not sufficient to predict A3B-dependent DSB formation, as 74% of the E2-induced A3B-independent DSBs also overlapped with A3B binding sites, suggesting there was an additional determinant for A3B dependency. We also observed a prerequisite for R-loop formation preceding A3B binding in the generation of A3B-modified DSBs, since a significant overlap (p<105 by Χ2 test) with R-loops was detected for these A3B-dependent DSBs compared to the A3B-independent E2-induced DSBs (Fig. 5c). Accordingly, both R-loop formation and A3B binding served as distinguishing factors between A3B-modified DSBs and A3B-independent DSBs (96% vs. 42% of all DSBs, Fig. 5d) and were reliable indicators of the impact of A3B depletion on DSB formation (Fig. 5e, f). Thus, our findings support the involvement of R-loop formation in the generation of A3B-induced DSBs.

Upon closer examination of the regions flanking DSBs, distinctive features associated with A3B-dependent DSBs emerged, compared with those at A3B-independent sites. These features included heightened bidirectional transcription activities (determined by global run-on sequencing, GRO-seq), increased occupancy of RNA polymerase II (determined by PolII ChIP-seq), and the presence of denser G-quadruplex-forming sequences (determined by G4Hunter sequence analysis), all of which promote R-loop formation46 (Fig. 5g). Utilising genome-wide analysis with our ChromHMM model, we observed that A3B-dependent DSBs exhibited greater enrichment at active or flanking transcription start sites (TSS) and transcriptionally active enhancers (Supplementary Fig. 18). We also found these same chromatin states, characterised by transcriptional activity, were enriched for E2-induced R-loops, suggesting a reliance on R-loops during the conversion of A3B editing sites into DSBs (Fig. 4h).

The observed association between A3B binding and transcriptionally active R-loop regions raised the possibility that the formation of A3B-dependent DSBs involves mechanisms linked to active transcription, such as TC-NER. The increased transcriptional activity and RNA polymerase II occupancy at A3B-dependent DSBs could thus provide substrates and conditions favourable for TC-NER, which has been implicated in R-loop resolution and subsequent generation of single-strand breaks (SSBs)6. To investigate this further, we analysed seven A3B binding sites that overlapped with R-loops and were associated with A3B-dependent DSBs (Fig. 5h). We examined the impact of RNAi-mediated depletion of Cockayne syndrome group B (CSB) protein and XPG/XPF endonucleases, which are components of TC-NER required for the conversion of R-loops into DNA strand breaks47. Using quantitative PCR analysis of DSBCapture-enriched DNA samples, we discovered that depletion of these TC-NER components was sufficient to reduce the formation of E2-responsive A3B-dependent DSBs in T47D and MCF7 cells (Fig. 5i, Supplementary Fig. 19). Single depletion of either XPG or XPF caused only a slight reduction in E2-induced A3B-dependent DSB formation, whereas combined depletion almost completely abolished DSB production at all examined loci (Supplementary Fig. 20). Because XPF and XPG can incise either strand of the R-loop without strict strand preference, the modest effect of single depletion suggests functional redundancy between these endonucleases, each being individually capable of mediating the incision events required for R-loop processing and subsequent DSB formation. These findings demonstrate that A3B-driven DSB formation requires cooperative activity with the TC-NER machinery.

Blocking the processing of A3B editing sites into DSBs impairs E2 response

The genomic localisation of A3B binding sites, particularly those in proximity to R-loops and associated with double-strand break (DSB) formation, implies a regulatory role for the downstream processing of A3B-mediated base editing. To examine the functional impact of blocking this processing step, we performed RNA-seq analyses using T47D cells containing the doxycycline-inducible, catalytically inactive A3B** mutant expressed together with hUGI (Fig. 6a). As shown previously (Fig. 1 and Supplementary Figs. 1–4), A3B** has no detectable enzymatic activity and does not interfere with endogenous A3B, thereby serving as a non-catalytic control that allows assessment of uracil repair-dependent processes. By co-expressing hUGI, we effectively blocked the conversion of A3B-induced uracil lesions into repair intermediates that give rise to DSBs (Supplementary Fig. 2). Following exposure to E2 for two hours, we collected RNA samples from cells with or without doxycycline induction of A3B** and performed RNA-seq. To assess differential gene expression, we employed the interaction modelling approach using the DESeq2 programme to examine the impact of loss of endogenous A3B activity on the E2 response42. Our results demonstrated that the expression of A3B**-hUGI dampened the E2-regulated response of 87% of genes (Fig. 6b), with 90 genes showing statistically significant interaction models based on DESeq2 analysis (Fig. 6c). Gene set enrichment analysis (GSEA)48 using the MSigDB hallmark genes49 identified a significant overlap between the 90 E2-responsive genes whose regulation is modulated by A3B-hUGI expression and those regulated by oestrogen response pathways (Fig. 6d), suggesting the involvement of uracil repair of A3B-mediated DNA deamination sites during ER transactivation.

To examine the impact of A3B-mediated DSB and A3B-bound R-loops on gene expression, we employed the predictive algorithm rGREAT to generate two groups of genes associated with previously identified regions (Fig. 7a). The first group comprised 86 genes associated with A3B-dependent DSBs, while the second group comprised 333 genes associated with E2-induced, R-loop-proximal A3B binding sites. Applying leading-edge analysis to the expression of these two gene groups identified a significant overlap with genes up-regulated by E2 (Fig. 7b). Importantly, genes from both groups were also enriched among genes whose E2 responsiveness was dampened upon expression of the bicistronic A3B**-hUGI construct, in which the hUGI peptide primarily exerts its effect by blocking BER processing of A3B-induced lesions. (Fig. 7b). These data are consistent with a mechanism of A3B-dependent DSB formation facilitating ER-regulated gene transactivation.

To further validate this hypothesis, we examined the effects of hUGI induction and A3B depletion on DSB formation, in parallel with expression of associated genes. Two groups of genes were analysed, E2-responder genes modified by A3B**-hUGI expression and those unaffected (Fig. 7c). Consistent with our model, we found that both depletion of A3B and blocking BER repair of its deamination sites prevented A3B-dependent DSBs from forming, leading to modified E2 responses in T47D (Fig. 7d) and MCF7 (Supplementary Fig. 21) cells. In contrast, no changes in E2 responses were seen for gene-associated A3B-independent DSBs despite A3B depletion or BER blockade (Fig. 7d and Supplementary Fig. 21). Collectively, our findings indicate that the processing of A3B-mediated C>U editing sites at R-loops, leading to DSB formation, is required for the E2 response of a subset of ER-regulated genes in cells.

To sum up, the results presented in this study support a model of how ER transactivation leads to DSB formation and its functional implications for gene transcription with the following steps (Fig. 8): (i) Upon ER activation, A3B is recruited to transcriptionally active enhancers and promoters via its interaction with ER. (ii) At these sites, A3B catalyses cytidine deamination on the exposed ssDNA, generating uridine lesions and initiating strand nicks. (iii) In parallel, the TC-NER endonucleases XPF and XPG, which can incise either the displaced DNA strand or the DNA:RNA hybrid, act cooperatively to introduce SSB. The convergence of these A3B- and TC-NER-mediated incisions results in DSB formation at E2-responsive enhancers. (iv) Repair of these DSBs through DDR pathways triggers chromatin remodelling and establishes epigenetic marks favourable for sustained transcription of ER target genes.

Methods
Cell culturing and cell line preparation
T47D (ATCC HTB-133) and SKBR3 (ATCC HTB-30) cells were obtained from ATCC. Lenti-X 293T cells were obtained from Clonetech. Cells were maintained in ATCC-recommended medium, except that Lenti-X 293T cells were cultured in DMEM supplemented with 10% v/v FBS (PAA) and 0.5% v/v pen/strep. All cells were cultured at 37°C with 5% CO2.

To generate stable inducible cell lines, T47D cells at the exponential growth phase were transfected with lentivirus at an M.O.I of 100, with 10µg/ml polybrene. After a 48h transduction period, selection was performed in medium supplemented with 4µg/mL puromycin (Gibco). Transduction efficiency was monitored by either turbo-RFP expression, A3B immuno-fluorescence microscopy or immunoblotting following induction of sample cells with doxycycline. After verification, lentivirus cassettes were maintained by culturing cells with 4μg/mL puromycin.

Drug treatments
For the induction of lentiviral protein expression, cells were pre-exposed to full culture medium supplemented with 1µg/mL doxycycline (Sigma) for 24h unless denoted otherwise. Follow-up drug treatments, if included, also contained the same doxycycline concentration throughout the entire treatment period.

For ER induction by E2, adherent cells were briefly washed with warm PBS twice before incubating with phenol-red-free RPMI-1640 medium (Gibco) supplemented with 10% charcoal-stripped FBS (Thermo Fisher) for 48h at 37°C. E2 was added to the culture medium at a final concentration of 100 nM, and further incubated for two hours (unless denoted otherwise) before the cells were harvested for follow-up studies.

Colony separation and WGS
0.25 million exponentially growing T47D cells harbouring lentiviral 3×Flag-A3B-P2A-hUGI-HA or 3×Flag-A3B**-P2A-hUGI-HA were seeded onto a 100mm plate for 24h, followed by induction with 1µg/ml doxycycline for 120h. Cells were harvested using trypsin digestion, serially diluted, and plated onto a 96-well plate (Corning) according to the manufacturer’s protocol. Colonies derived from single cells were picked, and the cloned cells were expanded with culture medium supplemented with 4µg/ml puromycin. Genomic DNA was extracted from each colony using a Qiagen Genomic-tip 500/G kit. Sequencing libraries for whole genome sequencing with DNA nanoball (DNB) technology were constructed (Beijing Genomics Institute (BGI) Inc., Hong Kong). Sequencing was carried out using BGISEQ-500 sequencer with a mean sequencing coverage of greater than 30× for each of the samples using 2×150bp configuration.

Plasmids cloning and lentivirus packaging
DNA encoding 3×Flag-A3B-P2A-hUGI-HA were synthesised by GeneArt service (Thermo Fisher Scientific Inc.). In order to distinguish the mRNA of exogenous proteins from their endogenous and boost their expression, codons for both A3B and hUGI were modified using the GeneArt design algorithm. AgeI restriction site and Kozak sequences were introduced at the 5’-end, and BspD1 restriction site was introduced at the 3’-end of the construct using PCR. An empty pTRIPZ plasmid was obtained from Horizon Discovery. The DNA fragment was introduced into the pTRIPZ plasmid using AgeI and BspD1 restriction digestion and ligation with enzymes from New England Biolabs (NEB). For the A3B** construct, two rounds of site-directed mutagenesis at E68Q and E255Q sites on A3B were performed on 3×Flag-A3B-P2A-hUGI-HA-containing pTRIPZ plasmid using the Quick-Change kit provided by Agilent. The resultant plasmids were sequenced and maintained in Mach1 T1 E. coli cells.

For lentivirus packaging, helper plasmids psPAX2 and pMD2.G were co-transfected with pTRIPZ encoding the protein of interest into Lenti-X 293T cells (Clonetech) using a calcium phosphate transfection kit (Promega). After changing the culture medium after 24h, the virus was harvested at 48 and 72h, pooled, precipitated using Peg-it reagent (System Biosciences) and resuspended in serum-free RPMI medium (Gibco). Multiplicity of infection (M.O.I) of viruses was measured according to the manufacturer’s recommendation (Horizon Discovery).

Immunoblotting
Cells were lysed in RIPA buffer (Millipore) containing complete protease inhibitor cocktail (Roche) and phosphatase inhibitors solution 2 (Sigma). Protein concentrations were measured by the BCA protein assay (Pierce). Samples were separated on SDS polyacrylamide gels and transferred to PVDF membranes using the iBlot2 system (Invitrogen). Membranes were washed with TBST (25mM Tris, 140mM NaCl, 0.1% Tween-20, pH 7.5) and probed with various primary antibodies overnight at 4°C, followed by binding of IRDye 680- or IRDye 800CW-conjugated secondary antibodies (LI-COR) for 1 hour at room temperature. Imaging and quantification of protein bands were performed using an LI-COR CLX infrared imaging system.

UdgX preparation and slot blotting
DNA encoding M. smegmatis UdgX protein with N-terminal 6×His-tag and C-terminal AviTag was synthesised by GeneArt (Thermo Fisher) and cloned into the pET24a plasmid. Biotinylated UdgX protein was expressed in the AVB101 E. coli strain, and sequentially purified using Ni-NTA and Superdex 75 columns on an ÄKTA Avant protein purifier.

Uracil-incorporated positive control DNA was obtained by extraction of pUC19 plasmid which was transformed into the K12 CJ236 E. coli strain (NEB). Genomic DNA from T47D cells was extracted and purified using the Qiagen Genomic-tip 500/G kit according to the manufacturer’s instructions. Sample DNA (100 ng for positive control and 4µg for genomic DNA) was loaded onto a PVDF membrane with Hoefer PR600 slot blot manifold linked to a vacuum pump. The membrane was blocked by Pierce protein-free blocking buffer (37572), followed by incubating with 1µg/ml biotinylated UdgX at room temperature for 1h. After probing with streptavidin-HRB conjugate (Pierce), the membranes were washed with TBST and detected with LI-COR CLX infrared imaging system using ECL-plus substrate (Pierce).

SNV calling and analysis
Raw sequences from BGISEQ-500 sequencer were aligned to the human genome (hg38) Burrows-Wheeler Aligner (BWA)62. The resultant bam files were then recalibrated using GATK 4.0.5 mutation calling kit following the ‘best practice’ advised by the developers63. Briefly, bam reads were reordered, marked with group and duplication status with Picard tools, followed by base quality score recalibration by the GATK BaseRecalibrator programme. Joint variant calling was performed using GATK Mutect2 programme, with doxycycline-induced colonies as ‘Tumour’ sample and non-induced colonies as ‘Normal’ sample. In order to improve mutation calling accuracy, resources common germline variant sites from Genome Aggregation Database (gnomAD) and a ‘panel-of-normal’ dataset consisting of several wild-type T47D mutation calling results were included in the Mutect2 analysis64. After calculating the potential sampling contaminations using the GATK CalculateContamination programme, the Mutect2-derived SNVs were then selected against 14 filters implemented by the GATK FilterMutectCalls. In order to exclude SNVs that are located on either ends of homopolymer DNA sequences and within simple repeating elements, both of which are likely to generate false-positive results, the SNVs passing through GATK filters were further filtered using SNPiR software65 with perl scripts filter_homopolymer_nucleotides.pl and pblat_candidates_ln.pl scripts only. The latter script was enabled by the pBLAT algorithm66 and the BEDTools programme67.

The resultant SNVs, encoded in vcf file format, were subjected to further analyses using programmes coded by R language (R Core Team). Calculation and visualisation of intermutational distance using a waterfall plot were performed with the R package ‘qqman’, with minor modifications to the source code68. Kataegis and omikli mutation clusters were detected using the hyperclust programme, as described previously22. The effect of variants was analysed using Variant Effect Predictor (VEP)36, with the assistance of the genome assembly lift-over tool provided by UCSC Genome Browser69. Random sampling and enrichment analysis by Fisher exact test for variant effects were conducted using bedtools. For replication timing analysis for SNVs, a method described previously22 was employed using the T47D Repli-seq dataset from ENCODE (ENCFF440QFG).

Mutational signature analysis
All SNVs detected in the M+, A+ and A- groups were used for identifying mutational signature contributions. Mutation signatures and spectrum analysis were analysed by the Bioconductor package MutationalPatterns with 30 COSMIC signatures following the standard workflow70. De novo mutational signature extraction was performed using the NMF algorithm71. Cosine similarities between de novo-extracted signatures and the COSMIC cancer mutational signatures were also calculated using MutationalPatterns. The absolute and relative contributions of each de novo signature were also determined.

Chromatin state analysis
Chromatin state characterisation was conducted using ChromHMM software37. To provide modelling input, raw reads from ChIP-seq results using six chromatin marks were downloaded from the GEO database and aligned to the GRCh37 (hg19) genome assembly using BWA. The resultant BAM files were binarised and used as a training set to build the Hidden Markov Model that segments and annotates the T47D genome. To validate the chromatin model generated, the degree of overlap between known genomic features and the model was carried out using a script in ChromHMM. The genomic features were predefined by the ChromHMM authors, except that the super-enhancer regions in T47D cells were derived as previously reported72. For computing the overlap between ChromHMM and genomic regions of interest, the full-length peak regions were used except for SNVs, where ±200 bp flanking regions were used.

Strand-specific RNA/DNA immunoprecipitation (ssDRIP) and sequencing
RNA/DNA immunoprecipitation was performed on T47D cells using an optimised method previously reported by Halász et al.73. The entire procedure followed the best-performing experiment number 5 in the report, the detailed protocol of which was provided in the supplementary information section, except that the Covaris E220 was used for fragmenting genomic DNA with the following settings: duty factor 10%; peak incident power 75 watts; cycles per burst 1000; time 50s; temperature 4°C. S9.6 antibody was purchased from Kerafast (ENH002), and mouse IgG (Thermo Fisher 31903) was used as a control (if required).

ssDRIP-seq was performed using immunoprecipitated DNA:RNA hybrids. To enable strand-specificity, the product was subjected to the second-strand DNA synthesis with uracil incorporation. More specifically, the DNA:RNA hybrid was reconstituted in 1× NEB first-strand synthesis buffer (NEB E7525) without the addition of primer and enzyme to a final volume of 20µl. The product was immediately used as the starting material of the directional second-strand synthesis method following the manufacturer’s instructions (NEB E7550). The resultant DNA was purified using SPRI beads (Thermo Fisher) at a volume ratio of 2.0 and eluted with TE buffer. After quality inspections with the BioAnalyzer 2100 (Agilent), the product was sent to BGI in Hong Kong, where strand-specific libraries were constructed using DNB technology. The resultant libraries were sequenced with the BGISEQ-500 instrument.

Data processing of ssDRIP-seq
Pair-end raw reads were aligned to the GRCh37 genome assembly using BWA, followed by file format exchange by SAMtools. The resultant alignment reads were separated by first-in-pair strand into two files corresponding to (+) or (-) strand R-loop (or input) reads using SAMtools flag identifiers. Peak calling was performed with the MACS2 package using the pair-end fragment setting for the input and ssDRIP samples. Signal tracks were generated using MACS2, with the score representing fold enrichment relative to the input control.

For quantification of ssDRIP experiments, consensus R-loops peaks were generated first. The resultant peaks for 12 replicates were served as input for MSPC, and recurrent peaks were filtered using parameters ‘-w 1E-4 -s 1E-8 -c 4’. Peaks on the same strand within 1000bp were merged using BEDTools, and converted to GTF format using UCSC tools (bedtogenepred and genepredtogtf). Read counting was conducted using Subread, and raw counts were analysed by DEseq2 using a generalised linear model with an interaction term.

GC skewness and G4 motif analysis
GC-skewness was calculated as previously described. Genomic GC-skew data, which is encoded as a bigwig file format with a window length of 200bp, was created by the R-loop DB74 and downloaded from the UCSC genome browser69.

The DNA G4 motif was predicted using G4Hunter75, an R package. A scan was performed using a G4 score threshold of 1.2 on the GRCh37 genome assembly, followed by composing the resulting file in bigwig format with a window length of 100bp for follow-up studies using the R scripts provided by the authors of G4Hunter.

Sequencing data visualisation
Bigwig files containing enrichment scores, GC-skewness score and G4 motif frequency across the GRCh37 genome assembly were used to plot the heatmaps and profile plots. The R package ‘seqplots’ was used to perform data visualisation via a graphical user interface76. For heatmaps, average data using a window length of 50 was used, whereas for profile plots, average data with standard deviation using a window length of 100 was used. Signal profiles across specific genomic regions were visualised by the Integrative Genomics Viewer (IGV)77.

S9.6 co-immunoprecipitation
S9.6 co-immunoprecipitation was conducted using the method as previously reported40, except that S9.6 antibody was purchased from Kerafast (ENH002) and IgG control from Thermo Fisher (31903). Immunoblotting was carried out using S9.6-co-immunoprecipitated proteins, with HRP-conjugated secondary antibodies and SuperSignal West Femto substrate (Thermo Fisher 34094), and visualised by LI-COR CLX infrared imaging system.

RNA/DNA hybrid slot blots
Genomic DNA from T47D cells was extracted and purified using the Qiagen Genomic-tip 500/G kit following the manufacturer’s instructions. 8µg DNA was treated with or without RNaseH (NEB M0297) at a final reaction volume of 400µl overnight at 37°C and loaded onto a pre-wetted Hybond N+ nylon membrane (Cytiva, RPN303B) with Hoefer PR600 slot blot manifold linked to a vacuum pump. The membrane was immediately transferred to a Stratalinker UV cross-linker (model 1800 Stratagene) and crosslinked using a total energy of 120,000 microjoules. The membrane was then wetted with TBST, and blocked with StartingBlock T20 blocking buffer (37543) at 4°C overnight, followed by probing with S9.6 antibody and anti-mouse-HRP conjugate (Cell Signalling 7076). The membrane was visualised using the LI-COR CLX infrared imaging system with SuperSignal West Femto HRP substrate (Thermo Fisher 34094).

Protein transient over-expression and siRNA transfection
ppyCAG_RNaseH1_WT plasmid was used for transient over-expression of RNase H1 protein and was a gift from Xiang-Dong Fu (Addgene plasmid #111906). This plasmid encodes V5-tagged human RNase H1 without the first 27 amino acids and replaced with a nuclear localisation signal. For protein transient overexpression, T47D cells were plated at a density of 5×104 cells per well in a 6-well plate and incubated for 24h. After brief washing with warm PBS and replacing the medium with RPMI-1640 medium supplemented with 10% charcoal-stripped FBS, 1.5µg of the vector was transfected into the cells by using Lipofectamine LTX PLUS (Invitrogen A12621) according to the manufacturer’s instructions. Cells were then treated with DMSO or E2 and proceeded with subsequent experiments.

For RNAi experiments with siRNA transfection, the same cell culture preparation and follow-up experimental procedures were used as plasmid transfection, except that Lipofectamine RNAiMAX transfection reagent was used according to the manufacturer’s instructions, and plasmid DNA was replaced with 20nM siRNA. For DSBCapture experiments, cell culture and transfection reagents were scaled up to 100mm plates according to manufacturer’s recommendations.

For RNAi experiments with siRNA transfection, the same cell culture preparation and follow-up experimental procedures were used as plasmid transfection, except that Lipofectamine RNAiMAX transfection reagent was used according to the manufacturer’s instructions, and plasmid DNA was replaced with 20nM siRNA. For DSBCapture experiments, cell culture and transfection reagents were scaled up to 100mm plates according to manufacturer’s recommendations.

Chromatin immunoprecipitation (ChIP)
ChIP protocol used in this study was adapted from the protocol used by Myer’s Lab at Hudson Alpha (protocol version v011014) which contributed to the ENCODE project8 with the following modifications. Firstly, fixed T47D cells were lysed in ChromaTrap lysis buffer designed for sonication (Porvair Science, 100001), and chromatin fragmentation was carried out on a Covaris E220 instrument with optimised settings for ChIP (duty factor 2%; peak incident power 105 watts; cycles per burst 200; time 20mins; temperature 4°C). The resultant chromatin was digested with proteinase K (Qiagen), and the DNA concentration was measured with a Qubit fluorometer (Thermo Fisher). Secondly, Pierce ChIP-grade protein A/G beads (Pierce 26162) were used for immunoprecipitation, and chromatin binding to beads was performed at 4°C overnight. Thirdly, for Flag-M2 antibody, optimised compositions of wash buffers are used and are as follows: 100mM Tris pH 7.5; 500mM LiCl; 1% NP-40; 1% sodium deoxycholate. Finally, the resultant immuno-precipitated DNA was purified by SPRI beads (Thermo Fisher) using a ratio of 2.0 and analysed using a BioAnalyzer 2100 (Agilent).

Single-strand DNA-associated protein immunoprecipitation (SPI)
SPI was carried out following the developers’ literature with modifications41 and using the same method used to capture chromatin-bound DNA as described in the previous section, except that the resultant DNA was denatured at 95°C and immediately quenched on ice.

Sequencing and data processing for ChIP/SPI-seq
ChIP and SPI samples were sent to Beijing Genomics Institute (BGI Inc.) for library construction and sequencing. For analysis of ChIP samples, a standard pipeline for the construction of DNB library was used, whereas for SPI samples, the strand-specific DNB library construction protocol was requested in order to include ssDNA species in the library. Sequencing was carried out on a BGI-500 instrument with single-end 50bp reads for both input and immunoprecipitated DNA.

For data analysis, raw reads were aligned to the GRCh37 genome assembly using BWA, followed by file format exchange by SAMtools78. Peak calling was carried out using the MACS279 package using paired input/IP sequences with modelled narrow peak settings and an extended fragment length of 200bp. Enrichment scores were computed by MACS2 using the resultant files from peak calling and represented as a fold change of IP over control. For n=2 replicated ChIP/SPI-seq data, reproducible peaks were picked using MSPC80. Peaks were filtering against a ‘black list’ region defined by the ENCODE project8. For quantitative analysis of the effect of E2 over DMSO on SPI-seq data, EdgeR81 statistics were performed with the R package Diffbind82, using a narrow peak produced by MACS2 and aligned sequence reads. To filter for the peak of interest, the top 25% peaks by base mean score from normalised sequencing read counts were included for the downstream analyses. Significant peaks were defined as FDR≤0.05 with fold change≥1.5 by Diffbind. Intersection between peak regions and Fisher’s exact test with randomised genomic regions was performed by BEDTools, and data were visualised by the R package SeqPlots. Transcription factor binding motif analysis was carried out using the AME83 programme from the MEME84 suite. The distance between peak features was analysed by BEDTools67, where proximity was defined as ±1.5 kb of distance. For genome-wide analysis of the ChIP-seq dataset, the R package ChIPseeker was used.

DSBCapture-seq
DSBCapture was conducted following a previous report43 except that the Covaris E220 was used for fragmenting adaptor-ligated DNA with the following settings: duty factor 10%; peak incident power 75 watts; cycles per burst 1000; time 50s; temperature 4°C.

Data processing for DSBCapture-seq
DNA libraries generated by the DSBCapture protocol were sequenced on an Illumina HiSeq 2500 by BGI. Raw reads were aligned to the GRCh37 genome assembly using BWA and processed by SAMtools. MACS2 was used to perform peak calling, with modelled narrow peak settings and an extended fragment length of 200bp. Signal tracks were generated by MACS2, with the score representing fold enrichment relative to the input control. For quantification, EdgeR analysis was performed with Diffbind, using the same procedure as for ChIP-seq. The heatmap was created using the R package ComplexHeatMap85.

RNA-seq and quantification
Total RNA from T47D cells were extracted using the MagNA pure 96 instrument (Roche), and RNA integrity number (RIN) was determined by BioAnalyzer 2100 (Agilent). RNA libraries were constructed and sequenced by BGI using poly-dT enrichment in conjunction with DNB technology on the BGISEQ-500 instrument. Raw reads were aligned to GRCh38 genome assembly and GENCODE86 GRCh38.p13 annotation using STAR programme87, followed by processing of the resultant file using SAMtools. Read counting on genomic features was carried out using Rsubread88, and the R package DESeq242 was used to perform statistic-based quantification. In DESeq2 analysis, in addition to terms to denote doxycycline and E2 effects in the design formula, an additional interaction term was added to denote the synergism effect between the two drugs following the user’s manual written by the DESeq2 authors (http://www.bioconductor.org/packages/devel/bioc/vignettes/DESeq2/inst/doc/DESeq2.html). Expressed genes in T47D cells were defined as those with a base-mean normalised read count greater than 50% of all genes. Differentially expressed genes were picked using an FDR cut-off of 0.05 and a fold change cut-off of 1.5. The heatmap was created using ComplexHeatMap.

Gene set analysis
Gene set analysis was carried out using MSigDB49,89 with gene set collection H using differentially expressed genes identified by RNA-seq. Cis-regulatory region-associated genes were predicted using rGREAT48 using differential binding peaks identified by SPI-seq and DSBCapture-seq. The resultant gene sets from rGREAT were used as input for the R package fGSE89 to for cutting-edge analyses.

BspH1 restriction enzyme-based biosensor assay for A3B activity
DNA cytidine deaminase activity in T47D and MCF7 cells was measured using a BspH1-based fluorescence biosensor assay90. Briefly, cells were lysed in ice-cold RIPA buffer supplemented with protease and phosphatase inhibitors, and lysates were clarified by centrifugation (12000×g, 15min, 4°C). Protein concentration was determined by BCA assay.

For each reaction, 31.8µL of cell lysate containing 100µg total protein was mixed with 317.9µL reaction buffer (50mM potassium acetate, 20mM Tris-acetate, 10mM magnesium acetate, 100ng/mL BSA, pH 7.9) in a low-binding microtube. To initiate the assay, 0.3µL of 100µM FAM-labelled A3B_probe substrate (5’-FAM-TA-TAAGTTATCATGATATATA-TAMRA-3’) was added and incubated at 37°C for 1h to allow cytidine deamination. dsDNA hybridisation and restriction digestion were achieved by adding 0.9µL of 100µM TAMRA-labelled complementary strand (A3B_Probe_CS) and 1µL of BspH1 enzyme (1µ/µL, NEB), followed by a 4h incubation at 37°C. Fluorescence emission (500–700nm) was recorded (excitation 492nm) using an LS55 spectrophotometer (Perkin Elmer). Negative controls without lysate or enzyme were included for background subtraction.

Statistics
All data were analysed using the R statistical software (R Foundation for Statistical Computing, version 4.4.1). The specific statistical test used, sample size (n), and P-values are reported in the corresponding figure legends. Unless otherwise indicated, tests were two-sided, and a significance level of α=0.05 was used. When extremely small exact P-values were encountered, we capped the reported value at P<0.0001 and denoted this level of significance with ****.

Datasets used in this study
Previously published datasets used in this study include: GC-skew data for GRCh19 from R-loop DB [http://r-loop.org/]; T47D ER ChIP-seq data GSE148277; T47D GRO-seq data GSE128452; T47D RNAPII ChIP-seq data GSE105793; T47D Repli-Seq data ENCFF440QFG; T47D super enhancers data GSM2862201; T47D Mnase-seq data GSE74308.

Antibodies, siRNA oligonucleotides and constructs
The following antibodies used for this study are provided in Table 1. For siRNA, A3B knockdown was performed with the siRNA sequence reported by Periyasamy et al.5, CSB, XPF and XPG with siRNA reported by Sollier et al.47. For the UdgX uridine-DNA sensor, the coding sequence corresponding to WP_011726794.1 was subcloned into the multiple-cloning site of the pET-24a expression vector. For the mammalian RNaseH I overexpression construct, the protein sequence encoded by NM_002936 was used.

qPCR quantification
qPCR was performed on an Applied Biosystems ViiA7 thermocycler using POWER SYBR-Green master mix (Thermo Fisher). The primer pairs used in this study are provided in Table 2.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "1) The overall logic/rationale behind this manuscript is questionable. If A3B has a role in ER function, many people in SE Asia with a complete deletion of the A3B gene should be impacted. This deletion is well-known and no one has reported a defect in ER function associated with any phenotypes in these individuals.\n2) Methods – how did the authors successfully deliver active APOBEC3B to target cells using a lentivirus. A3B is a potent restrictor of retroviruses and it is highly likely that the target T47D cells now express an unintended mutant of A3B. A3B functionality should minimally be confirmed in clones by sequencing the integrated construct (part of existing WGS) and showing that induced levels in extracts have high DNA deaminase activity (in comparison to the A3B** mutant).\n3) Why is the mutant (A3B**) cell line also showing an increase in TC-to-TT mutations? This is bizarre as this A3B double mutant should be defective in DNA deamination. The authors should have deleted the endogenous A3 locus before setting up this complicated experiment.\n4) Interpretation – Given that A3B is well known to bind ssDNA (and ssRNA) and associate with R-loop regions, isn’t the simplest interpretation of the results that estrogen induces ER-responsive genes and that a subset of these genes can form R-loop structures that A3B can bind and deaminate?\n5) Alternative interpretation – RNA polymerase was recently shown to induce DNA breaks that can become DSBs (independent of A3B) (PMID: 39972147). Pol II enrichment is shown here at DSBs, etc. Therefore, maybe RNA Pol II has a role in the DNA breaks in this study.\n6) The experiments with XPG/F are potentially interesting; do both of these nucleases need to be depleted for DSB formation to be compromised? Individual knockdowns would be informative.\n7) The mutant A3B** is implied to exert a dominant negative effect. This doesn’t make sense with #3 above. Moreover, it may be dominant negative because it is overexpressed and pulling other cellular factors into unfolded protein aggregates (or generally triggering an unfolded protein response). Given that a similar E-to-Q mutation makes A3A unstable (PMID: 36923917), a better control may be an E255A mutant.\n8) Most importantly, if R-loop DSB formation is a general property of A3B, then key experiments should be done in at least one other estrogen/ER-responsive cell line.\nMinor:\nP4, L10 – references incorrect; none of the papers cited here show that A3B binds to ERalpha.\nFigures – all blots need at least 2 size markers (one above and one below each relevant band)\nMethods – reference(s) and Genbank accession number should be provided for all constructs (eg. UdgX)", "reviewer_2": "In this manuscript, the authors used unbiased models to map APOBEC3B (A3B) editing sites and show that A3B binds to and deaminates cytosines in the single-stranded DNA (ssDNA) of R-loops. They further show that estrogen-induced R-loops facilitate A3B recruitment and editing at R-loops, leading to the formation of DNA double-strand breaks and activation of estrogen-responsive genes. Although some connections have already been reported, such as the induction of R-loops by estrogen (PMID: 27552054), the recruitment of A3B to R-loops (PMID: 37735199), and the role of A3B in estrogen-induced gene transcription (PMID: 26411678), this study provides new insights into the role of A3B in R-loop-mediated transcriptional regulation, and is of significant and general interest. This study is further strengthened by the use of multiple genome wide approaches, such as WGS, DRIP-seq, SPI-seq and DSBcapture-seq, which provide very compelling results. I have only one comment concerning the proposed model, which may benefit from further discussion.\nMinor point:\nThe proposed model of DSB formation by two SSBs on opposite DNA strands is interesting and plausible. However, there are some points and/or alternatives in this model that could be further discussed. For example:\n- XPF and XPG can cleave the ssDNA of R-loops (PMID: 25435140; 31533039), implying that TC-NER activity could also target the same strand as A3B. Why, then, would the cleavage be limited to the RNA/DNA hybrid strand in response to estrogen?\n- TOP2B has been reported to induce DSBs at the promoters of estrogen-responsive genes (PMID: 16794079), as well as at many other stimulus-induced promoters, to initiate transcription. How does this observation align with the proposed model? Could TOP2B facilitate the recruitment of XPF to R-loops, as recently reported at TAD regions (PMID: 37939182)?", "reviewer_3": "In this manuscript, Zhang and colleagues use a combination of sequencing approaches to provide a comprehensive genome-wide view of APOBEC3B activity in engineered T47D breast cancer cells. The study is very thorough, the data are of a high quality and the manuscript is generally very well-written and presented. This work builds upon that of Periyasamy et al (reference 5), who first reported a requirement for APOBEC3B deaminase activity at ER target genes in 2015 and proposed a mechanism for transcriptional activation based on the recruitment of DNA repair factors to facilitate chromatin remodelling following APOBEC3B-mediated generation of DNA double strand breaks. It is instructive to compare the graphical abstract of that paper with Figure 8 of the current manuscript to show the additional insight provided by this study and therefore refinement of the model; namely the importance of R-loops and the specific involvement of transcription-coupled DNA repair. This manuscript therefore provides important confirmation of the findings of Periyasamy and colleagues, which is significant, as studies confirming this intriguing mechanism of transcriptional regulation and potentially key function of APOBEC3B have been distinctly lacking since its publication a decade ago. Overall, I find this to be a timely and important contribution to our limited knowledge of the physiological functions of APOBEC3 enzymes beyond their roles in viral restriction.\nPoints for the authors to address:\nLine 137-140 – include ‘(M+ colonies)’ somewhere in this description to clarify what M+ refers to (this is shown in Figure 1B but would be useful to also state in the text).\nLine 175 – ‘arise’ should be ‘arising’?\nLine 279 – ‘shown’ should be ‘showed’\nDoes the immunoblot shown in Figure S10 relate to the cells used for the experiments shown in Figure 5 in addition to those shown in Figure S11? If not, the authors should provide demonstration of A3B knockdown for the experiments shown in Figure 5.\nIn lines 379 – 381 the authors state “To assess differential gene expression, we employed the interaction modelling approach using the DESeq2 program to examine the impact of loss of endogenous A3B activity on the E2 response”. While their speculation that “the enzymatic inactive A3B** can potentially exert a dominant negative effect by sequestering regulatory partners of endogenous A3B” (lines 375-377) is reasonable, this is not clearly demonstrated. If the demonstration of this lies (as I think the authors imply), “in Figure 1, S1, S3, S5” (line 373), the evidence should be pointed out more clearly to the reader – which specific results in these figures demonstrate this? It is important to address whether A3B** is indeed acting in a dominant negative fashion, as earlier in the manuscript, the M+ colonies (which co-express A3B** and hUGI) are described as demonstrating the “intrinsic A3B effect”. In this scenario, the authors imply that A3B** is not acting as a dominant negative, or at least that it is not completely suppressing the activity of endogenous A3B. The interpretation of the experiments shown in Figure 7 also depend on the assumption that A3B** suppresses the activity of endogenous A3B.\nIn line 401 “…we found that both depletion of A3B and blocking BER repair…” A3B has not been depleted (e.g. using siRNA) here (Figure 7), as was done in other experiments in the manuscript, rather the authors are assuming that its activity has been suppressed by A3B**. Provided that this assumption is justified (see point above) the authors should nonetheless modify their terminology here to avoid confusing the reader between experiments in which they have actually depleted A3B using siRNA versus inhibiting its function with the presumed dominant negative construct.\nFigure 1B – ‘seperate’ should be ‘separate’\nFigure 4A – ‘trasnfection’ should be ‘transfection’"}, "author_response": "1. The overall logic/rationale behind this manuscript is questionable. If A3B has a role in ER function, many people in SE Asia with a complete deletion of the A3B gene should be impacted. This deletion is well-known and no one has reported a defect in ER function associated with any phenotypes in these individuals.\nWe acknowledge the Reviewer’s concern regarding the high prevalence of the APOBEC3B germ-line deletion in South-East Asia, which appears to conflict with our conclusion that APOBEC3B (A3B) promotes oestrogen-receptor (ER) activity. We have reviewed the data on both the population genetics of the deletion and the mechanistic data on A3B in ER-positive breast cancer, and we are confident that the two sets of observations can be reconciled.\n1. Although the 29-kb deletion that removes the entire A3B coding region is indeed common in East Asian populations, most clinical and epidemiological research to date has focused almost exclusively on cancer risk. To our knowledge, no study has systematically measured classical ER-physiology end-points—pubertal timing, fertility, bone density, vasomotor symptoms or serum oestradiol—in individuals who are homozygous for the deletion. The frequent statement that “no defect has been reported” therefore reflects a lack of data, not positive evidence that ER function remains unimpaired in these subjects.\n2. Most tumours that arise in deletion carriers are heterozygous, not homozygous, for the structural variant. Even a single intact A3B allele appears to provide sufficient enzyme to engage the enhancer-remodelling loop we describe. Epidemiological surveys that pool heterozygotes and homozygotes therefore dilute any impact of complete A3B absence, making it unsurprising that they find no difference in ER status.\n3. Most significantly, the deletion is not a straightforward loss-of-function allele: it generates a chimeric APOBEC3A-B mRNA transcript where the highly active APOBEC3A open reading frame is fused to the stabilising 3′-UTR of A3B (Caval et al., PMID: 25298230). The hybrid mRNA is more abundant than native APOBEC3A mRNA, and the resulting protein has at least ten times the cytidine-deaminase activity of A3B. In normal tissues this additional APOBEC3A activity is likely to compensate for the absence of A3B, preserving basal ER programmes and masking any physiological deficit that might otherwise be detected.\n4. Our study focuses on the tumour context, rather than normal endocrinology. Super-enhancers, heightened transcriptional activity and replication stress characterise ER-positive breast cancers. Under these conditions, our data demonstrate that A3B is part of a positive-feedback loop: ERα induces A3B, A3B edits cytosines in R-loops at ER enhancers, and the resulting uracil triggers TC-NER–mediated strand breaks that accelerate chromatin remodelling. This circuitry provides a selective advantage to rapidly proliferating cancer cells and may represent an oncogenic contribution of A3B that is simply irrelevant to normal, slowly cycling mammary epithelium. When the gene is constitutionally absent, tumours appear to reach the same hyper-ER state through alternative routes such as greater reliance on other pioneer factors such as FOXA1 or GATA3 (Carroll et al., 2005, Cell; Kouros-Mehr et al., 2006, Cell), amplification of co-activator complexes (Anzick et al., 1997, Science), or utilisation of the hyper-active APOBEC3A–B chimaera described above. In other words, tumour evolution can bypass a requirement for A3B, but doing so requires additional changes that are unnecessary when A3B is present.\n5. Finally, the deletion does not protect against breast cancer; instead, it modestly increases overall risk and leaves the relative proportion of ER-positive tumours unchanged. These observations strongly argue that the absence of A3B does not impair ER signalling in vivo, but, importantly, they do not contradict our central finding that cancer cells expressing active A3B exploit it to enhance ER-driven transcription.\nWe have added a new paragraph to the Discussion (page x, lines y) explaining these points, and we now highlight the need for focused endocrine phenotyping of deletion homozygotes as an important future direction:\n(Line 3, Page 21):\n…“Within this mechanistic context, the high prevalence of the 29 kb germline deletion eliminating the APOBEC3B open reading frame (ΔA3B, rs12628403) in South East Asian populations appears inconsistent with an obligatory physiological role for A3B in ER signalling, as homozygous carriers show no endocrine abnormalities [56,57]. Existing studies have focused primarily on cancer risk, leaving classical ER related traits, such as pubertal timing, fertility and bone density, unexamined. Importantly, ΔA3B is not a true null: its breakpoint generates a stabilised APOBEC3A-B chimaera with elevated cytidine deaminase activity [58], probably buffering ER pathways in normal tissues. In contrast, our data indicate that ER positive tumours exhibit an oncogenic contribution from A3B, forming an amplifier circuit in which ERα induces A3B [5,59], and A3B edits R loops at ER enhancers to accelerate transcriptional cycling. Consistent with this compensatory model, ΔA3B does not reduce breast cancer incidence and may modestly increase risk, including in ER positive disease [56,57] . We therefore regard A3B as conditionally essential, dispensable under homeostatic endocrine signalling but strongly selected in the replication stressed, ER driven environment of cancer. Future work should combine rs12628403 genotyping with systematic endocrine phenotyping, clarify the respective contributions of A3A and A3B at ER enhancers, and test the therapeutic potential of targeting A3B or uracil processing steps in genotype stratified models.”…\n2) Methods – how did the authors successfully deliver active APOBEC3B to target cells using a lentivirus. A3B is a potent restrictor of retroviruses and it is highly likely that the target T47D cells now express an unintended mutant of A3B. A3B functionality should minimally be confirmed in clones by sequencing the integrated construct (part of existing WGS) and showing that induced levels in extracts have high DNA deaminase activity (in comparison to the A3B** mutant).\nWe acknowledge the Reviewer’s concern that lentiviral delivery of active A3B might provoke self-restriction or mutagenic inactivation of the viral genome, leading to the generation of unintended A3B variants in target cells. Indeed, our early attempts using constitutive CMV-driven vectors produced very low titres and heterogeneous transductants, consistent with leaky A3B expression compromising viral replication. To address this, we employed the tightly controlled pTRIPZ doxycycline-inducible backbone, which contains six tandem tet-operator sites upstream of a minimal CMV promoter and requires the rtTA3 trans-activator for transcriptional activation. Because producer HEK293T cells lack rtTA3 and virus production occurs in the absence of doxycycline, the A3B open reading frame remains completely silent during packaging, thereby preventing any A3B-mediated restriction or mutagenesis of the lentiviral genome.\nWhole-genome sequencing of all independent T47D clones confirmed intact proviral A3B integrations without any deep or non-synonymous substitutions or frameshifts within the coding region (NM_004900.5; new Supplementary Table 1).\nFunctional assays also verified enzymatic integrity. A fluorometric BspHI cytidine-deaminase biosensor demonstrated robust activity in lysates from doxycycline-induced A3B-expressing cells, whereas extracts from A3B** mutants were inactive (new Figures S1–S2). Additionally, slot-blot analysis using the biotinylated M. smegmatis uridine-sensor UdgX revealed high levels of A3B-mediated uracil incorporation in A3B-induced colonies, with only basal endogenous A3B signals in A3B**-induced clones (new Figure S3).\nTogether, these data confirm that the integrated lentiviral A3B allele is both sequence-intact and catalytically active. To include the additional data, we have edited the first and second paragraphs in the “Detection of A3B DNA deamination sites using a BER-deficient cancer cell model” results section, as follows :\n(Line 18, Page 5):\n…”To identify A3B deamination sites with potential functional impacts on protein coding or gene regulation, we conducted whole-genome sequencing (WGS) of ER-positive T47D human breast cancer cells expressing doxycycline-inducible A3B. As a control, we adopted an enzymatically inactive A3B E68Q/E255Q double mutant (A3B**), which was confirmed to lack detectable enzymatic activity in multiple cell line models (Figure 1A, S1). Bicistronic expression of the humanised bacteriophage PBS2 uracil glycosylase inhibitor (hUGI) peptide was employed to inhibit BER-associated uracil-DNA glycosylase (UDG/UNG) 20,30. We showed that lentiviral hUGI expression prevents the removal of A3B induced uracil bases, allows more effective capture of A3B editing events, and it is sufficient to block the majority of UDG/UNG activity within cells (Figure S2). The selection of the T47D cell line was based on its TP53 loss-of-function mutation, which is known to confer protection against synthetic lethality induced by ectopic A3B expression 35. Doxycycline-induced expression of A3B/hUGI and effective uridine incorporation were confirmed by immunoblotting and slot blotting using Mycobacterium smegmatis UdgX as a uridine DNA sensor (Figure 1A, S3A-B). The cells were then exposed to doxycycline for five days, and genomic DNA samples were sequenced from five individual T47D clones expressing A3B-hUGI (A+ colonies, Figure 1B). For control purposes, sequencing results from three uninduced colonies (A- colonies) were included to represent single-nucleotide variants (SNVs) arising from isolating individual clones. In addition, three T47D colonies co expressing hUGI and the enzymatically inactive A3B** (M+ colonies) were also included. A3B** neither interfered with the activity of endogenous A3B (Figure S1, S2) nor triggered an unfolded protein response (Figure S4), indicating that it has no functional impact on the endogenous A3B enzyme. The M+ colonies were therefore expected to capture base editing events mediated by endogenous APOBEC/AID family enzymes in T47D cells, predominantly A3B 5. Slot blots with UdgX uridine DNA sensor confirmed that both the endogenous and induced wild-type A3B in the individual colonies retained full enzymatic activity (Figure S3C-D)”…\nAdditionally, for the third paragraph of the same section, we have added the following sentence:\n(Line 20, Page 6):\n…”Analysis of the WGS data confirmed that the integrated lentiviral construct, including the A3B open-reading frame and flanking vector sequence, was completely sequence-intact, with no deep and non-synonymous mutations detected following A3B induction (Supplemental Table 1).”…\n3) Why is the mutant (A3B**) cell line also showing an increase in TC-to-TT mutations? This is bizarre as this A3B double mutant should be defective in DNA deamination. The authors should have deleted the endogenous A3 locus before setting up this complicated experiment.\nWe appreciate the Reviewer’s concern that expression of the catalytically inactive A3B** construct still results in an increased burden of TC>TT substitutions. The explanation lies in both the design of the A3B** fusion and the persistent activity of endogenous APOBEC enzymes within T47D cells. The A3B** protein carries double active-site substitutions (E255Q/E258Q) that abolish its own cytidine deaminase activity but is bicistronically expressed with hUGI, a potent inhibitor of uracil DNA glycosylases (UNG/UDG). Expression of UNG inhibits base-excision repair (BER), preventing the removal of uracils generated by the native endogenously expressed APOBECs, mainly A3B in T47D cells. These uracils persist into S phase and are replicated as thymines, leading to the characteristic TC>TT transitions observed under BER-deficient conditions.\nTo verify that these substitutions originate from endogenous A3B rather than the mutant A3B** construct itself, we analysed SKBR3 cells, which lack detectable APOBEC family activity (Periyasamy et al., 2015; PMID 26411678, ref 5 in main text). In this model, induction of A3B-GFP robustly induced deamination activity. In contrast, expression of the mutant A3B**-GFP showed no evidence for increased deamination. Consistent with the absence of endogenous A3B in the SKBR3 background, neither construct generated C>T mutations when BER was inhibited by adding excess exogenous UGI peptide (new Figure S1C). Conversely, T47D lysates, which contain active endogenous A3B, exhibited a restored deaminase signal upon addition of the UGI peptide, consistent with the unmasking of native A3B activity following BER inhibition.\nFurthermore, we verified that co-expression of hUGI effectively blocks BER and unmasks endogenous A3B-mediated deaminase activity (new Figure S2). Under these BER-deficient conditions, A3B**–hUGI expression resulted in the accumulation of uridine DNA incorporation, which was not further increased by additional UGI peptide. This result indicates near-complete blockade of BER in cell lysates by the bicistronically expressed hUGI peptide.\nTogether, these experiments demonstrate that the modest TC>TT enrichment observed in A3B** cells results from residual endogenous A3B activity, revealed by hUGI-mediated suppression of BER, rather than from unintended activity of the mutant enzyme itself.\nTo clarify this, we have added the following sentence to the main text :\n(Line 18, Page 7):\n…“In addition, analysis of the M+ colonies showed a measurable but weaker enrichment of C>T substitutions at the A3B-preferred 3′-TCW trinucleotide motif and partial retention of SBS2 and Signature B. This effect is attenuated relative to wild-type A3B induction and reflects deamination by the endogenous A3B enzyme, whose activity is unmasked when the bicistronically expressed hUGI peptide suppresses BER.”…\n4) Interpretation – Given that A3B is well known to bind ssDNA (and ssRNA) and associate with R-loop regions, isn’t the simplest interpretation of the results that estrogen induces ER-responsive genes and that a subset of these genes can form R-loop structures that A3B can bind and deaminate?\nWe concur with the Reviewer’s interpretation, and indeed, our description of A3B as part of an ER-driven positive-feedback loop that facilitates rapid enhancer remodelling in ER-positive tumour cells remains consistent throughout the manuscript. In the Summary we state that\n“（Line 30, Page 1）…A3B preferentially targets transcriptionally active regulatory regions in an R-loop-dependent manner… A3B binds to and deaminates single-stranded DNA within R-loops, facilitated by ER transactivation.”\nThe Research Highlights echoes the same point:\n(Line 14, Page 2)“R-loop formation by ER activation facilitates APOBEC3B binding” and\n(Line 17, Page 2) “APOBEC3B regulates ER activity by promoting DSB formation at E2-induced R-loops”.\nIn the Results section titled “ER-induced R-loop formation facilitates A3B binding” (Page 10), we document that ER-induced R-loops serve as the substrate for A3B binding and cytidine deamination, as shown by ssDRIP-seq and SPI-seq mapping. In this section, we explicitly note that\n(Line 12, Page 12)“…the availability of R-loops may dictate E2-induced A3B binding and editing chromatin in or adjacent to the R-loop….”\nThus, the mechanistic model the Reviewer proposes—E2 stimulates ER-responsive transcription; a subset of these loci form R-loops; A3B is recruited to the displaced ssDNA and deaminates it—is already the central narrative we convey in both the abstract and the main body of the manuscript.\n5) Alternative interpretation – RNA polymerase was recently shown to induce DNA breaks that can become DSBs (independent of A3B) (PMID: 39972147). Pol II enrichment is shown here at DSBs, etc. Therefore, maybe RNA Pol II has a role in the DNA breaks in this study.\nWe thank the Reviewer for raising this possibility and for drawing attention to the recent work by Liu and colleagues (2025; PMID 39972147), which demonstrates that paused RNA polymerase II (Pol II) can generate DNA termini that mature into double-strand breaks at specialised transcription-termination regions containing a CCTTTTTT-like pause motif. These regions are typically a few hundred bases downstream of poly-adenylation signals.\nIn contrast, the double-strand breaks documented in our study occur almost exclusively at promoters and oestrogen-responsive enhancers, lying within ± 1 kb of transcription-start sites or within Erα-bound enhancer chromatin. In contrast, almost none of the mapped sites were found within 1 kb downstream of annotated poly(A) sites. Mechanistically, the DSB we see require the complete A3B→UNG→XPF/XPG pathway; disruption of any component, catalytic A3B, uracil-DNA glycosylase, or the TC-NER nucleases, abolishes the DSB signal. The Pol II-associated breaks described by Liu and colleagues arise independently of cytosine deamination, do not involve uracil-excision repair, and are not processed by XPG/XPF. Therefore, while Pol II is necessarily present at our damage sites because these are transcriptionally active regions, the genomic localisation and strict dependence on A3B-mediated C>U editing indicate that the lesions we observe are generated via the A3B-initiated pathway, rather than by the termination-associated mechanism described by Liu and colleagues.\nWe have included a clarifying sentence in the Discussion acknowledging this complementary Pol II-dependent mode of break formation.\n6) The experiments with XPG/F are potentially interesting; do both of these nucleases need to be depleted for DSB formation to be compromised? Individual knockdowns would be informative.\nWe thank the reviewer for highlighting the importance of assessing the relative contribution of the TC-NER endonucleases XPF and XPG to A3B-dependent DSB formation. To address this point, we performed individual and combined siRNA knockdowns of XPF and XPG in T47D cells, followed by quantitative analysis of DSBs using the DSBCapture qPCR assay. The results show that single depletion of either XPF or XPG produced only a modest decrease in E2-induced, A3B-dependent DSB signals compared with control cells. In contrast, simultaneous knockdown of both nucleases led to a near-complete loss of DSB formation at all representative loci (new Figure S19). Western blotting confirmed efficient depletion of the individual proteins.\nThese data demonstrate that both nucleases contribute to the cleavage events required for A3B-dependent DSB generation. We also find redundancy, so that depletion of one can be partially compensated for by the other. This redundancy is consistent with the known structural rather than strict strand specificity of XPF and XPG described across different systems.\nThese results provide clear experimental evidence that both nucleases participate in the A3B-driven damage pathway and that loss of either enzyme alone is insufficient to block DSB formation.\nWe have updated the Results section to incorporate these findings as follows:\n(Line 5, Page 15)\n“Using quantitative PCR analysis of DSBCapture-enriched DNA samples, we discovered that depletion of these TC-NER components was sufficient to reduce the formation of E2-responsive A3B-dependent DSBs in T47D and MCF7 cells (Figure 5I, S18). Single depletion of either XPG or XPF caused only a slight reduction in E2 induced A3B dependent DSB formation, whereas combined depletion almost completely abolished DSB production at all examined loci (Figure S19). Because XPF and XPG can incise either strand of the R loop without strict strand preference, the modest effect of single depletion suggests functional redundancy between these endonucleases, each being individually capable of mediating the incision events required for R loop processing and subsequent DSB formation.”\n7) The mutant A3B** is implied to exert a dominant negative effect. This doesn’t make sense with #3 above. Moreover, it may be dominant negative because it is overexpressed and pulling other cellular factors into unfolded protein aggregates (or generally triggering an unfolded protein response). Given that a similar E-to-Q mutation makes A3A unstable (PMID: 36923917), a better control may be an E255A mutant.\nOur response:\nWe thank the reviewer for raising this important point regarding the function and cellular behaviour of the catalytically inactive A3B** (E68Q/E255Q) mutant. We fully agree that a catalytically inert but structurally stable construct is essential for accurate interpretation of our experiments. Accordingly, we have performed detailed biochemical and cellular characterisation of the A3B** mutant as described in the revised manuscript. The results were as follows:\n1. Consistent with the predicted loss of catalytic activity, detectable expression of the A3B** mutant protein displayed no detectable cytidine-deaminase activity when expressed in the APOBEC-null SKBR3 model. In contrast, expression of equivalent levels of the wild-type A3B protein resulted in significant deaminase activity in the same assay (new Figures S1C, S2B), indicating that the mutant protein is enzymatically inactive rather than unstable.\n2. To assess whether A3B** interferes with or suppresses endogenous A3B, we quantified total cytidine-deaminase activity in T47D cells with or without doxycycline-induced A3B** expression. Activity levels were unchanged, and the abundance of endogenous A3B protein remained constant (new Figures S1A–B). These data demonstrate that A3B** neither competes with nor inhibits the endogenous enzyme and therefore cannot act as a dominant-negative variant.\nThese results differs from our earlier expections.\n3. We directly evaluated the reviewer’s concern regarding possible proteotoxic stress. RNA-seq analysis of A3B**-GFP or A3B**-hUGI expressing cells (new Figure S3) revealed no induction of genes associated with unfolded protein, heat-shock, or general stress responses, showing that the expression of the mutant protein does not trigger an unfolded protein response.\nCollectively, these experiments demonstrate that A3B** is catalytically dead, biochemically stable, and biologically neutral with respect to endogenous A3B activity and overall cell homeostasis. Although related E>Q mutations in A3A have been linked with protein instability, the A3B paralogue displays distinct structural tolerance at these positions, as evidenced by our experiments.\nFor clarity, the revised manuscript explicitly describes the above validation data and refers to the newly included Figures S1–S3, which show no evidence of a dominant negative effect by A3B**.\nThe relevant text has been added to the Results section:\n(Line 5, Page 5):\n“As a control, we adopted an enzymatically inactive A3B E68Q/E255Q double mutant (A3B**), which was confirmed to have no detectable enzymatic activity in multiple cell line models (Figure 1A, S1)… …In addition, three T47D colonies co-expressing hUGI and the enzymatically inactive A3B** were also included (M+ colonies). A3B** neither interfered with the activity of endogenous A3B (Figure S1, S2) nor triggered an unfolded protein response (Figure S4), indicating that expression of the inactive A3B** mutant has no functional impact on the endogenous enzyme… ”\n(“Blocking the processing of A3B editing sites into DSBs impairs E2 response”)\n“…To examine the functional impact of blocking this processing step, we performed RNA-seq analyses using T47D cells containing the doxycycline-inducible, catalytically inactive A3B** mutant expressed together with hUGI (Figure 6A). As shown previously (Figure 1, S1–S4), A3B** has no detectable enzymatic activity and does not interfere with endogenous A3B, thereby serving as a non-catalytic control that allows assessment of uracil repair-dependent processes. By co-expressing hUGI, we effectively blocked the conversion of A3B-induced uracil lesions into repair intermediates that lead to DSBs (Figure S2)...”\n8) Most importantly, if R-loop DSB formation is a general property of A3B, then key experiments should be done in at least one other estrogen/ER-responsive cell line.\nFor an independent validation we selected MCF7, a prototypical luminal-A, ER-positive breast-cancer line with high basal A3B activity (Periyasami et al., 2015, ref 5 in main text; Burns et al., 2013, ref 20 in main text). We repeated the critical experiments, inducible A3B expression or knock-out, ssDRIP-qPCR, SPI-qPCR, DSBCapture-qPCR, and XPG/XPF depletion. In these cells, we obtained results consistent with those from the T47D model.\nThe new data and a side-by-side quantitative summary are presented in Figures S16, S18 and S20 of the revised manuscript.\nMinor:\nP4, L10 – references incorrect; none of the papers cited here show that A3B binds to ERalpha.\nWe thank the reviewer for noting this inaccuracy. The original reference list had inadvertently lost the citation to Periyasamy et al., 2015 (ref 5 in main text) during manuscript editing and reformatting. That study indeed demonstrated the interaction between A3B and ERα. We have now reinstated this citation in the revised version of the manuscript, and corrected the reference numbering accordingly. We apologise for this oversight.\nFigures – all blots need at least 2 size markers (one above and one below each relevant band)\nWe thank the reviewer for this helpful suggestion. In the revised figures, we have re-run the key immunoblot experiments to include clearly visible molecular-weight markers above and below each relevant band. In the revised figures, these markers are now shown for all immunoblots.\nMethods – reference(s) and Genbank accession number should be provided for all constructs (eg. UdgX)\nWe thank the reviewer for this helpful suggestion. In the revised Methods section, we have now included the GenBank accession numbers and corresponding references for all constructs used. Specifically, we have added UdgX (WP_011726794.1) and RNase I (NM_002936) to ensure complete transparency and reproducibility."}, {"round": 2, "reviewer_comments": {"reviewer_1": "The authors have done a good job of addressing reviewer comments and revising the manuscript. The elaboration of the NER endonucleases and the additional results with another ER+ cell line (MCF7) are particularly appreciated.\nRemaining (minor) comments:\n1) The title is a little strong; “influences” is a more appropriate word than “determines”\n2) The authors should double check kDa sizes in Fig 4A as endogenous A3B typically runs less than 40 kDa (e.g., ref.27, PMID 31544853, and many others)\n3) A prior study demonstrating R-loop mutagenesis by A3B is not adequately cited (ref.27). In particular, claims here of “first” such as p19, line4 should be eliminated.", "reviewer_2": "The authors have adequately addressed all my comments in their revised manuscript.", "reviewer_3": "The authors have addressed all my comments and I have no further concerns."}, "author_response": "RESPONSE TO REVIEWERS' COMMENTS\nReviewer #1 (Remarks to the Author):\nThe authors have done a good job of addressing reviewer comments and revising the manuscript. The elaboration of the NER endonucleases and the additional results with another ER+ cell line (MCF7) are particularly appreciated.\nOur response: We thank the reviewer for the positive evaluation of our revised manuscript and for recognising our efforts to address role of NER endonucleases and inclusion of a second cell line. The Reviewer’s constructive and critical comments led to revisions that have, in our view, strengthened both the presentation and the interpretation of our findings.\nRemaining (minor) comments:\n1) The title is a little strong; “influences” is a more appropriate word than “determines”\nOur response: We thank Reviewer 1 for this helpful suggestion regarding the strength of the title. In response, we have revised the title to use a more moderate word. Specifically, we have replaced “determines” with “modulates,” so that the title now reads:\n“R-loop editing by DNA cytosine deaminase APOBEC3B modulates the activity of oestrogen receptor enhancers”\n2) The authors should double check kDa sizes in Fig 4A as endogenous A3B typically runs less than 40 kDa (e.g., ref.27, PMID 31544853, and many others)\nOur response: We thank Reviewer 1 for pointing out this. We have re-examined the SDS–PAGE running buffer conditions and the manufacturer’s instructions for the protein ladder. We confirmed that the band we had previously labelled as 40 kDa should in fact be labelled as 37 kDa. We have now corrected this label in Fig. 4A (and anywhere else applicable in the manuscript).\n3) A prior study demonstrating R-loop mutagenesis by A3B is not adequately cited (ref.27). In particular, claims here of “first” such as p19, line4 should be eliminated.\nOur response: We thank Reviewer 1 for this important point regarding the prior study (ref. 27) on R-loop mutagenesis by A3B. In response, we have removed all priority-type wording (e.g., “first,” “new,” “novel”) from the manuscript, including the statement on page 19, line 4, and elsewhere as applicable.\nAlso, We have now strengthened the citation to McCann et al. (2023) (ref. 27) in the revised text. The relevant sentence has been updated to:\n“…This hypothesis gains support from studies utilising the APOBEC–Cas9 engineered system to edit cytosines in R-loop DNA 26 and a recent investigation revealing the influence of A3B on R-loop homeostasis and consequent R-loop–associated mutagenesis27….”\n“…While a previous study has established that A3B interacts with R-loops and contributes to transcription-associated mutagenesis, the precise mechanism through which A3B engages with R-loops and its functional consequences remain unclear 27...”\nAnd\n“…Recent findings have demonstrated that cells with high-level expression of A3B exhibit an accelerated clearance of steady-state or stimulation-induced R-loops 27…”\nTogether, these revisions more accurately acknowledge and integrate the contributions of McCann et al. (2023) to the understanding of A3B-mediated R-loop biology and mutagenesis.\nReviewer #2 (Remarks to the Author):\nThe authors have adequately addressed all my comments in their revised manuscript.\nOur response: We are grateful to Reviewer 2 for carefully evaluating our work and for the constructive feedback provided during the review process. The comments have been very helpful in refining and improving the manuscript.\nReviewer #3 (Remarks to the Author):\nThe authors have addressed all my comments and I have no further concerns.\nOur response: We sincerely thank Reviewer 3 for their positive feedback. We greatly appreciate the time and thoughtful comments that contributed to strengthening our manuscript."}]

Decision letter