Dual roles of DEAD-box RNA helicase Brr2 in genome stability regulation

Chen, Xiaolan; You, Jin; Ma, Qin; Lin, Jiamei; Ji, Yu; Li, Shi; Song, Zhenxing; Su, Rui; Shan, Ge; Huang, Chuan

doi:10.1038/s41467-025-64174-8

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Published: 15 October 2025

Dual roles of DEAD-box RNA helicase Brr2 in genome stability regulation

Xiaolan Chen¹^na1^nAff3^nAff4,
Jin You¹^na1,
Qin Ma¹,
Jiamei Lin¹,
Yu Ji¹,
Shi Li¹,
Zhenxing Song¹,
Rui Su¹,
Ge Shan ORCID: orcid.org/0000-0002-3561-2088² &
…
Chuan Huang ORCID: orcid.org/0000-0003-2565-9154¹

Nature Communications volume 16, Article number: 9133 (2025) Cite this article

Subjects

Abstract

R-loop is a common chromatin feature consisting of a displaced single-stranded DNA and an RNA-DNA hybrid, and dysregulation of R-loop surveillance results in genomic and transcriptomic instability. Although the RNA moiety of most R-loops originates from linear transcripts, circular RNAs (circRNAs), outputs from back-splicing, can also hybridize with the complementary strand of a DNA duplex. However, how circRNA-associated R-loops (ciR-loops) are monitored remains elusive. Here, we identify the DEAD-box RNA helicase Brr2 as an evolutionarily-conserved ciR-loop repressor with dual roles in inhibiting circRNA generation and resolving harmful ciR-loops. Accumulation of ciR-loops caused by loss-of-function of this dual-action factor induces antisense transcription and premature transcription termination for many genes and generates significant DNA damage, which further leads to a series of defects in DNA replication, cell division and cell proliferation. We propose that functional integration of multilayered regulation by a single protein can be an efficient double protection against genome instability.

Introduction

R-loop is a crucial three-stranded nucleic acid structure comprising a displaced single-stranded DNA (ssDNA) and an RNA-DNA hybrid, and is usually thought to be formed and erased during the process of transcription, DNA replication and/or DNA repair under normal conditions in a scheduled manner^1,2,3,4. Accidental perturbations in R-loop formation, resolution and distribution under pathological or stressed conditions are tightly associated with human diseases, such as cancer and neurological disorders^5,6,7. As a functional chromatin feature, R-loops prevalently exist along the genome, particularly at active genes^8,9,10,11, and have both positively and negatively regulatory effects on gene expression in different contexts through distinct mechanisms^1,2,3. For example, in the promoter regions, R-loops are rapidly generated to prevent DNA methyltransferases from methylating DNA at CpG islands, ultimately impeding transcription silencing^12,13; R-loops around the transcription end sites (TESs) can attenuate the read-through activity of elongating RNA polymerase II (Pol II) by inducing the dissociation of Pol II from these regions, thereby ensuring an efficient termination step and avoiding the aberrant enrichment of read-through transcripts¹⁴. By contrast, the excessive accumulation of nonscheduled pathological R-loops can interfere with genome and transcriptome, which further leads to an array of harmful consequences, such as premature transcription termination (PTT), DNA replication impairment and even DNA breaks^1,2,3. Therefore, the precise control of R-loop homeostasis is an essential aspect to sustain genomic and transcriptomic stability in living organisms.

Indeed, cells have developed several pathways through different factors to either restrict nonscheduled R-loop formation or resolve accumulated R-loops. The conserved endonuclease RNase H family, including RNase H1 and RNase H2, harbors an N-terminal hybrid binding domain (HBD) and C-terminal endonuclease motif and can specifically interact with and resolve R-loops by degrading the RNA moiety of RNA-DNA hybrids^15,16,17. The DNA repair factor RAD52 can recognize R-loops at the sites of DNA damage and in turn recruits the structure-specific endonuclease XPG for R-loop cleavage^18,19. Notably, some RNA processing factors are also involved in controlling R-loops. For example, the transcription elongation factor TFIIS ensures a high rate of RNA synthesis by reducing Pol II pausing and backtracking, which further diminishes the chance of R-loop formation²⁰; the splicing factor SRSF1 directly binds to the new-synthesized RNAs and promotes an efficient nuclear export process, thereby preventing them from re-annealing to their paired DNA templates to generate R-loops²¹. The aforementioned diversity indicates that the regulation of R-loop is likely more complicated than currently appreciated, and that additional layers of mechanisms might be underestimated.

Circular RNAs (circRNAs) are a type of natural exonuclease-resistant transcripts with a unique covalently closed structure. Most circRNAs (i.e., exonic circRNAs, also termed EcircRNAs) are generated from one or more exons of numerous eukaryotic protein-coding genes via a non-canonical splicing event called back-splicing (BS), during which a splice donor is connected to an upstream acceptor^22,23,24. In contrast to EcircRNAs, a small subgroup of circRNAs are outputs of certain intronic sequences^25,26,27, or even are produced from genomic regions originating from distinct genes^28,29,30. Although the first circRNA was identified in 1976³¹, only in the recent 10 years has it been appreciated that many circRNAs are vital modulators functioning in a wide range of molecular processes through diverse mechanisms, such as orchestrating transcription and splicing, controlling the activity of RNA-binding proteins (RBPs) and even serving as active translation templates to encode functional proteins^22,23,24,32. Some circRNAs are evolutionarily-conserved and have been found to exert similar physiological functions across different species. For example, conserved circRNAs derived from the reproductive gene Boule can mediate male fertility maintenance through the circRNA-heat shock protein regulatory axis in both mice and Drosophila³³. Dysregulation of circRNAs thus leads to a spectrum of severe consequences at the physiological and pathological levels^34,35,36,37. More interestingly, emerging studies have demonstrated that circRNAs are also capable of forming R-loops with genomic DNA^38,39,40. Given that circRNAs are generally much more stable than linear RNAs^41,42,43, nonscheduled circRNA-associated R-loops (ciR-loops) seem relatively difficult to be eliminated and may give rise to a drastic disturbance in genome and transcriptome, such as DNA breaks/damage and aberrant transcription events. Therefore, we speculate that cells require sophisticated mechanisms to protect themselves against ciR-loop accumulation, which remains a challenging question so far.

DEAD-box RNA helicases represent a large family of RBPs and ATP-dependent enzymes in eukaryotes, and are characterized by the specific DEAD motif (Asp-Glu-Ala-Asp). These family members share a highly conserved helicase core with several functional domains that mediate their binding abilities to RNA or ATP and enzyme activities to separate duplex oligonucleotides^44,45. The conventional viewpoint on DEAD-box proteins is that they are post-transcriptional regulators having central roles in almost every aspect of the life span of diverse RNA species^46,47. For example, the proper separation of RNA-processing bodies from stress granules in the cytoplasm is precisely monitored by DDX6, loss of which will lead to the dysregulation of translation and degradation of inefficiently translated mRNAs^48,49. However, it is increasingly clear that some of DEAD-box RNA helicases are chromatin-interacting proteins and can non-canonically function in transcription^50,51,52,53. For example, our previous study has demonstrated that DDX56 acts as a stress check-point, which can specifically maintain the basal, but not stress-induced, transcription of stress-responsive genes by forming an open chromatin structure and facilitating Pol II assembly to the transcriptional start sites (TSSs)⁵⁴; DDX41 ensures a continuous transcription process by removing physical barriers formed by the harmful R-loops in the promoter proximal regions⁵⁵. These cases indicate the unanticipated potential of DEAD-box RNA helicases in functional integration of two independent regulatory processes—transcriptional and post-transcriptional regulation. Based on these hints, we assume that certain RNA helicases may serve as a multifaceted regulator to coordinate and fine-tune ciR-loop homeostasis at two separated phases, during which circRNA metabolism and R-loop elimination were independently controlled by a single RNA helicase in a stepwise manner. But this hypothesis needs further investigation.

In this study, we identified the DEAD-box RNA helicase Brr2 as a potent circRNA biogenesis inhibitor by a systematic screening on all known Drosophila RNA helicases. The following high-throughput sequencing analysis revealed the pervasive regulatory effects of Brr2 on the balance between BS and linear splicing (LS). Loss-of-function of Brr2 led to a significant accumulation of harmful ciR-loops in the genome, which subsequently impaired genomic and transcriptomic integrity. Depletion of the human homolog of Brr2 (SNRNP200) resulted in similar effects, indicating the evolutionarily-conserved role of Brr2 in regulating circRNAs and ciR-loops. More interestingly, to efficiently prevent ciR-loop accumulation, Brr2 can act as a direct R-loop-unwinding factor in addition to its inhibitory role in circRNA generation, suggesting the unique ability of Brr2 in fine-tuning R-loop homeostasis at multilayered levels. Furthermore, we unveiled that the Brr2-involved regulatory axis is required for the proper control of DNA replication, cell division and cell proliferation. Altogether, our work reveals the dual roles of Brr2 in controlling genome stability, and provides a novel insight into how a single protein integrates transcriptional and post-transcriptional events.

Results

A systematic screening on DEAD-box RNA helicases reveals the potent and specific role of Brr2 in circRNA biogenesis

Although it is now known that (i) circRNAs play crucial roles in genomic instability and DNA damage^38,40,56, and that (ii) a group of RNA helicases represent essential regulatory factors responsible for maintaining R-loop homeostasis under genotoxic stress^55,57,58, the question of whether some RNA helicases may protect genomic integrity through a circRNA-involved regulatory nexus remains unclear. Since RNA helicases can influence the metabolism of many species of RNAs, we sought to systematically identify RNA helicases involved in circRNA expression and conducted an RNAi screening on all Drosophila RNA helicases, which belong to the FlyBase (FB2020_01) GO term “RNA helicase activity”, in S2 cells using double-stranded RNAs (dsRNAs) (Fig. 1a). RT-qPCR was then performed to quantify the levels of 6 Drosophila circRNAs with variable genomic and transcript length, exon count, GC content and nuclear-cytoplasmic distribution (Fig. 1b and Supplementary Fig. 1a). Interestingly, Brr2 was identified as the only one that exhibited a robust inhibitory effect on all 6 examined circRNAs (Fig. 1c). Next, we repeated Brr2 knockdown (KD) experiments using two independent dsRNAs, both of which successfully silenced Brr2 expression in cells (Supplementary Fig. 1b), and 5 additional circRNAs were investigated in the assay (Fig. 1d). In line with the screening data, the levels of all 11 circRNAs were found to be elevated (Fig. 1d, row 1 and 2). Moreover, overexpression (OE) of Brr2 significantly down-regulated these circRNAs (Fig. 1d, row 3), which was opposite to the phenotype of Brr2 KD cells and thereby ruled out the unspecific effects of Brr2 KD. These data collectively indicate the general role of Brr2 in circRNA generation. We also tried to generate a Brr2 knockout (KO) cell line using the CRISPR/Cas9 genome editing, but all edited colonies failed to grow or even died after rounds of selection, suggesting that Brr2 is an essential gene indispensable for cell proliferation. Given that the upregulation of circRNAs could be a simple consequence of the increased transcriptional activity of their host genes, we performed an expression analysis for the cognate mRNAs of Brr2-regulated circRNAs and found that the levels of these mRNAs were either unchanged or decreased after KD of Brr2 (Fig. 1d, row 4 and 5). This indicates that Brr2 specifically regulates circRNA biogenesis rather than gene transcription. RNA FISH of circPlexA(2), circpan(11,12), circNipped-B(11,12), circSarm(1,2) and their linear counterparts further confirmed the specificity of Brr2-mediated regulation in circRNA biogenesis at the single-cell level (Fig. 1e, f; and Supplementary Fig. 1c, d). In the screening, we noticed that Prp5, pit, pea, Hel25E and Ddx56 also somehow showed an inhibitory effect on circRNA expression, at least for a small subset of circRNAs (Fig. 1c). However, the expression levels of the cognate mRNAs were similarly increased upon the individual KD of these helicases (Fig. 1g–i; Supplementary Fig. 1e, f). These data suggest that Prp5, pit, pea, Hel25E and Ddx56 seem to indirectly affect circRNA expression by regulating transcription. Taken together, we concluded that the RNA helicase Brr2 serves as a potent and specific circRNA biogenesis repressor in eukaryotic cells, and we next concentrated on Brr2-mediated regulation in the subsequent study.

**Fig. 1: RNAi screening of RNA helicases that control circRNA metabolism.**

Depletion of Brr2 extensively promotes circRNA biogenesis

To explore whether Brr2-mediated regulation represents a widespread mechanism for circRNA generation, total RNA samples from Brr2 KD and control cells were subject to Ribo-Zero Next-Generation Sequencing, and the CIRI2 software was utilized to identify circRNAs and calculate their expression levels⁵⁹ (Supplementary Data 1). Because (i) the vast majority of circRNAs are derived from back-spliced exons^22,23,24 and (ii) the presence of reads from lowly-expressed circRNAs reduces the sensitivity of bioinformatics analysis, we focused on 625 high-confidence EcircRNAs, each of which must contain more than 20 unique reads in all samples, for the subsequent genome-wide investigation. As observed, about 42.40% of EcircRNAs exhibited an altered expression upon Brr2 KD (Fig. 2a, left panel; Supplementary Fig. 2a). The cutoff for differentially expressed EcircRNAs was set as P < 0.05 and |log₂(fold change (Brr2 KD/Ctrl))| > 0.5. Notably, among these dysregulated EcircRNAs, 81.13% were substantially upregulated (Fig. 2a, right panel; Supplementary Fig. 2a).

**Fig. 2: The global circRNA abundance is regulated by Brr2.**

The dysregulation of circRNAs could be a consequence of the altered expression of their pre-mRNAs⁶⁰, which will lead to a parallel change in corresponding linear mRNAs. To test the possibility, we performed Pearson’s correlation analysis, which revealed that the expression of linear RNAs, including long non-coding RNAs (lncRNAs) and mRNAs, in all three Brr2 KD groups was positively correlated with that of in control groups, while the EcircRNA expression displayed a relatively week correlation (Fig. 2b; Supplementary Data 1). This indicates that Brr2-mediated regulation is more specific for circRNAs but not linear RNAs. Furthermore, we plotted the fold change in EcircRNA abundance against that in abundance of the cognate mRNAs and found that about 70.24% EcircRNAs are produced from pre-mRNAs to a much greater extent than their linear counterparts when Brr2 was depleted from cells (Fig. 2c). Indeed, the global cognate mRNA expression was marginally but significantly reduced in the absence of Brr2 (Supplementary Fig. 2b), which was distinct from the EcircRNA profile (Supplementary Fig. 2c). Protein-coding genes KCNQ and mbl are two pertinent examples (Fig. 2d), whose circularized exons (but not other exons) were substantially enriched upon Brr2 KD. To provide experimental support for the aforementioned bioinformatics analyses, we performed RT-qPCR to examine several of the strongly regulated EcircRNAs and confirmed that the levels of all tested circRNAs were increased 5 to 50-fold (Supplementary Fig. 2d, higher panel), while 7 of 10 corresponding linear mRNAs were significantly reduced (Supplementary Fig. 2d, lower panel). The expression inconsistency between circRNAs and their corresponding linear mRNAs demonstrated that the increase in circRNA production caused by Brr2 KD was not due to the enhanced transcription of the host genes. The transcription inhibition assay, during which Actinomycin D (Act D) was added to cells over time, revealed that the half-lives of tested EcircRNAs exhibited no difference between the control and Brr2 KD group (Fig. 2e; and Supplementary Fig. 2e, f), eliminating the possibility of circRNA accumulation caused by dysregulated degradation. Indeed, the nuclear run-on (NRO) assay confirmed that the nascent expression of examined EcircRNAs was drastically upregulated (Fig. 2f; and Supplementary Fig. 2g, h). In addition, RNA FISH (Fig. 2g; Supplementary Fig. 2i) and nucleocytoplasmic separation experiments (Fig. 2h; and Supplementary Fig. 2j–m) showed that the nuclear and cytoplasmic signals of EcircRNAs were increased to a similar degree between each other, ruling out the influence of Brr2 in nuclear export of EcircRNAs.

Unlike EcircRNAs, circular intronic RNAs (ciRNAs) originate from a subgroup of intron lariats that fail to be debranched and exon-intron circRNAs (EIciRNAs) are products with both exons and incompletely processed introns^25,26,27. Interestingly, a same expression pattern was observed with these intron-containing circRNAs in Brr2 KD cells compared to EcircRNAs (Fig. 2i, j; and Supplementary Data 1). By contrast, ~82.9% fusion circRNAs, formed by exons or introns from at least two genes^28,29,30, were downregulated (Fig. 2k; Supplementary Data 1), indicating that Brr2-mediated regulation strongly prefers EcircRNAs and intron-containing circRNAs. These observations were further verified by RT-qPCR experiments (Supplementary Fig. 2n, o).

Taken together, these data unveil the selective regulatory effect of Brr2 on EcircRNAs and demonstrate the active regulation of circRNA-linear RNA balance by Brr2.

Overall balance between back-splicing and linear splicing is largely determined by Brr2

It is very important to note that we probably underestimate the regulatory potency of Brr2 on circRNA biogenesis by simply examining the expression levels of circRNAs, which could be exemplified by circRNAs shown in the lower right part of the third quadrant of Fig. 2c. In these instances, even though the expression of circRNAs dropped upon Brr2 KD, their corresponding linear mRNAs decreased to an even greater extent (Supplementary Fig. 3a, b). These particular examples clearly support that certain genes, whose transcription was attenuated by Brr2 KD, still tended to generate circRNAs versus linear mRNAs. In fact, previous studies have demonstrated that the circularization event somehow competes with LS^43,60. In agreement, further correlation analysis revealed that the BS reads were positively shifted toward increased abundance when Brr2 was depleted from cells (Fig. 3a), while the fold changes of the LS reads displayed a modest decline tendency (Fig. 3a; and Supplementary Fig. 3c), implying that Brr2 is very likely a regulatory factor which determines whether a gene tends to undergo BS or LS. Informed by these reasons, we next asked whether Brr2 globally modulates the BS/LS ratio in the following investigation.

**Fig. 3: Brr2 specifically inhibits the BS reaction.**

To this end, we used the BS rate (BSR) to estimate the genome-wide competition between circRNA biogenesis and canonical splicing in the control and Brr2 KD samples. The BSR of a circRNA locus was defined as dividing the number of the BS reads by the total number of splicing reads of the site (Fig. 3b; and Supplementary Data 1)⁶¹. To ensure the accuracy of our analysis, we took 597 high-confidence EcircRNAs, which harbor both BS and LS reads at the site, into consideration. As observed, the overall BSR arose upon Brr2 KD (Fig. 3c; Supplementary Fig. 3d), and over 70% EcircRNAs showed a rather significant increased BSR (log₂(Brr2 KD/Ctrl) > 1 and P < 0.05) (Fig. 3d). More importantly, the increased BSR was also found for EcircRNAs, whose expression levels were either declined or unchanged upon Brr2 KD (Fig. 3e, f), demonstrating that the BS reaction was genome-wide enhanced by loss-of-function of Brr2, even the steady-state EcircRNA expression was reduced by attenuated transcription. We also made use of CIRCscore, an alternative strategy that uses fragments per billion mapped bases (FPB) as quantitation parameters (Supplementary Fig. 3e)⁶², to examine the altered BS/LS ratio caused by Brr2 KD, and found that the CIRCscore of EcircRNAs in Brr2 KD cells was generally higher than that in control cells (Fig. 3g; and Supplementary Data 1), which further confirmed that the Brr2 KD-induced BSR upregulation can cause deficient LS. Interestingly, the regulatory potency of Brr2 on BS was significantly weakened in EcircRNAs containing over 6 exons (Fig. 3h), which was not likely owing to the length of EcircRNAs (Fig. 3i). This suggests that Brr2 preferentially affects EcircRNAs composed of fewer exons. The group of genes with increased BSRs were enriched for housekeeping genes^63,64 with relatively high level of expression, including categories such as “regulation of transcription by RNA polymerase II” and “phosphorylation” (Fig. 3j, k; and Supplementary Data 1), implying that Brr2 preferentially functions at highly transcribed genes. Collectively, we concluded that Brr2 acts as a circRNA biogenesis inhibitor through controlling the balance between BS and LS.

As a subunit of U5-snPNP, Brr2 is typically thought to unwind U4/U6 duplex during the phase of spliceosome activation in eukaryotic cells^65,66. Therefore, we reasoned that Brr2 might regulate circRNA formation indirectly through the spliceosome. To test the hypothesis, we performed correlation analysis between the number of recognition motifs of spliceosome components in circularizing exons and the fold changes of circRNA expression or BSRs. No significant correlations were observed (Supplementary Fig. 4a; Supplementary Data 2). Moreover, the overall expression of Brr2-regulated EcircRNAs was not significantly altered after KD of several of essential factors of the canonical splicing machinery, including SF1, SNRNP40 and Hrb27c (Supplementary Fig. 4b). We also constructed a vector which can express a Brr2 truncated mutant (Brr2_ΔH1) deleted of a functional domain responsible for U4/U6 unwinding^67,68,69 (Supplementary Fig. 4c), and the wild-type (WT) and ΔH1 vectors were introduced individually into S2 cells to generate corresponding stable cell lines. As observed, the expression of all examined circRNAs was significantly and robustly inhibited in both the WT and ΔH1 stable cells, relative to regular S2 cells (Supplementary Fig. 4d). These data suggest that Brr2 regulates circRNA biogenesis independent of its canonical role in spliceosome activation.

We also explored the potential RNA regulons involved in Brr2-mediated regulation, and all unbiased short motifs of circularizing exons were estimated by 7-mers-based sequence quantitation, a bioinformatics algorithm that can calculate the occurrence frequency of a sequence of interest. Note that only 7-mers detected in more than 40 EcircRNAs were taken into consideration in the assay (Supplementary Data 2). The rank-ordered plot reflected that the BSRs of EcircRNAs containing 7-nt RNA short elements, such as “CAUUGAG”, “AAUCCGA” and “AUAUUGC”, were more sensitive to Brr2 KD (Supplementary Fig. 5a). Furthermore, we found that the fold change of BSR was positively correlated with the number of types of top 10 7-mers in a circRNA (Supplementary Fig. 5b). Consistently, EcircRNAs with more types of top 10 short elements have a larger BSR, indicating the combinatorial control by these 7-mers short elements in Brr2-mediated circRNA generation (Supplementary Fig. 5c).

Dual roles of Brr2 in preventing circRNA-mediated R-loop accumulation

CircRNAs can directly interact with the genomic DNA via base-pairing^38,39,40. For example, circSMARCA5 has been found to form a stable R-loop structure at exons 15-16 of its host gene⁴⁰. Therefore, we reasoned that the dysregulation of circRNA expression may be associated with R-loop formation, and that Brr2 probably dictates R-loop homeostasis through circRNAs. To test the assumption, we first utilized a mouse S9.6 monoclonal antibody, which is broadly used for R-loop detection and quantification⁷⁰, to compare the intensity of nuclear R-loop signals of the control and Brr2 KD/OE cells. Indeed, Brr2 KD led to a significant R-loop accumulation in nuclei (Fig. 4a, b), and Brr2 OE reduced the basal level of nuclear R-loops (Supplementary Fig. 6a). Note that we also observed some unspecific cytoplasmic S9.6 signals in the immunofluorescence staining (IF) assay, which was consistent to prior reports^56,71. To ascertain whether the nuclear fluorescence signals were truly from R-loops, we then used RNase H, an endonuclease that can specifically catalyze the cleavage of the RNA within an R-loop⁷², to pre-treat cells before S9.6 IF. As expected, the nuclear S9.6 signals were almost completely eliminated in both the control and Brr2 KD cells (Fig. 4a, b), thereby confirming the validity and accuracy of this assay. In addition, we constructed an expression vector that can produce a Flag-tagged Drosophila RNase H1 mutant with a single point mutation at the catalytic core (D252N) (Supplementary Fig. 6b), which abolished the enzymatic activity but retained the binding ability to R-loops⁷³. This mutant could be used as a direct sensor of R-loops in cells through Flag tag-based methods. Brr2 KD was subsequently performed in cells stably expressing the D252N vector. The Flag IF assay showed a notable increase in nuclear signals of the catalytic dead-RNase H1 in Brr2 KD cells versus the control cells (Fig. 4c, d). It was not because Brr2 KD affected the protein level of dead-RNase H1 (Supplementary Fig. 6c). These data collectively suggest that Brr2 opposes R-loop formation.

**Fig. 4: Accumulated R-loops caused by Brr2 depletion are largely ciR-loops.**

Since both circRNAs and linear RNAs are capable of forming R-loops (Fig. 4e), we next asked whether the Brr2 KD-induced R-loops were generated by accumulated circRNAs using cells pre-treated with RNase R, an exonuclease that can specifically digest linear RNAs but not circRNAs⁷⁴. Although RNase R can impair linear RNA-associated R-loops (Supplementary Fig. 6d–f)^39,56,75, the increased R-loop formation was still observed in the nuclei of Brr2 KD cells after RNase R digestion (Fig. 4f, g). In particular, whether or not there was pretreatment with RNase R had very limited effect on the fold change of R-loops (Fig. 4f, g). Moreover, considering that the half-lives of linear RNAs are much shorter than that of circRNAs^41,42,43, we performed Act D treatment (0.5, 1 and 4 hr) to reduce the overall abundance of linear mRNAs in cells prior to S9.6 IF. In line with what was observed with RNase R treatment, the high level of the nuclear S9.6 signals was only modestly affected in Brr2 KD cells after Act D treatment (Fig. 4h, i; Supplementary Fig. 6g, h). In fact, the overall linear mRNA expression was almost unchanged upon Brr2 KD (Supplementary Fig. 6i), which extremely reduces their chances to induce more R-loops. Taken together, these results provide evidence that the R-loops generated in Brr2 KD cells were primarily formed by circRNAs. These circRNA-associated R-loops were referred to ciR-loops herein.

Drosophila Brr2 is about 74% identical to its human homolog SNRNP200 (Supplementary Fig. 7a). Likewise, KD of SNRNP200 resulted in a significant increase in the expression levels of all examined human EcircRNAs (Supplementary Fig. 7b), and led to the increased formation of R-loops (Supplementary Fig. 7c, d) that were sensitive to RNase H digestion (Supplementary Fig. 7c–g). In addition, a large proportion of R-loop signals remained in the nuclei of SNRNP200 KD cells pre-treated with RNase R (Supplementary Fig. 7c, d), which verified that the SNRNP200 KD-induced R-loops were mainly ciR-loops in human cells and further confirmed the evolutionarily-conserved role of Brr2 in prevention of ciR-loop formation.

It is widely acknowledged that certain RNA helicases have an RNA-DNA unwinding activity and are able to directly impair R-loop structure through the conserved helicase domain in higher eukaryotes^1,2,76. For example, DDX39B (also known as Hel25E in Drosophila) was recently reported as an R-loop resolvase in HeLa and S2 cells^56,77. Informed by this, we raised two potential mechanisms for Brr2-mediated R-loop regulation. One was merely based on the inhibitory role of Brr2 in circRNA biogenesis, which potently reduces the overall abundance of circRNAs responsible for ciR-loops formation. The other was that Brr2 may additionally function as a direct R-loop-unwinding factor (Fig. 5a). To distinguish between these hypotheses, we utilized the Brr2_ΔH1 stable cell line which can stably overexpress a Brr2 truncated mutant deleted of its helicase domain in the following study. Of note, we found that, in comparison with the WT control, the Brr2_ΔH1 mutant failed to prevent R-loop accumulation induced by KD of two different unrelated factors (i.e., Hel25E and RNase H1) (Fig. 5b, c; and Supplementary Fig. 8a, b). Furthermore, we took advantage of biotin-labeled synthetic R-loops (RNAs within R-loops were labeled with biotin) to perform an in vitro unwinding assay with the WT Brr2 and its ΔH1 mutant (Fig. 5d; and Supplementary Fig. 8c). Similar to the single-cell phenotype assessed by S9.6 IF, the WT efficiently unwound R-loops in vitro, whereas the ΔH1 mutant failed to do so (Fig. 5d). It is important to point out that Brr2_ΔH1 still possessed a strong inhibitory effect on circRNA biogenesis as the WT did (Supplementary Fig. 4c, d). These data thus support that the accumulated R-loops in Brr2 KD cells were not a simple consequence of the increased circRNA level and suggest the dual actions of Brr2 in regulating R-loop metabolism.

**Fig. 5: Dual actions of Brr2 in avoiding nonscheduled ciR-loop accumulation.**

Brr2 prevents R-loop-mediated DNA damage

Emerging evidence has proved that deleterious R-loops represent a major source of DNA damage, and that nonscheduled formation and accumulation of R-loops can trigger DNA breaks and threaten genome stability^1,2,3,78,79. Therefore, we first conducted γH2A.V (a classic marker used to evaluate the genomic integrity of individual Drosophila cells^80,81) IF and western blotting experiments to assess the impact of Brr2 on genome stability. As observed, the nuclear fluorescence signals of γH2A.V in Brr2 KD cells were much higher than those in the control cells (Fig. 6a, b), and western blotting yielded a similar phenotype (Supplementary Fig. 9a). Next, we asked whether this increase was dependent on R-loops. For this purpose, we performed Brr2 KD experiments using cells stably overexpressing RNase H1. Notably, the phenotype of the increased γH2A.V level was no longer observed upon Brr2 KD (Fig. 6a, b; and Supplementary Fig. 9b), supporting that the accumulated ciR-loops induced by Brr2 KD may serve as a driver for genomic instability. Brr2 KD had no effect on the overexpression level of RNase H1 in the stable cells (Supplementary Fig. 9c), ruling out the possibility of dysregulated RNase H1 in the assay. To explore the evolutionarily-conserved role of Brr2 in regulating R-loop dependent DNA damage, we conducted same experiments using human cells. As expected, SNRNP200 KD caused a significant increase in the γH2A.X (the homolog of Drosophila γH2A.V) level (Fig. 6c, d; and Supplementary Fig. 9d), and the additional comet assay (i.e., single-cell electrophoresis) further confirmed the serious consequence in the genome caused by SNRNP200 KD (Fig. 6e, f). Consistent with what was observed with Drosophila cells, the accumulated γH2A.X signals caused by SNRNP200 KD were rescued by transfection with a vector overexpressing RNase H1 (Fig. 6g–i). Taken together, we demonstrated that Brr2 plays an essential and evolutionarily-conserved role in preventing R-loop-mediated genome instability.

**Fig. 6: Brr2 protects cells from R-loop-dependent DNA damage.**

Brr2 depletion leads to antisense transcription and premature transcriptional termination

Apart from constituting a source of genome instability, R-loops are critical for the regulation of transcriptomic integrity in eukaryotes^1,2,3,82. For example, some TSS proximal R-loops possess an intrinsic promoter activity to induce Pol II transcription of antisense RNAs⁸³. Since there are usually two distinct pre-initiation complexes (PICs) simultaneously assembled in opposite orientation in the active promoter-proximal regions^84,85,86,87, the aforementioned antisense transcription could be partly explained by the mechanism that Pol II is blocked by an R-loop downstream of the TSS in the sense direction, leading to more Pol II being recruited to the antisense direction. Having established that Brr2 KD resulted in a robust ciR-loop formation (Figs. 4 and 5; Supplementary Fig. 6–8), we thus proposed that Brr2 may somehow function in antisense transcription initiation and obtained the reads of antisense transcripts (ATs) generated from DNA 500 bp in front of the TSS region to analyze the activity of antisense transcription (Supplementary Fig. 10a). As observed, the AT transcription of about 26.94% EcircRNA host genes was significantly activated upon Brr2 KD (Fig. 7a; Supplementary Data 3), using the Pde11 gene as an example (Fig. 7b). These EcircRNA host genes were referred to AT-increased genes herein. Further RT-qPCR analysis confirmed the Brr2 KD-induced upregulation of EcircRNA-associated ATs at the single gene level (Supplementary Fig. 10b). Notably, over 93% AT-increased genes showed an increased BSR upon Brr2 KD (Fig. 7c), implying that the upregulation of circRNAs caused by Brr2 KD may contribute to the activation of AT transcription. In support of this, overexpression of circPde11(3,4,5) and circPIP4K(2,3,4,5,6) led to an increased level of the ATs of Pde11 and PIP4K, respectively (Supplementary Fig. 10c). More importantly, circRNA loci from AT-increased genes with a BSR_{Brr2 KD}/BSR_Ctrl ratio over 3 had shorter distance to the TSS than those from AT-unchanged or decreased genes (Fig. 7d; Supplementary Fig. 10a). By contrast, we did not observe a similar phenomenon in gene groups with a BSR_{Brr2 KD}/BSR_Ctrl ratio less than 3 (Fig. 7e). These observations were consistent with the theory of TSS proximal R-loop-mediated antisense transcription⁸³.

**Fig. 7: Depletion of Brr2 results in an altered transcriptomic profile.**

Transcription elongation is a discontinuous molecular process during which Pol II may be paused, stalled or even improperly released at different genomic sites, such as physical barriers imposed by specific chromatin structures⁸⁸. Indeed, a recent study has demonstrated that nonscheduled R-loops formed by linear mRNAs can impair transcription elongation by inducing PTT within the gene body region⁸⁹. Due to the covalently closed structure of circRNAs, ciR-loops seem to be more stable than R-loops from linear RNAs. Therefore, we reasoned that at least a subset of accumulated circRNAs induced by Brr2 KD may cause PTT within the body of their host genes. To test the possibility, we took advantage of the premature termination rate (PTR), which was calculated by taking the read density in the gene body upstream from a circRNA locus divided by the downstream read density (Supplementary Fig. 10d), to evaluate the PTT effect induced by Brr2 KD in the following analysis. The host genes of about 24.38% EcircRNAs were found to have a significantly increased PTR_{Brr2 KD}/PTR_Ctrl ratio, and thus these EcircRNAs were considered to effectively trigger PTT upon Brr2 KD (Fig. 7f; Supplementary Data 3). The host genes of these EcircRNAs were referred to PTR-increased genes herein. In particular, the BSRs of these PTR-increased genes were mostly upregulated (Fig. 7g). Take circAlh(2,3,4,5) as an example. The read density at sites downstream of the circularizing exons were robustly declined after Brr2 KD (Fig. 7h). RT-qPCR analysis examining the upstream/downstream expression ratio of the circrl(6,7,8), circeIF5B(7,8), circAlh(2,3,4,5) and circPIP4K(2,3,4,5,6) locus yielded a phenotype which was consistent with the bioinformatics data (Supplementary Fig. 10e). Moreover, for PTR-increased genes, the distance between the TSS and the circRNA locus was much shorter in comparison with genes with decreased PTRs (Fig. 7i). Importantly, the phenotype was particularly obvious among genes with increased BSRs (Fig. 7j), but not for genes with unchanged or decreased BSRs (Fig. 7k). These data suggest that circRNAs, whose loci are closer to the TSSs, are more likely to induce PTT upon Brr2 KD. More interestingly, for the host genes with increased BSRs, the activity of antisense transcription was positively correlated with the extent of PTT, since PTR-increased genes tended to have a higher AT expression level compared to others (Fig. 7l). Overall, our data propose that (i) loss-of-function of Brr2 can result in a dysregulated transcriptomic profile, such as an increased extent of antisense transcription and PTT, and that (ii) the transcription directionality of at least a subset of genes can be determined by the Brr2-circRNA regulatory axis (Fig. 7m), which expands our view of additional regulatory pathways for the precise coordination of transcription programs.

Brr2-mediated regulation is essential for proper DNA replication, cell division and cell proliferation

DNA replication forks can be impeded by harmful R-loops, leading to replication stress and DNA damage in a cell^90,91,92,93. In order to gain an insight of Brr2-mediated regulation in DNA replication, 5-Ethynyl-2-deoxyuridine (EdU), a thymidine analog, was utilized to label the replicated DNA and compare the DNA replication activity of the control and Brr2 KD cells. As observed, the EdU signal was significantly reduced upon Brr2 KD, suggesting that loss-of function of Brr2 impairs the DNA replication process (Fig. 8a). The phenotype was also observed in human cells after SNRNP200 KD (Supplementary Fig. 11a). Of note, Brr2 KD no longer triggered a higher level of the DNA replication defect in cells stably overexpressing RNase H1 (Fig. 8b) but not its catalytic-dead mutant (Fig. 8c), which confirmed that the DNA replication defect was closely related with the harmful nuclear-accumulated ciR-loops induced by Brr2 KD. Considering that the proper control of DNA replication is a prerequisite for a successful cell division, we next explored the relationship between Brr2 and cell cycle. We used human MCF7 cells to investigate the process of cell cycle and chromosome segregation, as they are a type of adherent cells and an ideal model with well-defined cell cycle. As expected, SNRNP200 KD arrested cells at the S and G2/M phase (Fig. 8d; and Supplementary Fig. 11b). Moreover, chromosome segregation errors, such as multipolar mitoses and polar asymmetry, exhibited a rather higher frequency of the occurrence in the SNRNP200 KD cells in comparison with the control (Fig. 8e), and, correspondingly, cell proliferation was significantly inhibited upon Brr2 KD (Fig. 8f; Supplementary Fig. 11c). It was reminiscent that all Brr2 KO cell colonies failed to grow after rounds of selection. Importantly, these errors or defects were rescued by overexpression of RNase H1 (Fig. 8g, h; Supplementary Fig. 11d) rather than its catalytic-dead mutant (Supplementary Fig. 11e). These results further support that Brr2 ensures cell division through its role in R-loop metabolism.

**Fig. 8: Brr2-mediated regulation in cell division and proliferation.**

Discussion

Despite being considered as canonical post-transcriptional modulators of RNA metabolism^94,95,96, a group of RBPs can function directly or indirectly in the transcriptional process, which is becoming the center of attention in the field of RNA biology lately^{50,51,52,53,97,98}. A pertinent example in point is the chromatin-interacting RBP gawky (also known as GW182), which acts as a universal cytoplasmic decay factor for both linear and circular transcripts under unstressed conditions^42,99, but rapidly moves into the nucleus to induce a huge transcriptional reprogramming especially for stress-responsive genes once cells encounter adverse stimuli¹⁰⁰. The functional diversity of versatile RBPs raises a possibility that some of them may be ideal monitors for biological events that require multilayered surveillance at different stages. This theory is exactly applicable to R-loop homeostasis, which needs the precise regulation of both R-loop formation and removal^1,2,3. In particular, ciR-loops are more stable than R-loops generated from linear RNAs (i.e., ciR-loops are more resistant to Act D treatment) (Fig. 4h, i; and Supplementary Fig. 6g, h) and thus seem to be more dependent on a rapid, strong and efficient regulatory mode. Indeed, we here found that ciR-loop homeostasis can be tightly-controlled by the DEAD-box RNA helicase Brr2 at two distinct levels (Fig. 9a). On the one hand, Brr2 inhibits circRNA (but not its parental linear mRNA) expression by specifically repressing BS, which immensely reduces the RNA source for ciR-loop formation. On the other hand, Brr2 functions as an unwinding factor at chromatin, where it resolves accumulated harmful circRNA-DNA hybrids. Notably, the Brr2_ΔH1 mutant, despite losing the ability to resolve R-loops (Fig. 5b–d; and Supplementary Fig. 8a, b), still potently inhibited circRNA biogenesis (Supplementary Fig. 4c, d), indicating that the two roles of Brr2 rely on distinct domains and thus are independent of each other. The dual roles of Brr2 support a theory that a single RBP could act as a vital bridge for rapidly and effectively integrating transcriptional and post-transcriptional regulatory events, eventually ensuring cellular homeostasis.

**Fig. 9: A model for the essential role of Brr2 in ciR-loop homeostasis and genome stability.**

Prevailing evidence has demonstrated that the overabundance of R-loops, in most cases, is a key reason to induce aberrant transcription events^1,2,3,82. For example, some R-loops around TSSs were reported to have the promoter activity to recruit Pol II and multiple transcription factors (TFs) and in turn facilitate the transcription of a substantial proportion of ATs⁸³. Here, we indeed found that loss-of-function of Brr2 significantly activated the antisense transcription of a subset of circRNA-encoding genes. More consistently, genes with circRNA loci that are closer to TSSs yielded a stronger antisense transcription upon Brr2 KD (Fig. 7a–e). Besides, we observed an induced PTT phenotype in Brr2 KD cells (Fig. 7f–k). In line with our observations, R-loops formed by exogenous antisense oligonucleotides (ASOs) were found to release Pol II at ASO-target sites and induce a PPT event in the sense direction through a 5’ exonuclease-mediated pathway (also known as the “torpedo” mechanism), in which the target nascent RNA is cleaved by RNase H and the resulted uncapped RNA fragment is then degraded gradually by the Drosophila 5’ exonuclease Rat1 (or its human homolog XRN2) until Rat1/XRN2 catches up and dissociates elongating Pol II from chromatin^101,102,103. A following study has confirmed that naturally generated R-loops can trigger PTT genome-wide through a similar mechanism⁸⁹. Furthermore, R-loops are able to induce PTT by establishing repressive chromatin compaction and histone modifications where they are located^104,105,106. For example, R-loops at the human β-actin gene can act as a signature/platform for the recruitment of components of the RNAi machinery, thereby generating the H3K9me2 repressive mark for efficient transcriptional termination¹⁰⁴. Informed by these findings as well as the structural nature of R-loops, we speculate that R-loops could be physical obstacles in sense strand of the dual-orientation promoters. Because PICs are typically loaded on both the sense and antisense strands at the same time in the vicinity of the promoter regions^84,85,86,87, the upregulated antisense transcription caused by Brr2 KD is very likely a result of blocked transcription by ciR-loops in the sense direction. In support of this idea, the Brr2 KD-induced PTT tended to occur at genes with TSS-proximal circRNA loci and the incidence of PTT was positively correlated with that of antisense transcription (Fig. 7i–l). Note that the current understanding of divergent transcription is largely limited to U1-polyadenylation signal (U1-PAS)-mediated model, which can be explained by the asymmetric distribution of U1 and PAS sites flanking the TSS^107,108. Our work, together with other recent reports, implies that the transcription directionality of at least a subset of eukaryotic genes may also be partially defined by ciR-loops and Brr2 (Fig. 9b), thereby providing a novel clue into the diverse regulation of bidirectional promotors.

Nonscheduled pathological R-loops can also interfere with genome integrity and are a major genomic threat^1,2,3,78,79. This was initially documented by several early studies about the downstream phenotypes of mRNP processing or export defects, which give rise to an increased extent of R-loop-dependent DNA damage^109,110,111. The widely recognized mechanism behind this is that R-loops can serve as strong barriers to stall the progression of DNA replication forks and ultimately induce DNA breakage at forks^90,91,92,93. Focusing on a new class of R-loops, the present study proposes that ciR-loops are also a non-negligible inducement of DNA damage and replication stress (Fig. 9b), which implies that ciR-loops may induce DNA instability through a mechanism mirroring the one by R-loops generated from linear mRNAs. However, this assumption needs deeper investigation in the future, since circRNAs within R-loops might bind and recruit endonucleases in an R-loop independent manner. Moreover, we uncovered that the mitotic segregation disorders, including multipolar mitoses and polar asymmetry, and the impaired cell proliferation are closely accompanied with Brr2 KD-induced ciR-loop accumulation (Fig. 8e–h), which extends the understanding of the downstream impacts of diverse R-loop types.

It is also interesting to note that many circRNAs function as transcription/genome regulators through a variety of mechanisms^22,23,24. For example, two EIciRNAs (circEIF3j and circPIPK3) can form a complex with U1 snRNP via base-pairing (RNA-RNA interaction), which further facilitates the loading of Pol II and general TFs to promoters of their host genes to initiate transcription²⁵; a class of metal-responsive element-containing circRNAs (MRE circRNAs) are capable of repressing stress-induced transcription as a group by sequestering the transcription activator gawky away from metal-inducible genes¹¹²; circARHGAP35 indirectly affects oncogene activation through its encoded protein which binds and modulates the functional activity of the TF TFII-I in the nucleus¹¹³. Here, our study suggests that the formation of ciR-loops represents an alternative and pervasive mode for circRNA-involved gene regulation. In fact, individual circRNAs have been recently reported to have the capacity to generate R-loops at the single gene level^38,39,40. A case in point is circMLL which can hybridize with the complementary DNA strand of the MLL locus, thereby slowing or even shutting down Pol II elongation near R-loop sites³⁸. Our present study is different in that it reveals the connection between perturbations of overall circRNA abundance and gene transcription activity as well as genome stability at the genome-wide scale. Based on these findings, we speculate that, in principle, any factors that globally influence circRNA life span (e.g., degradation and subcellular localization) may alter ciR-loop homeostasis and genome stability to some extent. Nevertheless, the question of whether there are additional Brr2-like dual-action regulators that control both ciR-loop formation and removal is unclear. Therefore, extensive efforts are required to fill the gap.

Methods

Cell culture

S2 cells used in the study were a gift from Dr. Qingfa Wu’s lab (University of Science and Technology of China), and were cultured in Schneider’s Drosophila Medium (SDM; Sigma, S9895) containing 10% (v/v) fetal bovine serum (FBS; Excell Bio, FSP500) and 1% (v/v) penicillin-streptomycin (Thermo Fisher Scientific, 15140122) at 25 °C. The human breast adenocarcinoma cell line (MCF7) used in the study were a gift from Dr. Aimin Yang’s lab (Chongqing University), and were maintained in high-glucose Dulbecco’s modified Eagle’s medium (Biosharp, BL301A) supplemented with 10% (v/v) FBS (ExCell Bio, FSP500) and 1% (v/v) penicillin-streptomycin (Thermo Fisher Scientific, 15140122) at 37 °C with 5% CO₂.

RNAi

To KD the indicated Drosophila protein-coding genes, 1.5 × 10⁶ S2 cells were incubated with 8 μg of corresponding dsRNA in 600 μL serum-free medium for 30 min, and then 400 μL of medium supplemented with 20% (v/v) FBS was added to maintain cell growth. For RNAi screening of all Drosophila RNA helicases, untreated S2 cells served as a negative control. For RNAi experiments targeting other Drosophila protein-coding genes, cells treated with β-gal dsRNA served as a negative control. After 3 days of treatment, the cells were harvested for later use. For dsRNA preparation, DNA templates for dsRNAs selected from the Drosophila RNAi Screening Center (DRSC: https://www.flyrnai.org/cgi-bin/DRSC_gene_lookup.pl) were obtained by polymerase chain reaction (PCR) using primers incorporating the T7 promoter sequence (TAATACGACTCACTATAGGG) at the 5ʹ end. DsRNAs were subsequently generated through in vitro transcription using the ScriptMAX® Thermo T7 Transcription Kit (TOYOBO, TSK-101) according to the manufacturer’s protocol. The primer pairs used for dsRNA generation are provided in Supplementary Data 4. For siRNA-mediated KD, 50 nmol of the indicated siRNAs (SNRNP200 KD: 5ʹ-GCCUACCUCUAUAUCCGAAUG-3ʹ; SNRNP200 KD2: 5ʹ-GUGAUUCAGAUUGAGUCCU-3ʹ) or negative control siRNAs were transfected into MCF7 cells using LipoRNAi™ (Beyotime, C0535) according to the manufacturer’s instructions.

RNA isolation, complementary DNAs synthesis and RT-qPCR

RNA was isolated using RNAiso Plus reagent (TaKaRa, 9108). Complementary DNAs (cDNAs) were synthesized using PrimeScript RT Master Mix (TaKaRa, RR036A). For the reverse transcription reaction of antisense transcripts, the Hifair^® III 1st Strand cDNA Synthesis Kit (gDNA digester plus) (YEASEN, 11119ES60) was employed with gene-specific primers. cDNA was subjected to qPCR analyses using Hieff^® qPCR SYBR Green Master Mix (YEASEN, 11201ES03) on a CFX Connect Real-Time PCR System (Bio-Rad). Detailed information regarding all RT-PCR and RT-qPCR primers used in this study is provided in Supplementary Data 4.

Vectors and stable cell line construction

The circRNA expression vectors used in this study were generated by cloning the indicated sequences into the Hy_pMtnA_laccase2 MCS vector. Drosophila protein-coding gene expression vectors were generated by inserting the indicated sequences of Drosophila protein-coding genes into the Hy_pMtnA/pAct Flag MCS vector. Prokaryotic expression vectors were generated by inserting the indicated sequences into the pQE-80L vector. Human protein-coding gene expression vectors were generated by inserting the indicated sequences into the pcDNA3.1_3 × Flag vector. Detailed information for all vectors used in this study is provided in Supplementary Data 5. Transfection of expression vectors was performed with Lipo6000^TM reagent (Beyotime, C0526) according to the manufacturer’s instructions. Stable cell lines expressing the indicated genes in S2 cells were generated by selection with 150 μg/mL of hygromycin B (Biofroxx, 1366ML010) for 3-4 weeks following transfection with the indicated vectors for 3 days. To induce expression, a final concentration of 500 μM copper sulfate (CuSO₄; MACKLIN, C805782) was added and incubated for 12 hr.

RNA sequencing (RNA-seq) and bioinformatic analysis

The quality of total RNA extracted from Brr2 KD and control cells (β-gal KD) was verified using the Agilent Bioanalyzer 2100 system. Qualified RNA was then subjected to Ribo-Zero Next-Generation Sequencing. Briefly, ribosomal RNA (rRNA) was removed by TIANSeq rRNA Depletion Kit (Animal) (TIANGEN, NR101-T1), and surplus RNA was purified and subjected to library construction using TruSeq Ribo Profile Library Prep Kit (Illumina, 20020599). Paired-end sequencing (150 nt) was performed using an Illumina NovaSeq 6000 System (Novogene, China).

Adapter sequences and low-quality reads were removed using Trimmomatic (version 0.36)¹¹⁴ with parameters LEADING:3 TRAILING:3 SLIDINGWINDOW:4:15 MINLEN:36. The cleaned reads were then aligned to the reference genome (Drosophila melanogaster genome (BDGP6)) from Ensembl (https://www.ensembl.org) using TopHat2 (version 2.1.0)^115,116. Gene expression levels for each library were estimated using StringTie (version 2.1.7)¹¹⁷. Differential gene expression analysis was performed using DESeq2 R package (version 1.32)¹¹⁸. BigWig files with FPKM normalization were generated from the BAM files using deepTools (version 3.3.0)¹¹⁹ and visualized using Integrative Genomics Viewer (IGV)¹²⁰. CIRI2 (version 2.0.6) with default parameters was employed to identify circRNAs⁵⁹. CircRNAs with more than 20 BS junction reads across all 6 samples (3 β-gal KD and 3 Brr2 KD samples) were retained for further analysis. Differential circRNA expression analysis was performed using the DESeq2 R package (version 1.32), and statistical significance was determined using the two-sided Wald chi-squared test in R software. The annotation of circRNA was based on the location of circRNA’s BS junction by using R software. The length, GC content, and exon number of circRNA were calculated by R software based on the annotation of circRNA.

To obtain housekeeping genes, gene expression data encompassing 30 developmental time points and conditions of Drosophila melanogaster, generated by the modENCODE Consortium⁶³, were analyzed. A gene was defined as housekeeping if its expression exceeded the 50th percentile of expression in every condition^63,64. This resulted in 4617 housekeeping genes. Gene Ontology (GO) analysis was carried out using DAVID bioinformatics resources¹²¹.

Subcellular localization analysis of circRNA

To quantify the subcellular localization of circRNAs, we normalized the circRNA-level CPM output using a method that employs the whole cell extract (WCE) samples to fit normalization coefficients for cytoplasm and nucleus samples¹²². WCE, nuclear, and cytoplasmic data were collected from our previous dataset (BioProject: PRJNA1039137)¹¹². Briefly, circRNAs with expression levels in the top 50 or bottom 50 of the nuclear samples were excluded. The remaining circRNAs were subjected to linear regression analysis to calculate regression coefficients. Finally, absolute localization values were obtained by normalizing the cytoplasmic and nuclear data using these coefficients.

BS and LS analysis

CIRI2 (version 2.0.6.) with default parameters was used to identify the BS sites and obtain the associated BS reads and LS junction reads (exon-exon junction reads) of each BS site. All LS sites and LS junction reads of a gene were identified using TopHat2 (version 2.1.0)¹¹⁵ with default parameters. The identified LS sites and associated LS junction reads were then annotated using BEDTools (version 2.15.0)¹²³.

CIRCscore

Fragments per billion mapped bases (FPB) were used to evaluate circular and linear RNA expression individually by counting fragments mapped to circRNA-specific BS sites or linear RNA-specific splicing junction sites⁶². Based on this, a CIRCscore was generated by dividing FPB_circ by FPB_linear, which could indicate the relative circRNA expression level using linear RNA expression level as the background.

Motif and 7-mers enrichment analysis

For analyzing the enrichment of known RNA binding motifs¹²⁴ in circRNAs regulated by Brr2, FIMO¹²⁵ was used to detect the number of examined motifs in each circRNA sequence. For 7-mers analysis, Jellyfish (version 2.3.1)¹²⁶ was used to count 7-mers in each circRNA sequence. 7-mers with at least 40 occurrences across all circRNA sequences were retained for further analysis.

Antisense transcription analysis

To analyze antisense transcription of the circRNA’s parental gene, the read density within the 500 bp antisense region upstream of the transcription start site (TSS) of the circRNA’s parental gene was analyzed, and differential expression analysis was performed. AT regions with more than 40 reads across all 6 samples (3 β-gal KD and 3 Brr2 KD samples) were retained for further analysis.

Premature transcription termination analysis

For analyzing the effect of circRNA on the premature transcription termination of its parental gene, the read density in the gene body upstream and downstream from a circRNA locus was obtained using StringTie (version 2.1.7)¹¹⁷. PTR was calculated by dividing the read density in the gene body upstream of a circRNA locus by the downstream read density. CircRNAs without upstream or downstream regions were excluded. The fold change of PTR upon Brr2 KD was calculated by dividing PTR_{Brr2 KD} by PTR_Ctrl.

Nuclear and cytoplasmic fractionation

Cellular fractionation of Brr2 KD and control (β-gal KD) cells was performed as previously described^100,127. Briefly, Brr2 KD or control cells were washed twice with 1 mL of ice-cold 1 × PBS buffer (phosphate buffer saline, pH 7.4, 137 mM NaCl, 2.7 mM KCl, 10 mM Na₂HPO₄ and 1.8 mM KH₂PO₄) and resuspended in ice-cold lysis buffer B (10 mM Tris-HCl, pH 8, 1.5 mM MgCl₂, 140 mM NaCl, 0.5% IGEPAL CA-630 (v/v) and 1 mM dithiothreitol). Sample separation was performed by centrifugation at 1000 × g for 3 min at 4 °C. The supernatant was collected as the cytoplasmic fraction, and the crude nuclei were pelleted in 1 mL of ice-cold lysis buffer B supplemented with 100 μL of detergent (3.3% (w/v) sodium deoxycholate, 6.6% (v/v) Tween 40). After 10 s of slow vortexing and 5 min of incubation on ice, the pellets were collected by centrifugation at 1000 × g for 3 min at 4 °C. The collected pellets were then washed with 1 mL of ice-cold lysis buffer B. The final washed pellets served as the nuclear fraction. Efficient fractionation of nuclear and cytoplasmic RNAs was verified by measuring levels of U6 RNA and 18S RNA.

Estimating RNA half-life

To measure circRNA half-lives, Brr2 KD and control cells were treated with Act D (1 μg/mL; Sigma, A4262) for the indicated amounts of time (0, 2, 4, and 8 hr). RNA was then extracted, and RT-qPCR was performed to quantify the expression of indicated circRNAs. Data were normalized to the circRNA levels observed at 0 hr of Act D treatment.

NRO assay

NRO assay was used to quantify the expression of nascent nuclear circRNA transcripts and was conducted as previously described with minor modifications^54,100,128. Briefly, after a total of 2.5 × 10⁷ S2 cells had been treated with 24 μg of Brr2 or β-gal dsRNA for 3 days, the cells were harvested and washed twice with 1 mL of ice-cold 1 × PBS buffer. Subsequently, nuclear and cytoplasmic fractionation was performed to obtain the nuclear fraction. Nuclei were then resuspended into 100 μL of nuclear run on buffer (50 mM Tris-HCl, pH 7.5, 5 mM MgCl₂, 150 mM KCl, 0.1% srkosyl (w/v), 10 mM dithiothreitol and 80 units/mL RNase inhibitor) containing each of 1 mM NTPs (ATP, GTP, CTP, and BrUTP (Sigma, B7166)). The mixture was incubated at 28 °C for 5 min to allow the incorporation of BrUTP into nascent transcripts. 500 μL of RNAiso Plus were then added to terminate the NRO reaction and isolate nuclear RNA. The nuclear RNA was resuspended into 500 μL of RNA immunoprecipitation buffer (20 mM Tris-HCl, pH 7.5, 200 mM NaCl, 2.5 mM MgCl₂, 0.5% NP40 (v/v) and 10% glycerol (v/v)) and incubated with 2 uL DNase Ⅰ. BrUTP-labeled nascent RNA was then purified using 500 μL of RNAiso Plus and pulled down using Dynabeads® Protein G (Thermo Fisher Scientific, 10003D) and anti-BrUTP (Abcam, ab1893).

Western blotting

Harvested cells were washed twice with 1 mL of ice-cold 1 × PBS buffer. Protein extraction was performed using RIPA buffer (50 mM Tris-HCl, pH 7.4, 150 mM NaCl, 0.1% sodium dodecyl sulfate (SDS; w/v), 1% sodium deoxycholate (w/v), and 1% Triton X-100 (v/v)). Protein samples mixed with protein loading buffer (62.5 mM Tris-HCl, pH 6.8, 10% glycerol (v/v), 0.01% bromophenol blue (w/v), 2.15% SDS (w/v), 1.55% dithiothreitol (w/v), and 5% 2-hydroxy-1-ethanethiol (v/v)) were denatured at 100 °C for 5 min. The denatured samples were then separated by SDS-PAGE, and transferred onto polyvinylidene fluoride (PVDF) membranes (BioRad, 1620177). Non-specific binding was blocked by incubating with 5% milk in TBST buffer (Tris-buffered saline with Tween-20, pH 7.6, 0.242% Tris-HCl (w/v), 0.8% NaCl (w/v), and 0.05% Tween-20 (v/v)) for 1 hr at room temperature. After incubating with primary antibodies overnight at 4 °C, the membranes were incubated with secondary antibodies at room temperature for 1 hr. The membranes were then subjected to the ECL protocol using the Bio-Rad ChemiDoc Imaging System. Protein level quantifications were carried out by ImageJ¹²⁹. Antibodies used in this study were: anti-FLAG (Beyotime, AF519; 1:2500 dilution), anti-γH2A.X (Beyotime, AF1201; 1:2500 dilution), anti-α-Tubulin (Beyotime, AF0001; 1:2500 dilution), anti-SNRNP200 (Santa Cruz Bitotechnology, sc-393170; 1: 500 dilution), HRP-labeled goat anti-rabbit IgG (H + L) (Beyotime, A0208; 1:3000 dilution), and HRP-labeled goat anti-mouse IgG (H + L) (Beyotime, A0216;1:3000 dilution). Uncropped and unprocessed scans of blots are provided in the Source Data file.

FISH

RNA probes targeting the BS site of circRNAs were generated through in vitro transcription using the ScriptMAX® Thermo T7 Transcription Kit (TOYOBO, TSK-101) according to the manufacturer’s protocol. The probes were then labeled with Alexa Fluor 546 dye using the ULYSIS® Nucleic Acid Labeling Kit (Thermo Fisher Scientific, U21652) and denatured at 95 °C for 5 min before use. FISH was carried out as previously described^112,130. Briefly, cells were seeded on a coverslip coated with concanavalin A (ConA; Solarbio, C8110) and fixed in fixative solution (75% methanol (v/v) and 25% glacial acetic acid (v/v)) at room temperature for 10 min. The cells were then permeabilized with 0.1% Triton X-100 (v/v) in TBST buffer. Permeabilized cells were denatured at 80 °C for 3 min. Subsequently, they were incubated with right amount of denatured probe at 42 °C overnight. Finally, the cells were stained with 1 μg/mL DAPI (Beyotime, C1002) for 5 min. Fluorescence signals were detected using the Leica TCS SP8 system. The integrated density of the whole-cell and nuclear FISH signals served as the analytical parameter in this study. Nuclear regions were defined by manual tracing of DAPI staining, and cellular regions were defined by manual tracing of cells in bright-field. The whole-cell and nuclear FISH signals were first quantified using the ImageJ software¹²⁹, and then the cytoplasmic signals were calculated by subtracting nuclear signals from whole-cell total signals. The FISH probes used in this study are provided in Supplementary Data 4.

IF

The IF protocol was adapted from^56,112,130. Cells seeded on a coverslip were fixed in fixative solution for 10 min at room temperature. Fixed cells were then subjected to permeabilization with 0.1% Triton X-100 in TBST buffer at room temperature for 10 min and incubated with primary antibody in the presence of 5% bovine serum albumin (w/v; Beyotime, ST025) at 4 °C overnight. Subsequently, cells were incubated with the fluorophore-conjugated secondary antibody for 2 hr and stained with 1 μg/mL DAPI (Beyotime, C1002) for 5 min at room temperature. Fluorescence signals were detected by using the Leica TCS SP8 system and quantified with ImageJ¹²⁹. For RNase H treatment, cells on coverslips were incubated with RNase H (Beyotime, R7090S, 2.5 U per coverslip) at 50 °C for 40 min before incubation with primary antibody. For RNase R treatment, cells on coverslips were incubated with RNase R (Epicenter Biotechnologies, RNR07250, 5 U per coverslip) at 37 °C for 1 hr before primary antibody incubation. For UV irradiation treatment, cells were exposed to 254 nm UV light at a dose of 100 J/m² for 3 min using a LYUV07-II UV crosslinker. The following antibodies were used in IF: anti-S9.6 (Kerafast, ENH001; 1:200 dilution), anti-γH2A.X (Beyotime, AF1201; 1:200 dilution), anti-FLAG (Beyotime, AF519, 1:200 dilution), Goat Anti-Mouse IgG H&L (Alexa Fluor® 594; 1:200 dilution) (Abcam, ab150080) and Goat Anti-Rabbit IgG H&L (Alexa Fluor® 488; 1:200 dilution) (Abcam, ab150077).

Protein purification and in vitro unwinding assays

His-tagged wild-type Brr2 (WT) and Brr2_ΔH1 mutant plasmids were transformed into E. coli strain Rosetta (Tsingke, DLC204). Protein expression was induced by adding isopropyl β-D-1-thiogalactopyranoside (IPTG) to a final concentration of 0.4 mM, followed by incubation at 37 °C for 14 hr. Cells were harvested and resuspended in lysis buffer (300 mM NaCl, 50 mM NaH₂PO₄·2H₂O and 10 mM imidazole, pH 8.0), followed by sonication and centrifugation. His-fused proteins were purified using NTA-μSphere (pae, P003). The synthetic 5′ biotin-labeled RNA and unlabeled DNA used for preparation of R-loops were purchased from Tsingke. Oligonucleotide sequences are provided in Supplementary Data 4. R-loops were assembled by annealing combinations of labeled RNA and unlabeled DNA oligonucleotides in annealing buffer (10 mM Tris-HCl, pH 7.5, 50 mM NaCl). The mixture was incubated at 95 °C for 10 min and then cooled from 95 °C to 10 °C at a rate of 1 °C/min using a PCR machine. For the in vitro unwinding assay, the purified WT or the ΔH1 mutant was incubated with biotin-labeled R-loops in reaction buffer (10 mM Tris-HCl, pH 7.5, 10 mM MgCl₂, 10% glycerol (v/v), 0.2 μg/μL BSA and 1 mM DTT) for 30 min at 37 °C. 1 μL of 10 mM ATP (Beyotime, D7378) was lastly added to initiate the reaction. The reactions were terminated by adding 1 μL of 1% SDS (w/v) and 1 μL of 10 mg/mL Proteinase K (Beyotime, ST535), followed by incubation at 37 °C for 15 min. Subsequently, 2 μL of loading dye (100 mM Tris-HCl, pH 7.5, 10 mM EDTA, 50% glycerol (v/v), 0.15% Orange G (w/v)) was added. The reaction products were resolved in 10% native polyacrylamide gel by running at 100 V until the dye front reaches the bottom of the gel. Immerse the gel electrophoresis apparatus in an ice bath to maintain low temperature during electrophoresis. After electrophoresis, the oligonucleotides were transferred onto Hybond-N+ membranes (GE Healthcare, RPN303B) and fixed by UV irradiation (254 nm, 0.12 J, 1 min) in a LYUV07-II crosslinker. Non-specific binding was blocked by incubating with blocking buffer (Beyotime, GS009B) for 15 min at room temperature. After incubating with HRP-labeled streptavidin (Beyotime, A0305) at room temperature for 15 min, the membranes were subjected to the ECL protocol using the Bio-Rad ChemiDoc Imaging System. Uncropped and unprocessed scans of gels and blots are provided in the Source Data file.

Comet assay

The comet assay was performed using Comet SCGE assay kit (Enzo, ADI-900-166) as previously described⁵⁶. Cells were washed with ice-cold Ca²⁺/Mg²⁺ -free PBS buffer and mixed with molten LMAgarose (1:10 dilution). Prior to use, the LMAgarose was boiled at 100 °C for 5 min and incubated at 37 °C for 20 min. The mixture of cells and LMAgarose was immediately transferred onto a Comet Slide and placed at 4 °C for 30 min. After immersion in freshly prepared alkaline solution (300 mM NaOH, 1 mM EDTA, pH >13) in the dark at room temperature for 40 min, the slides were washed twice with 1 × TBE buffer (89 mM Tris base, 89 mM boric acid, 2.5 mM EDTA) and then electrophoresed in 1 × TBE buffer for 10 min. Following electrophoresis, the slides were dehydrated in 70% ethanol for 5 min and dried at room temperature. The dried slides were then incubated with 100 μL of CYGREEN Dye (dissolved in deionized water; 1:1000 dilution) at room temperature for 30 min and dried at 37 °C in the dark. Images were captured using the Leica TCS SP8 system. The comet tail length was quantified using the CASP software (http://casplab.com/)¹³¹.

EdU assay

EdU assay was carried out according to the manufacturer’s protocol for the BeyoClick™ EdU Cell Proliferation Kit with Alexa Fluor 555 (Beyotime, C0075S). In brief, cells were exposed to 10 μM EdU for 4 hr at 25 °C (for S2 cells) or 2 hr at 37 °C (for MCF7 cells). Next, the cells were seeded on a coverslip and fixed in 4% paraformaldehyde for 15 min. Cells were then permeabilized with 0.3% Triton X-100 in PBS buffer for 15 min at room temperature. Subsequently, cells were incubated with 50 μL of Click Additive Solution (Click reaction buffer:CuSO₄:Azide 555:Click Additive Solution = 430:20:1:50) for 30 min in the dark and stained with Hoechst 33342 for 10 min. Finally, the EdU-stained cells were visualized and counted using the Leica TCS SP8 system.

Flow cytometry analysis of the cell cycle

Cells were collected after the indicated treatments and then fixed in 70% ethanol overnight at −20 °C. After incubation with propidium iodide (PI) solution (Beyotime, C1052) containing RNase A for 30 min at 37 °C, the cells were analyzed using the BD Accuri C6 flow cytometer (BD Biosciences) and ModFit LT 4.1 software (Verity Software House, USA).

Cell proliferation analysis

Cells were stained with trypan blue (Biosharp, BS924) and counted using a Bright-Line™ Hemacytometer (Sigma, Z359629) after the indicated treatments.

Statistical analyses

Statistical tests for each analysis are indicated in the legend of the figures. Differences were considered to be statistically significant: P < 0.05. Significance: *P < 0.05; **P < 0.01; N.S., no significance.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

The high-throughput sequencing data used in this study have been deposited in the Gene Expression Omnibus (GEO) database under accession number GSE279102 and the BioProject database under accession number PRJNA1039137 (https://www.ncbi.nlm.nih.gov/bioproject/1039137). Source data are provided with this paper.

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Acknowledgements

We thank Dr. Jeremy E. Wilusz (University of Pennsylvania and Baylor College of Medicine), Dr. Qingfa Wu (University of Science and Technology of China), Dr. Fei Xia (Chongqing University) and Mr. Xingze Huang (Chongqing University) for providing useful expression vectors, cell lines or constructive comments. We also thank Dr. Qin Deng at Analytical and Testing Center of Chongqing University for the assistance with confocal microscopy analyses. This work was financially supported by National Natural Science Foundation of China (32270601, 32070633 and 32570667 to C.H.; 32200442 to X.C.), Ministry of Education of the People’s Republic of China (Q2024230 to C.H.), Natural Science Foundation of Chongqing, China (CSTB2025NSCQ-JQX0023, CSTB2024NSCQ-MSX0710 and cstc2022ycjh-bgzxm0140 to C.H.; CSTB2024NSCQ-KJFZMSX0087 to X.C.), Outstanding Young Medical Talents Program of Chongqing (YXQN202401 to X.C.), Fundamental Research Funds for the Central Universities (2024IAIS-QN006 to C.H.) and China Association for Science and Technology (CAST) Young Talent Support Program (156-O-180-0000675-3 to J.L. and 156-O-180-0000673-7 to Z.S.).

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Xiaolan Chen
Present address: Department of Obstetrics and Gynecology, Chongqing Health Center for Women and Children, Chongqing, China
Xiaolan Chen
Present address: Department of Obstetrics and Gynecology, Women and Children’s Hospital of Chongqing Medical University, Chongqing, China
These authors contributed equally: Xiaolan Chen, Jin You.

Authors and Affiliations

School of Life Sciences, Chongqing University, Chongqing, China
Xiaolan Chen, Jin You, Qin Ma, Jiamei Lin, Yu Ji, Shi Li, Zhenxing Song, Rui Su & Chuan Huang
School of Basic Medical Sciences, Division of Life Science and Medicine, University of Science and Technology of China, Hefei, China
Ge Shan

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Xiaolan Chen
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Jin You
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Qin Ma
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Jiamei Lin
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Yu Ji
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Shi Li
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Zhenxing Song
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Ge Shan
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Chuan Huang
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Contributions

C.H. conceived this project, supervised its execution and provide the major funding. X.C., J.Y., Q.M., J.L., Y.J., S.L., Z.S., R.S. and G.S. performed experiments, analyzed data or provided the experimental material. X.C., J.Y. and C.H. wrote the manuscript with input from the other co-authors.

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Correspondence to Chuan Huang.

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Nature Communications thanks Aldo Bader, Hui Shen and the other, anonymous, reviewer(s) for their contribution to the peer review of this work. A peer review file is available.

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Chen, X., You, J., Ma, Q. et al. Dual roles of DEAD-box RNA helicase Brr2 in genome stability regulation. Nat Commun 16, 9133 (2025). https://doi.org/10.1038/s41467-025-64174-8

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Received: 31 October 2024
Accepted: 05 September 2025
Published: 15 October 2025
DOI: https://doi.org/10.1038/s41467-025-64174-8