Walking a tightrope: the complex balancing act of R-loops in genome stability

Joshua R Brickner; Jada L Garzon; Karlene A Cimprich

doi:10.1016/j.molcel.2022.04.014

. Author manuscript; available in PMC: 2023 Jun 16.

Published in final edited form as: Mol Cell. 2022 May 3;82(12):2267–2297. doi: 10.1016/j.molcel.2022.04.014

Walking a tightrope: the complex balancing act of R-loops in genome stability

Joshua R Brickner ¹, Jada L Garzon ¹, Karlene A Cimprich ^1,²

PMCID: PMC9233011 NIHMSID: NIHMS1798025 PMID: 35508167

Summary

Although transcription is an essential cellular process, it is paradoxically also a well-recognized cause of genomic instability. R-loops, non-B DNA structures formed when nascent RNA hybridizes to DNA to displace the non-template strand as ssDNA, are partially responsible for this instability. Yet recent work has begun to elucidate regulatory roles for R-loops in maintaining the genome. In this review, we discuss the cellular contexts in which R-loops contribute to genomic instability, particularly during DNA replication and double-strand break (DSB) repair. We also summarize the evidence that R-loops participate as an intermediate during repair and may influence pathway choice to preserve genomic integrity. Finally, we discuss the immunogenic potential of R-loops and highlight their links to disease should they become pathogenic.

While R-loops are a well-recognized source of genomic instability, replication stress, and DNA damage, there is a growing appreciation for how they contribute to genome maintenance. Here, Brickner et. al. navigate this dichotomy and discuss the consequences of R-loop dysregulation, highlighting links to the immune response and disease progression.

Introduction

How cells respond to the host of endogenous and exogenous insults that threaten to damage their genomes is particularly important to preserve genome stability. Loss of genome integrity can lead to a host of complications, from cancers to neurological disorders (Aguilera and Gómez-González, 2008). Paradoxically, transcription, a fundamental cellular process, can contribute to genomic instability. Although transcription-associated instability can result from many mechanisms, including increased DNA accessibility and interference between replication and transcription, the deregulated formation of non-B DNA structures known as R-loops has emerged as a potent source of instability (Crossley et al., 2019; García-Muse and Aguilera, 2019). R-loops are thought to occur when a nascently transcribed RNA strand invades duplex DNA behind RNA polymerases and hybridizes to the template DNA, forming an RNA-DNA hybrid (hybrid) (Figure 1). Consequently, the non-template DNA is displaced as a strand of single-stranded DNA (ssDNA). While R-loops are generally thought to form behind the polymerase, some evidence suggests that they can form anterior to RNA Polymerase II (RNAPII) during polymerase backtracking (Zatreanu et al., 2019). Hybrids can also form without displacing ssDNA, for example during Okazaki fragment synthesis. Importantly, most techniques to detect so-called R-loops, outside of bisulfite-based sequencing techniques, probe for the hybrid itself and thus cannot readily distinguish between a bona fide R-loop or a hybrid (Chédin et al., 2021). Hence in some scenarios, it is not clear what structure is actually formed. In this review, we will discuss the roles of both hybrids alone as well as R-loops, referring to the specific structure when it is clear which is known to form.

The use of imaging and sequencing-based techniques has allowed researchers to quantify R-loop levels and characterize their genomic occupancy in cells (Chédin et al., 2021; Crossley et al., 2020, 2021; Paull, 2019). These studies have shown that hybrid formation is widespread, with hybrids occupying 5–10% of the genome, and single-molecule experiments suggest they vary in length from under 60 base pairs to several kilobases. Initial attempts to quantify hybrids in unperturbed HeLa cells have put a lower estimate on their abundance at ~300 R-loops/cell, with an average half-life of 11 minutes. This suggests that cells resolve on the order of 27,000 R-loops per day (Crossley et al., 2020; Sanz et al., 2016). As products of transcription, R-loops are frequently genic in nature, although they have been found at centromeres, telomeres and other repetitive genomic regions, as well as enhancers (Liu et al., 2021a; Niehrs and Luke, 2020; Tan and Lan, 2020). They also occupy different genomic regions depending upon growth conditions or cell type. Furthermore, they tend to form in regions of high GC content and GC-skew, and the displaced ssDNA is prone to form secondary structures such as G-quadruplexes (G4s) (Castillo-Guzman and Chédin, 2021).

As dynamic structures, R-loops can be controlled at many levels, including in their formation and resolution (Figure 1). Multiple factors and pathways are involved in this regulation (reviewed in Bayona-Feliu and Aguilera, 2021; Crossley et al., 2019; García-Muse and Aguilera, 2019). These include RNA processing pathways and export machinery, which suppress hybrid formation by sequestering nascent RNA from the template, promoting chromatin compaction, and packaging and removing RNA from the nucleus. Topoisomerases also suppress hybrid formation by minimizing supercoiling. Modification of the RNA strand of the hybrid by the RNA methyltransferases METTL3 or TRDMT1 helps regulate R-loop formation (Abakir et al., 2020; Chen et al., 2020; Zhang et al., 2020). Once formed, a host of additional proteins resolve or degrade R-loops. Two nucleases dedicated to resolving R-loops in mammalian cells, RNase H1 and RNase H2, specifically degrade the RNA moiety of an RNA-DNA hybrid. RNase H2 also acts in ribonucleotide excision repair, removing single mis-incorporated ribonucleotides from DNA (Hyjek et al., 2019). Numerous helicases can also unwind R-loops and hybrids once formed, and several proteins that are involved in separate processes, such as chromatin remodelers and DNA repair nucleases, have been implicated in their resolution (García-Muse and Aguilera, 2019; Paull, 2019; Uruci et al., 2021). The many factors implicated in R-loop formation and resolution suggest the cell needs to dynamically balance these processes.

R-loops have both beneficial and harmful impacts on the cell. They are important intermediates in immunoglobulin class-switch recombination in B-cells and in mitochondrial replication (Yu et al., 2003). They have also been implicated in regulating gene expression, where they can prevent DNA methylation of promoter CpG islands and subsequent gene silencing, regulate transcriptional termination, and regulate the recruitment of chromatin remodelers (Bayona-Feliu and Aguilera, 2021; Crossley et al., 2019). Recent work has highlighted the importance of R-loops in maintaining telomeric and centromeric integrity (Liu et al., 2021a; Niehrs and Luke, 2020), and also suggests that R-loops or hybrids formed at DNA double-strand breaks (DSB) help promote DNA repair (Marnef and Legube, 2021). By contrast, R-loops can act as a source of DNA damage and genomic instability. As first shown upon perturbation of genes involved in RNA biogenesis and export, R-loop accumulation can dramatically increase DNA damage, specifically DSBs (Huertas and Aguilera, 2003; Li and Manley, 2005; Paulsen et al., 2009). Furthermore, overexpression of RNase H1 in cells can rescue R-loop induced damage in many scenarios. Precisely how R-loops increase DNA damage and genome instability is still a matter of intense investigation, and different mechanisms may be at play depending on the context in which the R-loop or hybrid forms.

In this review, we will discuss the mechanisms by which R-loops may either promote or prevent genome maintenance and stability, focusing primarily on eukaryotic processes. Excellent reviews on their physiological roles have been published, and we refer the reader to these and others on the factors that regulate R-loop formation (Bayona-Feliu and Aguilera, 2021; Crossley et al., 2019; García-Muse and Aguilera, 2019; Niehrs and Luke, 2020; Zong et al., 2020). Here, we will focus on how R-loops cause genomic instability, emphasizing their impact on DNA replication. Additionally, we will highlight advances in how R-loops are thought to promote DSB repair. Finally, we will discuss the pathological consequences of R-loop dysregulation.

R-Loops and the Replication Stress Response

During S-phase, replication forks may encounter R-loops, leading to replication stress and DNA damage. Although mechanisms exist to spatially and temporally separate transcription and replication, conflicts between these processes may occur at some frequency since they take place on the same DNA template (Hamperl and Cimprich, 2016; Lalonde et al., 2021). Indeed, numerous studies demonstrating the impact of R-loops on replication stress responses support the idea that R-loops are a source of replication stress and genome instability (reviewed in Crossley et al., 2019; García-Muse and Aguilera, 2019; Lalonde et al., 2021; Rinaldi et al., 2020). Perturbations that alter R-loop processing genes generally cause the most DNA damage in S-phase cells, suggesting that the R-loop itself affects fork progression. Early studies also demonstrated that replication forks slow at sites corresponding to an induced R-loop and that RNase H1 overexpression can restore normal fork progression in cells with deregulated R-loop formation (Gan et al., 2011; Gómez-González et al., 2011; Wellinger et al., 2006). Extending this further are observations showing that RNase H1 expression reverses fork slowing and/or fork asymmetry caused by hormone stimulation, bacterial infection, oncogene-induced replication stress, and Top1 mutations, all of which can induce R-loop accumulation (Bauer et al., 2020; Kotsantis et al., 2016; Stork et al., 2016; Tuduri et al., 2009). In recent years, much emphasis has focused on understanding the molecular mechanism by which these structures induce a block and how these barriers can be overcome.

R-loop-induced fork stalling: the nature of the block

One important question concerns the impact of replication fork orientation relative to an R-loop. Forks can encounter the transcription machinery in a head-on (HO) or co-directional (CD) orientation, leading to distinct outcomes (Figure 2A). In mammalian cells, transcription induction on an episome containing an R-loop forming region exacerbated the damage associated with both HO and CD transcription-replication conflicts (TRCs) as compared to an episome with transcription but no R-loop induction (Hamperl et al., 2017). This suggests the R-loop or R-loop-stalled transcription machinery is more of a problem than transcription itself. Moreover, although both HO and CD encounters led to DNA damage, particularly when transcription was strongly induced, HO conflicts were more detrimental. Different damage signaling pathways were also induced: HO encounters activated ATR rather than ATM, which was instead induced by CD conflicts (Figure 2A). ATM activation is likely a result of DSB formation at R-loops, but how ATR is activated is unclear. It could involve recognition of ssDNA arising at the stalled fork, on the R-loop itself, or from DSB formation and resection. As RPA binds to the ssDNA in an R-loop and recruits RNase H1 (Nguyen et al., 2017b), one possibility is that it could also recruit the ATR-ATRIP complex, contributing to ATR activation. In cells in which splicing was inhibited or splicing factors were downregulated, ATR was activated as opposed to ATM (Chen et al., 2018; Matos et al., 2020; Nguyen et al., 2018). This could indicate that fork stalling induced by HO encounters is the more problematic issue in human cells. Importantly, the activation of these kinases, in particular ATR, has been shown to prevent genome instability through multiple mechanisms described further herein.

Figure 2. — Factors Contributing to Replication Fork Stalling at R-loops. (A) Illustrates the two orientations in which a replication fork can encounter a R-loop, leading to a TRC. Head-on conflicts (HO, left) activate ATR and codirectional conflicts (CD, right) activate ATM. HO conflicts with R-loops are associated with greater genome instability. (B) Illustrates potential mechanisms that cause fork slowing in the HO (left) and CD (right) orientation. Flags represent epigenetic markers that promote chromatin compaction.

The molecular events that occur when a fork meets the transcription machinery in the HO and CD scenarios are distinct, providing some insight into the different cellular responses. In the HO orientation, the CMG helicase, which translocates on the leading strand in eukaryotes, approaches an R-loop from the opposite strand that is engaged by the transcription machinery (Figure 2B). In fact, the orientation of this approach has led some to envision that R-loops may not pose a problem for the fork. However, RNA polymerase normally engages both DNA strands, making this protein complex a likely block to CMG and/or the replicative polymerase (Barnes et al., 2015). Converging transcription and replication machineries could also lead to topological stress in the HO orientation, preventing further progression of each process. Both replication and transcription cause positive supercoiling to accumulate ahead of the respective machineries and negative supercoiling to accumulate behind them. When converging, the cumulative topological stress between the machineries may become too large to be compensated by topoisomerase activity, causing the fork to stall (Figure 2B). Furthermore, the negative supercoiling that builds up behind the replisome could promote hybrid formation in this orientation. Indeed, HO conflicts seem to enhance hybrid formation (Hamperl et al., 2017). Work in B. subtilis supports this idea and also provides insight into the mechanism by which stalling occurs. Here surprisingly, the effects of a HO vs CD encounter are similar to eukaryotic organisms, despite movement of the replicative helicase on the lagging strand (Lang et al., 2017). HO encounters induced by RNaseH loss and R-loop stabilization led to cell death and a complete failure of DNA replication in this system. Furthermore, it is topoisomerase II-dependent resolution of the positive supercoiling that promotes R-loop formation by allowing diffusion of negative supercoils behind RNAPII (Lang and Merrikh, 2021). By contrast, upon CD conflicts topological stress would be relieved. Nevertheless, in certain contexts CD encounters can be problematic. RNA polymerase may be stabilized by backtracking and pose a stronger block to fork progression (Figure 2B). Consistent with this idea, RECQL5, a helicase that associates with RNAPI and RNAPII, suppresses backtracking to reduce the incidence of TRCs and genome instability (Saponaro et al., 2014; Urban et al., 2016). Similarly, increased backtracking leads to R-loop dependent DNA damage in bacteria (Dutta et al., 2011).

There are scenarios in which an R-loop can contribute to damage regardless of orientation. For instance, RNAPII accumulation at a stalled or backtracked polymerase could exacerbate both CD and HO conflicts (Figure 2B). Additional factors may also contribute to orientation-independent damage. Studies in yeast and mammalian cells suggest that chromatin condensation contributes to the ability of R-loops to cause DNA damage. R-loop accumulation is associated with histone modifications that promote chromatin compaction, including H3S10 phosphorylation (H3S10P) and H3K9 dimethylation. Moreover, histone H3S10 phospho-mutants do not accumulate DNA damage, despite increased R-loop formation (Castellano-Pozo et al., 2013; García-Pichardo et al., 2017). One possibility is that chromatin condensation driven by these modifications contributes to R-loop-associated fork stalling (Figure 2B). Of note, R-loops themselves are not thought to be bound by nucleosomes, so these modifications are likely to be either adjacent to the R-loop, as part of a larger chromatin domain or be temporally distinct (Dunn and Griffith, 1980). By contrast, work in mammalian cells showed that H3S10P can also be protective. In this scenario, the oncogene MYCN activates Aurora A to phosphorylate H3S10, ultimately suppressing R-loop formation and TRCs (Roeschert et al., 2021). While these results seem discrepant, how R-loops promote or are regulated by chromatin condensation may be context specific, and Aurora A-mediated H3S10P may be limited to MYCN-driven genes, where avoiding damage is important to support tumorigenesis.

Another question is whether it is the R-loop alone or the stalled RNA polymerase together with the R-loop that challenges replication fork progression. As there are mechanisms for removal of stalled RNAP in the cell, both scenarios should be considered. Recent observations bring new insight to this question. Using a reconstituted Saccharomyces cerevisiae (S. cerevisiae, budding yeast)-based replication system, replication forks have been challenged by deproteinated or “naked” R-loops lacking RNA polymerase and containing a G4 on the non-template strand (Kumar et al., 2021). In this context, leading strand replication forks pause at both HO and CD R-loops but can bypass these structures and restart DNA replication on the leading strand downstream of the R-loop. Using RNase H1 to probe the hybrid’s role in these pauses, it was shown that leading strand hybrids (CD conflict) directly impede fork progression (Figure 2B). Surprisingly, however, fork stalling was still observed following RNase H1 expression when the hybrid was on the lagging strand (HO conflict). This observation, taken together with subsequent studies, suggests that in the HO orientation, the G4 formed on the leading strand as a result of R-loop formation on the lagging strand is the true cause of the fork block (Figure 2B). Hence, G4 formation in the displaced ssDNA of an R-loop may be an important determinant of an R-loop’s impact on the replication fork. Studies using a bacterial replication system are generally consistent with these findings, supporting the idea that naked R-loops can be bypassed in both orientations. They also demonstrate that a RNAP-bound R-loop poses a greater threat to fork progression than a naked R-loop, particularly in the HO orientation (Brüning and Marians, 2020). Although biochemical studies with a RNAP-bound R-loop have not yet been done in higher eukaryotes, a dCas9-associated R-loop also blocks the fork in both orientations using the reconstituted S. cerevisiae-based replication system (Schauer et al., 2020).

Mechanisms for restart

In principle, there are several ways for replication to continue when a fork encounters an R-loop. These include bypass of the R-loop and replication restart, removal of the block, either through R-loop resolution or removal of RNAPII, completion of DNA replication through use of dormant origins, or alternatively various TRC avoidance mechanisms, all of which are discussed below.

i. Fork bypass of R-loops and replication restart

It seems likely that cells require some sort of R-loop bypass mechanism to complete the replication of long genes, which can take more than one cell cycle to transcribe. Biochemical studies using the budding yeast system described above provide some basic insight into how this bypass or fork restart may occur (Kumar et al., 2021). In the CD orientation, the CMG/replisome can translocate over a naked RNA-DNA hybrid if the 5’-RNA end is flush, and if a free 3’-OH is present on the RNA strand, DNA polymerase α can extend it, allowing fork progression (Figure 3A). By contrast, if the 5’-end of the RNA is not annealed to the DNA template, CMG can unwind the flap-containing structure (Figure 3B). How the presence of RNAPII and other transcriptional regulators would affect the outcome is unclear. When RNA polymerase is present, CMG translocation along the hybrid duplex would bring it in closer proximity to the stalled RNAPII-hybrid complex without resolving the hybrid. In this case, restart would require either bypass and later dissolution of this complex, or alternatively, RNAPII removal and RNA extension using Pol α or another polymerase (Figure 3C). Alternatively, if CMG unwinds the hybrid containing the free 5’-end, it would effectively resolve that hybrid, freeing RNA polymerase to continue transcription (Figure 3D). As these machines move at similar speeds in eukaryotic cells, RNA polymerase could continue with the helicase behind it (Conti et al., 2007; Muniz et al., 2021). Whether this can occur in the genome is unclear but it is consistent with observations that R-loop levels are decreased in S-phase when the fork meets a CD R-loop in an episomal system (Hamperl et al., 2017).

Figure 3. — Models for the Restart of Replication Forks Stalled at R-loops. (A-D) illustrate potential bypass mechanisms in the CD orientation. (A) The CMG helicase translocates over and bypasses a “naked” hybrid when the 5’-end of the RNA strand is annealed to the DNA. Once the hybrid is bypassed, the RNA strand can be extended by Pol α. (B) The CMG helicase unwinds a hybrid when the 5’-end of the RNA strand is exposed as a flap, allowing fork progression to continue. (C) CMG translocation over the hybrid would lead to its arrest at RNAP. Fork progression at this point would require CMG to bypass the stalled RNAP or for RNAP to be removed. (D) CMG unwinding of a hybrid bound to RNAP would allow RNAP to continue and the replication fork to progress behind transcription. (E-G) illustrate potential restart mechanisms in the HO orientation. (E) CMG bypass of RNAP would leave ssDNA exposed, allowing PrimPol recruitment. Primer synthesis by PrimPol allows the fork to continue, leaving behind ssDNA that can be replicated by a gap-filling mechanism once transcription is complete. (F) ssDNA resulting from bypass and repriming or a failure to reprime, and which persists behind the replication fork, could serve as a template for the formation of hybrids. Hybrids could form through *de novo* synthesis by RNAP (top) or through the association of nascent RNA resulting from transcription after the fork has passed (bottom). (G) Restart of a stalled fork initiated by SLX4 and MUS81-mediated fork cleavage is thought to relieve the torsional stress, resulting in resolution of the R-loop formed during a HO conflict. Religation of the fork by XRCC4/LIG4 and ELL-mediated restart of transcription allows both replication and transcription to continue. Prior to fork cleavage, RAD51 may promote fork reversal. RECQ1 is needed to reset the fork and RECQ5 promotes removal of RAD51.

In the HO orientation, a fork can progress past a naked R-loop on the lagging strand as long as no secondary structure is present on the leading strand to impede unwinding or polymerase progression. If such a structure is present, the fork can restart downstream of the impediment, generating a gap. However, when RNAPII is present, restart is more complex. In bacteria, a full halt to DNA replication is observed in the HO orientation, and in mammalian cells HO conflicts lead to ATR activation, which is indicative of fork stalling. Since RNAPII interacts with both strands of duplex DNA, some alternative restart mechanism would need to be employed. If CMG could bypass the RNAPII block as it can bypass protein-DNA crosslinks (Huang et al., 2013; Sparks et al., 2019), restart might happen using PrimPol, an enzyme with both primase and polymerase activities (Figure 3E). Indeed, PrimPol suppresses extensive R-loop formation at GAA repeats, where R-loops can promote formation of a triplex structure (Šviković et al., 2019). PrimPol may mediate repriming in the wake of the uncoupled replicative helicase, suppressing ssDNA formation and hybrid formation on that ssDNA. Consistent with this idea, two factors which minimize ssDNA formation by promoting gap filling during post-replicative repair, RAD18 and UBE2B, also suppress hybrid formation in a post-replicative fashion (Figure 3E) (Barroso et al., 2019). These observations suggest that ssDNA forming behind the fork as a result of gap formation and/or gap resection may be a template for hybrid formation. In this context, RNAs transcribed on the other strand could bind to form a hybrid, or RNAPII could be recruited to the DNA for de novo RNA synthesis and hybrid formation (Figure 3F) (Barroso et al., 2019; Delamarre et al., 2020; Kemiha et al., 2021). As RNAPII can initiate transcription on ssDNA in the absence of a promoter sequence in vitro and has been shown to act on the ssDNA of an R-loop to initiate antisense transcription (Kadesch and Chamberlin, 1982; Tan-Wong et al., 2019), it could bind DNA gaps to synthesize RNA behind the fork.

The nuclease MUS81 also plays a role in fork restart at R-loops, initiating a fork cleavage and religation cycle that restores both transcription and replication (Figure 3G) (Chappidi et al., 2020). SLX4, MUS81 and its partner protein EME1 are proposed to cleave the fork, relieving torsional stress and resolving the R-loop generated by a HO conflict. Following ELL-mediated transcriptional restart and LIG4/XRCC4-mediated fork religation, replication restarts, allowing replication and transcription machineries to traverse each other. MUS81-EME1 acts with RAD52 and POLD3 in this process to promote fork restart at R-loops. These factors are proposed to mediate parental DNA strand annealing and DNA synthesis respectively, promoting a form of semi-conservative DNA replication. Prior to initiating this restart pathway, RAD51-mediated replication fork reversal may also occur (Figure 3G) to stabilize the stalled fork (Chappidi et al., 2020). When this occurs, RECQ1 and RECQ5 are required to reset the reversed fork and remove RAD51 at the R-loop stalled fork, respectively, such that cleavage and restart can occur.

ii. Resolution of the R-loop or transcription block

Fork bypass of the R-loop is one mechanism for allowing both continued replication and transcription, but RNA polymerase removal, transcription termination and auxiliary helicases can also facilitate fork progression. While these events could occur in all cell cycle phases, there are instances in which they are coupled to replication fork progression. In budding yeast, RNAP mutants with increased chromatin binding capacity exhibit increased TRC-associated DNA damage and genomic instability, suggesting the persistence of RNAP on chromatin is problematic for the fork (Felipe-Abrio et al., 2015). Mec1, the budding yeast ortholog of ATR, regulates the chromatin remodeler Ino80 and the transcription complex PAF1 to remove chromatin-bound RNAPII at certain transcribed genes upon HU-induced replication stress (Figure 4A) (Poli et al., 2016). This promotes fork progression and restart, consistent with the idea that RNAPII removal helps to resolve TRCs. Yeast RNAPIII may also be regulated by Mec1 to limit hybrid accumulation and control its chromatin binding (Hurst et al., 2021).

Figure 4. — Potential Mechanisms for Resolution of R-loop Induced Transcription-Replication Conflicts. (A) In budding yeast, RNAP removal and proteasomal degradation can be initiated by activation of ATR and mediated by Paf1c and Ino80. RNAP phosphorylation also promotes its degradation, which is counteracted by the protein phosphatase PP1 and its regulatory subunit PNUTS and WDR82. (B) ATR activation at stalled forks is dependent upon MUS81-EME2-mediated fork cleavage, which is needed to generate ssDNA. Activation prevents additional and excessive fork cleavage by MUS81 and suppresses TRC formation. Other characterized functions of ATR may also promote fork recovery. (C) A fork stalled at an HO conflict can be rescued by a fork approaching the R-loop from the CD orientation. (D) Fork pausing at the 3’-end of a gene (highlighted in gray) located near an origin allows time for transcription to complete before the fork progresses into the gene. The converging fork and transcription lead to torsional stress that is relieved by TOP1. Pausing also activates ATR, which may reinforce the slowdown by inhibiting elongation. (E) Histone deacetylation suppresses R-loop formation in diverse ways, preventing genome instability. (F) SWI/SNF complexes including the subcomplex containing ARID1A act in a pathway with FANCD2 and SETX to suppress R-loop formation and TRCs. ARID1A recruits TOP2A to forks to relieve torsional stress.

Mechanisms for RNAPII removal after transcription stress are also known in higher eukaryotes. For example, one mechanism for RNAPII removal involves its ubiquitination and degradation, which can occur following UV damage or other types of transcription blocking lesions (reviewed in Noe Gonzalez et al., 2021). RNAPII phosphorylation at S5 was shown to prevent its degradation and turnover. Depletion of factors that promote S5 dephosphorylation, including the protein phosphatase PP1 and its regulatory subunits PNUTS and WDR82, increase RNAPII stability on chromatin and promote TRCs (Figure 4A) (Landsverk et al., 2020). Furthermore, INO80 can promote R-loop turnover in mammalian cells when it is tethered to an R-loop site, and its knockdown increases R-loop formation (Prendergast et al., 2020). Whether these pathways help restart forks stalled at R-loop associated TRCs is still unclear. It is also unclear if ATR regulates any of these pathways in a manner analogous to what occurs in yeast, although ATR-dependent repression of transcription in the vicinity of stalled forks, which could suppress TRCs and R-loop formation, has been reported (Im et al., 2014).

Although ATR’s role in controlling transcription is poorly understood, it has better known roles in fork stabilization and restart that may be important at R-loop stalled forks. Interestingly, MUS81 may cooperate with ATR in a negative feedback loop that protects cells from R-loop induced damage (Matos et al., 2020). MUS81, acting with its partner protein EME2, generates ssDNA at stalled forks through fork cleavage, activating ATR (Figure 4B). This activation suppresses additional R-loop formation and prevents fork collapse mediated by the MUS81-EME1 complex. As ATR inhibitor treatment led to more DSB formation when R-loops accumulate, as well as a defect in fork restart and increased TRCs, it seems likely that ATR activation serves multiple roles in this context. Though some may be related to ATR’s suppression of origin firing and arrest of cell cycle progression, ATR may also suppress the activity of other enzymes, including the MUS81-EME1 complex. Alternatively, or in parallel, it could suppress the restart of forks cleaved by MUS81-EME1. Clearly further work is required to sort out this complex pathway and elucidate how MUS81-EME1 can have both beneficial (as in the fork cleavage and religation model) and toxic roles.

Transcription termination is another mechanism for RNAPII removal, and in fact, termination defects are associated with R-loop formation and genome instability (Hatchi et al., 2015; Morales et al., 2016; Skourti-Stathaki et al., 2011). Numerous mechanisms for premature termination following transcription stress have been described that involve RNA cleavage and RNAPII disengagement (reviewed in Noe Gonzalez et al., 2021). In these cases, specialized helicases and nucleases, such as SETX (Hatchi et al., 2015; Skourti-Stathaki et al., 2011), DHX9 (Cristini et al., 2018), DDX5 (Mersaoui et al., 2019), and XRN2 may play important roles (Skourti-Stathaki et al., 2011).

MYCN, an oncogenic transcription factor often amplified in neuroblastomas, may have an analogous function, acting as a transcription terminator to resolve replication blocks. In MYCN-driven cells, MYCN acts via a two-tiered mechanism to facilitate RNAPII removal. First, MYCN recruits the exosome to stalled and backtracked RNAPII to promote transcription termination and to prevent fork stalling and DSB formation (Papadopoulos et al., 2022). Should breaks occur, MYCN indirectly recruits BRCA1 to promoter-proximal pause sites, which further recruits mRNA decapping complexes to suppress R-loop formation at these regions. Although this occurs in a cell cycle independent manner, it could also limit TRCs by suppressing the accumulation of stalled RNAPII (Herold et al., 2019; Papadopoulos et al., 2022).

Finally, SETX and other auxiliary helicases and translocases may help remove RNAP and/or the R-loop outside the context of termination and play a more direct role in the resolution of TRCs (Figure 1). SETX is associated with stalled replication forks in mammalian cells and in budding yeast, and its loss leads to increased R-loop formation and genome instability (Alzu et al., 2012; Costantino and Koshland, 2018; Skourti-Stathaki et al., 2011; Yüce and West, 2013). The FANCM helicase/translocase, or its yeast homolog Mph1, also plays a role at TRCs to suppress R-loop formation. FANCM’s ability to resolve R-loops in vitro helps to release RNAP from transcriptional arrest or promote its removal from chromatin (Lafuente-Barquero et al., 2017; Schwab et al., 2015). Furthermore, FANCM may be important to resolve TRCs at telomeres in ALT cells by disrupting TERRA R-loops (Pan et al., 2019). RTEL1 can also act in a pathway with the polymerase δ-associated factor Poldip3 to resolve R-loops formed at TRCs. This may be particularly important at common fragile sites and other loci that are difficult to replicate or which may form G4s (Björkman et al., 2020; Kotsantis et al., 2020; Wu et al., 2020). Similarly, the Pif1 helicase, which translocates on the strand opposite the CMG helicase, can assist in removing R-loops bound by Cas9 in both orientations, raising the possibility it could help remove RNAPII at forks (Schauer et al., 2020). A host of other helicases and translocases that can resolve R-loops have been identified that may also play a role in this process, including AQR, BLM, WRN, SMARCAL1, DDX1, DDX5, DDX19, DDX21, DDX23, DHX9, and UAP56 (Figure 1) (Chakraborty and Grosse, 2011; Chang et al., 2017a; Cristini et al., 2018; Hodroj et al., 2017; Kim et al., 2020; Marabitti et al., 2019, 2020; Matson and Zou, 2020; Mersaoui et al., 2019; Pérez-Calero et al., 2020; Pugliese et al., 2019; Sollier et al., 2014; Song et al., 2017; Sridhara et al., 2017; Tan et al., 2020a). Interestingly, the PCNA unloader ATAD5 may regulate the action of several of these helicases, helping recruit them to the replication fork to suppress R-loops and TRCs (Kim et al., 2020). Why the activity of so many helicases is required and how their activity is coordinated remain open questions.

iii. Origin firing and genome organization

Another mechanism for the rescue of forks stalled at R-loops is related to eukaryotic genome organization and the bidirectional nature of origin firing. In prokaryotes, most genes are replicated in a codirectional fashion. Those oriented for replication in a head-on fashion are generally stress responsive genes for which there is pressure to select for beneficial adaptive mutations that promote stress tolerance or increased virulence (Lang et al., 2017). Similarly, origins are often located upstream of the promoter regions of highly expressed genes in higher eukaryotes. Hence, replication is often codirectional with transcription, an arrangement that minimizes the problems associated with TRCs (Chen et al., 2019; Petryk et al., 2016). As forks approach a gene from both directions, a fork approaching from the opposite direction may complete replication if R-loop-induced fork stalling occurs (Figure 4C) (Brambati et al., 2018). Many TRCs in the eukaryotic genome could be resolved by the very nature of this genomic organization.

iv. TRC avoidance by fork pausing

Active fork pausing may be a(nother) mechanism to avoid a TRC. Recent work suggests that forks may pause in the HO orientation at the 3’-end of genes containing R-loops when located in close proximity to an origin (Figure 4D) (Promonet et al., 2020). This pausing may be an active process that can prevent TRCs by allowing for the completion of transcription, which has been shown to occur in bursts (Tantale et al., 2016). ATR-dependent RPA phosphorylation is seen specifically at these R-loop-containing transcription termination sites within the genome, and may result from the fork pausing that arises in response to topological stress building up upon convergence of RNA and DNA polymerases (Promonet et al., 2020). While ATR activation could help to slow fork progression, it may also promote RNAPII displacement ahead of the fork in a manner analogous to what has been reported in S. cerevisiae (Poli et al., 2016). Interestingly, H3K4 methylation in yeast, which is abundant in highly transcribed genes, acts in a similar manner to suppress TRCs in checkpoint mutants by slowing the rate of DNA replication (Chong et al., 2020). Like fork pausing, altering fork speed could allow replisomes to avoid the transcription machinery.

v. TRC avoidance by chromatin remodeling

Cells also alter the local chromatin state to avoid R-loop associated TRCs (Bayona-Feliu and Aguilera, 2021). Among the factors involved, several regulate histone post-translational modifications, such as acetylation. Initial observations revealed that mutation of THO, mSin3 and other proteins involved in regulating histone lysine deacetylation leads to R-loop accumulation and R-loop dependent DNA damage (Salas-Armenteros et al., 2017; Wahba et al., 2011). Several other chromatin regulators have also been implicated in suppressing R-loop formation and TRCs (Figure 4E). Whether they resolve the R-loop at a TRC or simply prevent R-loop formation to avoid TRCs is not clear. Among those factors involved are the yeast SIRT6 homologs Hst3 and 4 (Feldman and Peterson, 2019), the mammalian SIRT7 (Song et al., 2017), and BRD4, a bromodomain containing protein that recognizes acetylated lysines (Edwards et al., 2020; Lam et al., 2020). BRD4 may prevent R-loop formation through its interaction with the elongation factor, pTEFb. Changes to histone acetylation status may therefore control R-loop levels by altering chromatin compaction and suppressing transcription, thereby minimizing the ability of the nascent RNA to interact with the DNA template upon TRCs.

Several proteins directly involved in chromatin remodeling, which is important for nucleosome positioning and chromatin relaxation, may also suppress R-loop associated TRCs (Figure 4F). Among these are components of the SWI/SNF complex and its catalytic subunit BRG1, which appears to act with FANCD2 and SETX to suppress R-loop formation and TRCs. BRG1 is found at stalled forks and limits chromatin accessibility at R-loops sites, suggesting it may reduce the ability of nascent RNA to bind to the template DNA (Bayona-Feliu et al., 2021). Members of the cBAF and PBAF SWI/SNF complex subtypes, ARID1A and PBRM1 respectively, may also be involved (Chabanon et al., 2021; Tsai et al., 2021). ARID1A helps to recruit topoisomerase IIa (TOP2A) to chromatin and may suppress R-loop associated TRCs by promoting the resolution of topological stress at the conflicts. Finally, the remodelers INO80 (Poli et al., 2016; Prendergast et al., 2020), and ATRX have been implicated in TRC resolution and/or R-loop suppression (Nguyen et al., 2017a; Yan et al., 2022).

R-loops and fork progression

An emerging question in the field is whether fork slowing at an R-loop is actually problematic for the cell or whether post-replicative hybrid formation is an additional culprit (Kemiha et al., 2021). Although the idea that forks slow upon approaching an R-loop is certainly the dominant model, it is important to better establish whether slowing and fork reversal actually occurs in front of the R-loop. Genetic studies are consistent with the involvement of fork reversal factors, but forks reversed at an R-loop have not yet been observed by electron microscopy. Independent verification by this or another single-molecule method is needed. One intriguing possibility is that fork slowing is alternatively, or additionally, due to post-replicative hybrid formation on the ssDNA that forms upon bypass of the R-loop (Figure 3F).

Recent data raise questions about the idea that R-loops pose a direct block to fork progression. Loss of many R-loop resolution factors leads to fork slowing and asymmetric fork progression as measured by DNA fiber assays, indicating that not all forks stall. Consistent with the idea that R-loops are involved in this slowing, the block is relieved by RNaseH expression (Salas-Armenteros et al., 2017; Tuduri et al., 2009). Surprisingly, however, most forks slow under these conditions, despite the fact that not all will be in the vicinity of an R-loop. This suggests there may be some type of global mechanism for fork slowing. Local slowing at R-loops may activate a checkpoint, which in turn slows global elongation. Evidence for such a global fork response has been reported under other conditions: ATR globally slows fork progression by regulating fork reversal in response to interstrand crosslinks (Mutreja et al., 2018), and checkpoint dependent control of both initiation and elongation have been observed (Bacal et al., 2018; Can et al., 2019; Seiler et al., 2007).

How hybrids formed behind the replication fork would slow DNA replication is not clear. One possibility is that they interfere with homologous recombination-mediated fork restart if not efficiently removed. Alternatively, they could suppress histone deposition, leading to other changes that promote fork slowing. Clearly, further work is needed to investigate this and other possibilities.

Replication-Independent Mechanisms for R-Loop Induced DNA Damage and Genome Stability

Although the impact of R-loops on genome stability is largely related to their effects on DNA replication, R-loops form outside of S-phase and may also threaten genome stability in a replication-independent manner (Hamperl et al., 2017; San Martin-Alonso et al., 2021). One mechanism by which R-loops could cause damage outside of S-phase involves nucleolytic processing. The endonuclease activity of two factors involved in nucleotide excision repair, XPF and XPG, is responsible for the DNA DSBs formed when R-loops are induced, and the absence of XPG increases cellular R-loop levels (Makharashvili et al., 2018; Sollier et al., 2014; Stork et al., 2016). This suggests that these flap-endonucleases process these structures, although the mechanism by which they act is not well understood (Figure 5A). Should XPF and XPG cleave opposite strands of the R-loop, this processing would directly generate a DSB. Conversely, XPF and XPG may cleave the same strand upstream and downstream of the hybrid, respectively, generating a single strand break (SSB). If two R-loops in close proximity are processed in this way, it could produce a DSB outside of S-phase. We note however that some DSBs arising from XPF and XPG-mediated processing may not be S-phase independent, as the SSB could remain until the cell progresses into S-phase, where it leads to DSB formation upon encountering the replication fork. It was shown that the splicing factor XAB2 helps recruit XPG and XPF to R-loops, suggesting that these factors might act outside the context of classical nucleotide excision repair (Goulielmaki et al., 2021).

Figure 5. — Potential Mechanisms of Replication-Independent R-loop-mediated DNA Damage. (A) The TC-NER nucleases XPG and XPF initiate R-loop processing by cleaving the 5’ or 3’-flap of the R-loop-template DNA junction, respectively. Cleavage may occur on either the template strand, excising the hybrid, or on the non-template strand, excising the displaced ssDNA to generate a single-strand break. Conversely, XPG and XPF may cleave opposing DNA strands of the R-loop, generating a DSB either 5’ or 3’ to the R-loop. (B) R-loop processing by several nucleases and TOP1-mediated DNA nicking in response to camptothecin generate two SSBs on opposing DNA strands, which mimic a DSB. (C) The displaced ssDNA of a transcription bubble is more accessible to damage from reactive oxygen species (ROS). ROS-generated SSBs in the displaced ssDNA promote unscheduled R-loops and G4s. (D) AID and APOBEC proteins can convert cytosine to uracil in the displaced ssDNA. Downstream processing of uracil results in an abasic site, which can cause SSBs. Additionally, the RNA editor ADAR has some activity towards the DNA strand of the hybrid. Base modifications are represented by stars.

R-loops also contribute to DSB formation in quiescent cells treated with camptothecin (CPT), a topoisomerase I (TOP1) inhibitor. R-loops induced by CPT contain a trapped TOP1 cleavage complex. Proteolytic removal of this complex leads to a SSB on one strand, while R-loop processing on the other strand generates a second SSB (Figure 5B). In this case, the endonucleases XPF, XPG, FEN1 and MRE11 all contribute to break formation (Cristini et al., 2019). Similarly, the flap endonuclease activity of the Sae2/CtIP protein, normally involved in DSB resection, can suppress R-loop and DNA break formation in both untreated and CPT-treated human cells (Makharashvili et al., 2018). In all cases, whether nucleolytic processing is required for repair/removal of the R-loop or whether the nucleases are acting in a gratuitous and pathological fashion simply because the structure is accessible for processing is not clear. The accumulation of R-loops in the absence of CtIP and XPG suggests that this may be a step in the repair of R-loops (Makharashvili et al., 2018; Sollier et al., 2014), although with CPT, processing may be more pathological. In either case, the nature of repair could depend on the cell cycle stage, with recombination initiated after DSB formation in S-phase, and some sort of R-loop excision or unwinding initiated by break formation outside of S-phase. Clearly further work is needed to understand how these nucleases contribute to both repair and genome instability. As the ssDNA of an R-loop structure is no longer protected by the other DNA strand or bound proteins, it may also be more accessible and prone to damage via endogenous metabolites such as reactive oxygen species (ROS) or certain enzymatic activities. For instance, DNA nicks, such as those generated by ROS, can stabilize and promote R-loop formation (Roy et al., 2010). Targeting ROS to actively transcribed genes or telomeres promotes G4 and R-loop formation, resulting in replication-independent damage at these sites (Figure 5C) (Tan et al., 2020a, 2020b; Teng et al., 2018). Furthermore, damage in the extruded ssDNA could arise due to the action of cytidine deaminases such as the APOBEC proteins or AID (Saini et al., 2017; Yu et al., 2003; Zheng et al., 2017). The resulting uracil that forms would be subject to glycosylase processing and base excision repair, ultimately causing SSB formation (Figure 5D). Indeed, AID acts on R-loops in the switch regions during immunoglobulin class-switch recombination (Yu et al., 2003). RNA editing enzymes, such as ADAR1, may also modify the DNA moiety of a hybrid, providing an alternative mechanism of SSB formation (Zheng et al., 2017).

R-loops and Mitochondrial Genomic Stability

The impact of R-loops on genome stability is not restricted to the nuclear genome but extends to the mitochondrial DNA (mtDNA), where R-loops are important intermediates in mtDNA replication (Holt, 2019). Mitochondrial DNA is circular and consists of a heavy (H) and a light (L) strand, each of which contain an origin, OriH and OriL, respectively. Intriguingly, mtDNA replication is predominantly asynchronous, beginning on the H-strand. Before replication, R-loops form across OriH (Holt, 2019) then are processed by RNase H1 to generate the RNA primer from which the replicative polymerase initiates nascent DNA synthesis (Lima et al., 2016; Posse et al., 2019). Synthesis of the L-strand begins once replication of the H-strand reaches OriL (Falkenberg, 2018). Thus, mitochondrial R-loop regulation is vital to ensure that the mtDNA is properly replicated and that mitochondrial levels are maintained.

As a regulator of mtDNA replication, RNase H1 is particularly important in maintaining mitochondrial genome stability, and its levels need to be properly regulated. In Arabidopsis thaliana, loss of the RNase H1 homologs AtRNH1B and AtRNH1C caused mitochondrial R-loop accumulation and elevated levels of mitochondrial recombination. R-loop accumulation reduced mitochondrial copy number and caused embryonic lethality, suggesting that the mitochondrial genome is unstable (Cheng et al., 2021). Loss of RNase H1 in the liver of mice also led to R-loop-induced mitochondrial genomic instability. This instability was associated with mitochondrial dysfunction, including altered fusion and fission dynamics, and liver degeneration (Lima et al., 2016). In contrast, expression of catalytically inactive RNase H1 mutants in human cells reduced R-loop levels (Akman et al., 2016). In these cells, mitochondria aggregation was observed, indicating that mtDNA segregation is impaired, another feature of genomic instability. Perturbations of other genes may also cause RNase H1-mediated mitochondrial dysfunction. In BRCA2-deficient cells, ROS levels are elevated, generating 8-oxoguanine. This inhibits RNase H1 recruitment to OriH, thus mimicking RNase H1-deficiency (Renaudin et al., 2021). These cells accumulate R-loops in the mtDNA, leading to its instability. While the molecular mechanism by which RNase H1 loss destabilizes the mitochondrial genome is not clear, recent work suggests that mtDNA replication initiation is promiscuous in the absence of RNase H1, which likely accounts for this instability (Posse et al., 2019). Additionally, persistent hybrids can interfere with initiation and progression of L-strand DNA synthesis (Chang et al., 2020). Further investigation into this mechanism is required.

In addition to RNase H1, several other factors are implicated in maintaining the mtDNA integrity. In mammalian cells, members of the mitochondrial degradosome, such as SUV3 and PNPase, act analogously to RNase H1 to prevent R-loop accumulation in the mtDNA. In the absence of the degradosome, mtDNA synthesis was impaired and mitochondrial DNA content was lost, suggesting that the degradosome maintains mitochondrial genome stability (Silva et al., 2018). Proper localization of the helicase RECQ4 to the mitochondria is also required to regulate R-loop formation and resolution and maintain genome stability (Chang et al., 2020). As mitochondrial dysfunction is associated with multiple diseases, delineating how different R-loop resolution factors contribute to the maintenance of the mitochondria is imperative.

R-loops and DNA Break Repair

Double-strand breaks (DSBs) are one of the most toxic forms of DNA damage and are a primary driver of genome instability. Although early observations showed that unscheduled R-loops can cause DSB formation, DSBs have also been shown to elicit elevated R-loop levels. Cells employ a variety of strategies to repair DSBs, including homologous recombination (HR) and non-homologous end joining (NHEJ) (Chang et al., 2017b; Chapman et al., 2012). During HR, DSB ends are resected by nucleases, generating ssDNA. RPA and RAD51 bind to ssDNA and facilitate strand invasion into the sister chromatid, providing a template for error-free repair of the break. HR is primarily restricted to the S and G2 phases of the cell cycle, consistent with the requirement for a sister chromatid. During NHEJ, which is potentially error-prone, DNA DSB ends are initially recognized by the KU complex and a host of other factors, including 53BP1, that initiate processing of damaged ends but protect them from extensive resection prior to religation. Importantly, there is growing recognition that R-loops may directly influence the DSB repair process. Indeed, evidence exists that, while their deregulation can impede DSB repair, R-loops and hybrids are important intermediates that facilitate this process.

R-loops hinder DNA repair and augment genomic instability

The role of R-loops and hybrids in DSB repair is debated. On one hand, hybrids have been proposed to hinder the repair of breaks once formed. Recent work using a reporter in budding yeast where DSBs were induced at transcriptionally active or silenced regions demonstrated that hybrids accumulate at transcriptionally active breaks (Ortega et al., 2021). When R-loop resolution factors were lost, transcription impaired recombination and increased genome instability at induced DSBs. This effect could be rescued by overexpressing RNase H1, clearly suggesting that hybrids can interfere with DSB repair processes.

Although mechanisms by which hybrids impede repair are not fully understood, several studies suggest they may alter end resection processes. DNA ends are recognized by MRE11-RAD50-NBS1 (MRN) and CtIP, which initiate resection by nicking the DNA on either side of the break (Reginato and Cejka, 2020). EXO1 and DNA2 subsequently promote long-range resection, in a process requiring the BLM helicase (Nimonkar et al., 2011; Sturzenegger et al., 2014). The loss of certain R-loop resolution factors, such as DDX5, DHX9, and HNRNPD/SAF-A leads to hybrid accumulation and impaired end resection at induced DSBs (Figure 6A) (Alfano et al., 2019; Matsui et al., 2020; Sessa et al., 2021; Yu et al., 2020). In each case, hybrid retention diminished RPA and RAD51 loading and was associated with increased genomic instability. Therefore, accumulated hybrids at break sites may block nucleases from binding to the DNA ends, preventing resection. They may also mask ssDNA recognition by RPA, thereby inhibiting downstream RAD51 nucleofilament formation and homology search. In support of the former, in vitro experiments demonstrated that long hybrids at DNA ends cannot be unwound by BLM and inhibit EXO1 activity (Daley et al., 2020).

Figure 6. — Unscheduled R-loops Interfere with DNA Repair. (A) Various helicases, including DDX5 and DHX9, act with BRCA1 to resolve hybrids that form when a DSB occurs in an actively transcribed locus. Loss of DDX5 or DHX9 results in a persistent R-loop at the break site, which inhibits MRN/CtIP-mediated end resection, leading to HR defects. (B) (i) In budding yeast, Sen1 and RNase H (RH) resolve hybrids. When both factors are lost, persistent R-loops result in a DSB forming downstream of the R-loop. While resection can occur normally on the hybrid-free end, the R-loop blocks MRN/CtIP activity, preventing resection and resulting in genomic instability and cell death. (ii) In wild-type cells, BRCA2 recognizes an R-loop near a DSB and recruits RNase H2 (RH2) and EXO1. EXO1 resects the DNA strand, while RNase H2 degrades the RNA, promoting RAD51 nucleofilament formation and proper HR. Upon loss of BRCA2 or RNase H2, the hybrid persists, blocking RAD51 loading and impairing HR. (C) In normal cells, BRCA1 associates with RNAP to promote transcription termination and resolve R-loops at termination sites. Upon DSB induction, BRCA1 dissociates from RNAP and traverses to the break site to promote repair. In Ewing’s Sarcoma patients, EWS-FLI prevents BRCA1 from dissociating with RNAP, even in the presence of DNA damage.

The impact of RNase H1 and RNase H2 on end resection further supports a model whereby unscheduled hybrids inhibit resection. In budding yeast, persistent R-loops caused by co-depletion of RNaseH and Sen1, the yeast ortholog of SETX, led to DSB formation downstream of the R-loop, leaving only one end of the break with a hybrid (Amon and Koshland, 2016; Costantino and Koshland, 2018). While resection proceeded normally on the hybrid-free end, it was inhibited at the hybrid-containing end. It seems likely that the hybrid needs to be removed for resection to occur (Figure 6B). In mammalian cells, RNase H2 has also been implicated in removing hybrids at DSB sites to allow RAD51 loading (Figure 6B) (D’Alessandro et al., 2018). Consistent with this result, RNase H1/2 overexpression in Saccharomyces pombe (S. pombe) enhanced resection (Ohle et al., 2016). Taken together, these studies suggest that hybrid clearance is a common event at DSB sites, but if hybrids become deregulated, they can interfere with resection, leading to defects in repair and genome instability.

Interestingly, a number of well-known HR proteins interact with factors implicated in hybrid turnover at DSBs. For instance, BRCA2 stimulates the helicase activity of DDX5 toward hybrids, and perturbing this interaction prevents DDX5-mediated R-loop dissolution (Sessa et al., 2021). Similarly, the deubiquitinase USP42 promotes DHX9 activity towards R-loops at DSBs and is needed to recruit BRCA1 to break sites (Matsui et al., 2020). The close links between these HR and R-loop processing factors suggests that hybrid resolution is an integral part of DSB repair.

Another, albeit indirect, mechanism by which R-loops may impede repair is by sequestering repair proteins and preventing their recruitment to DSB sites. For instance, R-loops accumulate in cells expressing EWS-FLI, the fusion protein driving Ewing’s sarcoma, in response to pro-tumorigenic transcription (Gorthi et al., 2018). In EWS-FLI cells, BRCA1 remained associated with R-loop sites and could not relocalize to DSB sites after damage induction, while BRCA1 recruitment to DSBs was not altered in normal cells (Gorthi et al., 2018). Unscheduled R-loops in patient cells may sequester BRCA1, preventing repair (Figure 6C). Of note, auxin treatment in A. thaliana induces expression of the long non-coding RNA APOLO (Ariel et al., 2014, 2020). APOLO can invade duplexed DNA distal to its site of transcription to form R-loops, which are recognized by Polycomb Repressive Complexes (PRCs). APOLO sequesters PRCs away from auxin-responsive genes, upregulating their expression. As protein sequestration by APOLO represents a normal developmental process while sequestration by EWS-FLI is pathological, there may be multiple scenarios in which R-loops sequester proteins to control their function.

R-loops mediate the DNA damage response

In contrast to the idea that R-loops impede DSB repair is their emerging role in promoting the DNA damage response (DDR) and DNA repair (Marnef and Legube, 2021; Paull, 2019). In fact, hybrids have been proposed to be required for both HR and NHEJ, and their presence may affect repair pathway choice.

i. Non-homologous end joining

In certain contexts, R-loops may be involved in NHEJ. SETX loss triggered an increase in 53BP1 foci while inhibiting RAD51 foci formation (Cohen et al., 2018). These changes were accompanied by increased translocations, suggesting that hybrid persistence may favor error-prone repair. In agreement, Sen1 loss led to increased KU occupancy at break ends, activating an alternative resection pathway that promotes classical and microhomology-mediated NHEJ (Rawal et al., 2020). Interestingly, KU exhibits robust RNA and DNA binding activity, and weakly binds hybrids in extracts (Wang et al., 2018). In conjunction with the RNA binding protein RBM14, KU facilitated RNAPII and 53BP1 recruitment to DSBs, as well as hybrid formation (Jang et al., 2020; Schellenbauer et al., 2021; Simon et al., 2017). It is therefore possible that KU promotes both the formation and stabilization of hybrids at DSBs, creating a positive feedback loop that promotes NHEJ (Figure 7A). Of note, loss of other R-loop resolution factors, such as DDX1 or EXOSC10, augmented NHEJ efficiency at the cost of HR efficiency, and loss of USP42/DHX9 increased 53BP1 foci (Domingo-Prim et al., 2019; Li et al., 2016; Matsui et al., 2020). Excessive hybrids may therefore favor NHEJ in these contexts. By contrast, XRN-depleted glioblastoma cells, which also accumulate R-loops, exhibit decreased KU binding and impaired NHEJ, suggesting not all instances of unscheduled hybrids promote NHEJ (Dang and Morales, 2020).

Another mechanism by which hybrids and RNAs may promote NHEJ at a DSB is through the RNAi machinery (Figure 7B). Initial investigations found that the RNAi factors Drosha and Dicer process RNAs generated following break induction into DSB-induced small non-coding double-stranded RNAs, termed DDRNAs (Francia et al., 2012; Michelini et al., 2018). DDRNAs likely derive from the break site, as they have the same sequence as the damaged locus. More recent evidence suggests that DDRNAs originate from precursor RNAs, called damage-induced long non-coding RNAs (dilncRNAs) (Michelini et al., 2017; Pessina et al., 2019). De novo RNAPII transcription at a DSB leads to synthesis of dilncRNAs, which are processed into pre-DDRNAs by Drosha and then into DDRNAs by Dicer (Michelini et al., 2017, 2018). How dilncRNAs become double stranded is not well understood, but dilncRNAs can anneal to the DNA ends from which they derive, forming bona fide RNA-DNA hybrids. Loss of either Dicer or Drosha, or RNAPII inhibition, precludes the formation of 53BP1 or RIF1 condensates after DSB induction (Francia et al., 2016; Lu et al., 2018; Michelini et al., 2017). It is likely that NHEJ is compromised in these cells, as phase separation of repair proteins is a prerequisite DNA repair (Fijen and Rothenberg, 2021). These observations suggest a model whereby hybrids shift the balance of repair pathway choice towards NHEJ, by helping to both restrict resection and recruit NHEJ factors.

Hybrids may regulate alternative repair pathways, such as microhomology-mediated end joining, by controlling the localization and clustering of DSBs at the nuclear periphery. DSBs relocate to the nuclear periphery in order to perform such alternative repair pathways (Caridi et al., 2017). mRNA processing and export proteins, which suppress hybrid formation, also localize with nucleoporins and the nuclear periphery (García-Benítez et al., 2017), suggesting that hybrids may normally localize to the nuclear periphery as part of their resolution. Additionally, RNAs containing the m6A modification, which is also observed on hybrids, promote DSB clustering and phase separation (Garcia-Jove Navarro et al., 2019; Ries et al., 2019). Thus, hybrid-driven DSB clustering may be another mechanism by which hybrids promote repair, which has indeed been observed for NHEJ (Pessina et al., 2019). Exactly how they promote clustering, however, remains unclear.

ii. Homologous recombination

R-loops and hybrids also have been reported to direct cells towards homologous recombination. Early studies in S. pombe demonstrated that hybrids were needed transiently at DSBs to restrain resection and modulate RPA loading for efficient HR (Ohle et al., 2016). While in apparent contrast to its role in inhibiting resection, these findings may indicate that hybrid formation and turnover is needed to control resection. Another class of small RNAs, called DSB-induced small RNAs (diRNAs), may also regulate repair through hybrid formation (Figure 7C) (Gao et al., 2014). diRNAs map to sequences distal to a break and are thus thought to be distinct from dilncRNAs/DDRNAs. Here, the RNAi processing protein Ago2, but not Drosha or Dicer, recognizes diRNAs and mediates RAD51 recruitment to DSBs, promoting HR. While the exact mechanism by which Ago2 regulates RAD51 is not clear, it has been hypothesized that the diRNA-Ago2-RAD51 complex hybridizes to free DNA ends, forming a hybrid that drives its recruitment. Of note, Drosha processing has also been implicated in regulating HR, and its loss diminishes BRCA1 and RAD51 loading (Lu et al., 2018). As such, how small RNAs and hybrids help regulate repair requires further investigation.

Further supporting a role for R-loops in HR is work using a novel HR reporter in mammalian cells which revealed that HR was stimulated by local active transcription (Ouyang et al., 2021). HR was dependent upon the RAD51-associated protein 1 (RAD51AP1) and UAF1 protein complex, which were originally found to stimulate RAD51-mediated strand invasion (Liang et al., 2016; Ouyang et al., 2021). RAD51AP1 was shown to promote invasion of ssRNA into the template DNA, suggesting that RNA could invade the sister chromatid to form a type trans R-loop (Figure 7D). Concurrent invasion of ssDNA at these regions also occurs, forming a DR-loop. DR-loops enhanced RAD51-mediated invasion of the template DNA by ssDNA to generate a D-loop, an important intermediate in DSB repair. While it is tempting to speculate that de novo transcription drives DR-loop formation in this context, a tethered RNA molecule was sufficient to robustly stimulate DR-loop-driven HR. This result leaves open the possibility that the RNA transcripts involved in DR-loop formation have already been transcribed at the respective locus.

Recent evidence also implicates RNAPIII-driven hybrid synthesis in the regulation of DNA repair (Liu et al., 2021b). Indeed, RNAPIII, which transcribes transfer RNAs and other small RNAs, but not RNAPII, was recruited to DSBs, where it transcribed de novo hybrids in an MRE11 and CtIP-dependent manner from the 3’-strand (Figure 7E). Loss of RNAPIII activity and hybrid formation impaired RPA and RAD51 loading, suggesting that hybrid formation is necessary for HR. It was also proposed that RNAPIII-dependent transcription protects the 3’ DNA strand from exonucleolytic attack and the loss of genetic material, thereby maintaining genome stability.

R-loops can also propagate an alternative HR pathway known as transcription-associated homologous recombination repair (TA-HRR) (Yasuhara et al., 2018). During TA-HRR, R-loops that form near break sites in actively transcribed loci are recognized by RAD52, which promotes XPG-mediated hybrid removal to generate ssDNA in what might be considered the first step of resection (Figure 7F). Downstream HR factors, such as BRCA1, are also recruited by RAD52 to prevent the binding of RIF1 and 53BP1. A related model proposes that RAD52 recruitment following localized ROS damage requires hybrid recognition by the TC-NER factor CSB (Teng et al., 2018). After binding ROS-stabilized hybrids, RAD52 recruits RAD51 to promote strand invasion and HR. R-loops have also been shown to recruit RAD52 and CSB to ROS-induced breaks that occur near telomeres to promote their repair (Tan et al., 2020b), although this is more akin to break-induced repair (BIR) rather than canonical HR. Together, these works illuminate the need for transcription and R-loops to promote faithful DNA repair, particularly in highly transcribed regions.

iii. Single-strand break repair

A third class of damage-induced RNAs, single-stranded, DNA damage-associated sRNAs (sdRNAs) has been identified and implicated in SSB repair (Figure 7G) (Hatchi et al., 2021). In this pathway, BRCA1 and its partner BARD1, in conjunction with Dicer and Ago1, are thought to promote sdRNA synthesis at SSBs found at transcription termination sites (TTSs). BRCA1/BARD1 may then transfer the sdRNA moiety to the PALB2/RAD52 complex, which facilitates repair of the SSB. While the link between R-loops and sdRNA synthesis is unresolved, it is likely that R-loops have an important role in this pathway. sdRNAs map to SETX and BRCA1 occupied TTSs, which are prone to R-loop formation. Furthermore, previous work demonstrated that BRCA1 recruits SETX to TTSs to promote the repair of R-loop induced SSBs at these sites (Hatchi et al., 2015). Additional work is required to elucidate the role of R-loops in promoting SSB repair.

iv. RNA-templated DNA repair

An RNA molecule itself may be used as a template to direct the repair of the gene from which it is derived (Figure 7H). In budding yeast, reverse transcriptase can synthesize complementary DNA (cDNA) from an existing mRNA, which is then used as the homologous template for repair in a process called cDNA-templated DSB repair (Keskin et al., 2014; Meers et al., 2020). Conversely, the RNA itself can serve as the template during a process called RNA-templated repair (R-TDR). During R-TDR, the RNA molecule hybridizes with the free ssDNA end via an inverse strand exchange mechanism stimulated by RAD52, forming a hybrid. DNA is then directly synthesized from the RNA by Polymerase ζ (Mazina et al., 2017; Meers et al., 2020). Of note, Polθ can also reverse-transcribe cDNA from an RNA template (Chandramouly et al., 2021). Importantly, only RNA transcribed at the same locus as the break, and not homologous RNA transcribed at a distal locus, can undergo inverse strand exchange (Keskin et al., 2014; Meers et al., 2020). RAD52 and CSB also recognize hybrids to drive RNA-directed recombination in G0/G1 mammalian cells, providing additional context in which RNA could be used to drive error-free repair (Wei et al., 2015).

RNA-DNA hybrid formation at DSB sites

To resolve questions regarding how R-loops facilitate repair, the mechanism by which hybrids form at break sites must be determined. Specifically, whether hybrids are products of de novo transcription that occurs after the break forms or whether they result from preexisting transcription at the break site remains an intriguing question.

In support of the first hypothesis, recent work provides evidence of de novo transcription at DSBs in mammalian cells (Liu et al., 2021b; Michelini et al., 2017). In response to damage, MRN can recruit the transcriptional preinitiation complex (PIC), including the transcriptional coactivator Mediator and the RNAPII kinases CDK7 and CDK9, to DNA ends (Pessina et al., 2019; Sharma et al., 2021). During normal transcription, the PIC, CDK7, and CDK9 act in concert to phosphorylate and activate RNAPII, and they may have a similar role at DSBs (Larochelle et al., 2012). The PIC, in conjunction with MRN, promotes RNAPII association with break sites, resulting in local dilncRNA synthesis (Pessina et al., 2019). It is thought that dilncRNAs hybridize with DNA at the break site, accounting for the hybrids observed at DSBs. Surprisingly, MRN’s nuclease activity seems to be dispensable for RNAPII recruitment, which instead requires MRN’s DNA melting activity (Sharma et al., 2021). Consistent with this finding, work in S. pombe demonstrates MRN-dependent RNAPII-mediated hybrid formation at DSBs, although in this case MRN’s end processing activity was required (Ohle et al., 2016). Similarly, the synthesis of diRNA and sdRNA appears to occur de novo at DSBs (Gao et al., 2014; Hatchi et al., 2021). Work in mammalian cells also supports a role for RNAPIII in hybrid generation at break sites (Liu et al., 2021b). RNAPIII localization to DSBs required the nuclease activities of MRN as well as CtIP (Liu et al., 2021b). In each instance, de novo hybrid formation promotes the recruitment of downstream repair factors to break sites, providing insight into the potential mechanisms by which hybrids propagate repair signaling pathways.

While these works support the concept of de novo RNAPII transcription and hybrid formation at break sites, the exact mechanism by which processed DNA ends support transcription is still unclear. There are mechanisms for RNAP-mediated transcription at ssDNA sites lacking a promoter sequence (Kadesch and Chamberlin, 1982). It is also likely that various helicases are required to promote new transcription and hybrid formation at DSBs. For example, the helicase UPF1 promotes hybrid formation at regions where a DSB has been induced (Ngo et al., 2021). Although further studies are needed, UPF1 was proposed to both unwind dsDNA to promote RNAPII elongation and to translocate along the nascent RNA to remove RNA binding proteins, both of which would facilitate hybrid formation.

Alternatively, R-loops observed at DSBs may be an unwanted consequence of altered transcriptional dynamics in response to break induction. Indeed, R-loops may arise due to changes in promoter proximal pausing or termination even in the absence of damage (Shivji et al., 2018; Skourti-Stathaki et al., 2011; Zhang et al., 2017). As such, DSB-induced transcriptional repression, which occurs both globally and locally, could drive R-loop formation (Shanbhag et al., 2010; Silva and Ideker, 2019). Consistent with this idea, hybrids were elevated only at DSBs occurring in actively transcribed regions upon SETX loss (Cohen et al., 2018). DRIP-seq analysis similarly revealed elevated hybrid occupancy specifically at DSBs in transcriptionally active loci (Bader and Bushell, 2020). This finding was corroborated by mNET-seq, which showed that elongating transcripts are only found at DSBs induced in actively transcribed genes (Burger et al., 2019). While the above techniques utilize enzyme-mediated DSB formation, DSBs induced by ROS also stimulate R-loop formation in a transcription-dependent manner (Teng et al., 2018). These findings suggest that DSBs in actively transcribed loci may induce hybrid formation by either changing where the polymerase resides within that gene or slowing elongation rates. R-loops and hybrids may arise as a consequence of these altered dynamics.

R-loops and DSB crosstalk: in trans or in cis?

While there is a general consensus that R-loops can form in cis at sites of DNA breaks, an intriguing possibility is that these structures can also form in trans. Recent work has shown that trans R-loops can form in a variety of contexts and these R-loops may have physiological roles. As described previously, in vitro experiments that demonstrate RAD51AP1-mediated RNA invasion of template DNA using a ssRNA oligo suggest that trans-forming R-loops may help promote DSB repair (Ouyang et al., 2021). Another example of trans R-loop formation is the well-known telomeric lncRNA TERRA, which can invade short telomeric repeats in trans in a RAD51-dependent manner to maintain telomere integrity (Feretzaki et al., 2020). Furthermore, the APOLO lncRNA in A. thaliana can form trans R-loops via a mechanism described previously (Ariel et al., 2020). Whether trans hybrid formation is consistently associated with a physiological role remains to be determined, but these works highlight the capability of RNA to invade DNA in trans.

Less well-known, however, is the propensity for trans-forming R-loops to drive DSB formation and genome instability. An important example of this is trans R-loop formation during CRISPR/Cas9-guided DSB induction (Jinek et al., 2012; Nishimasu et al., 2014). Here, Cas9 opens the DNA duplex, allowing the guide RNA to base pair with its complementary DNA sequence and displace ssDNA. As hybrids direct Cas9 nuclease activity, they represent a source of DNA damage in this context. Intriguingly, trans R-loops have also been directly implicated as a source of genomic instability. Initial studies using a yeast artificial chromosome (YAC) system showed that Rad51 facilitated the trans invasion of RNA transcribed from the yeast genome into YAC DNA, forming an R-loop that ultimately led to YAC loss, a readout for genomic instability (Wahba et al., 2013). Other studies, however, found that only cis R-loops generate DNA damage, and that this occurs in a RAD51-independent process (Lafuente-Barquero et al., 2020).

R-Loops and Immune Signaling

Given that R-loops cause DNA damage, it seems likely that they may also have an effect on the innate immune response, a pathway known to protect cells from pathogens and viruses by detecting foreign nucleic acids in the cytoplasm (Zhou et al., 2017). Normally, recognition of nucleic acids through pattern recognition receptors, including the Toll Like Receptors (TLRs) and the cGAS/STING pathway, activates downstream inflammatory signaling mediated by IRF3, NF-kB, and MAPK (Brubaker et al., 2015; Takeuchi and Akira, 2010; Zhou et al., 2017). Recognition of self-nucleic acids resulting from DNA damage and replication stress has also been shown to induce innate immune signaling (Hopfner and Hornung, 2020; Ragu et al., 2020). Although the mechanism by which damage activates innate immune signaling is not fully understood, several sources of endogenous nucleic acids that activate this response have been identified in the cytoplasm. For example, micronuclei arising from DNA damage, ssDNA arising from stalled fork processing, and mitochondrial RNA released after damage-induced mitochondrial herniation can activate cGAS or RIG-I (Coquel et al., 2018; Harding et al., 2017; Mackenzie et al., 2017; Tigano et al., 2021).

These studies raise the question of whether R-loops may also contribute to innate immune signaling. Consistent with such an idea, RNase H1 overexpression reduced the level of cytosolic DNA and the interferon response in tumor cells (Shen et al., 2015) and in cells in which R-loop accumulation was induced by DNA damage (Chatzidoukaki et al., 2021). Similarly, loss of the DEAD-box helicase 41 (DDX41) led to RNase H-reversible nuclear R-loop accumulation and the expression of genes involved in the inflammatory response in both zebrafish embryos and human cells (Weinreb et al., 2021). Intriguingly, inflammatory signaling driven by excessive R-loop production was also shown to increase hematopoietic stem and progenitor cell production in zebrafish models, raising the possibility that R-loops may control developmental processes through the inflammatory response. Factors that dampen the inflammatory response induced by hybrids also exist. Hybrid binding to lysyl tRNA synthetase reduces the inflammatory response through two mechanisms: by suppressing the interaction of hybrids with cGAS, thereby reducing cGAMP production in cells, and by stimulating diadenosine tetraphosphate production, which can act as an antagonist to STING and reduce STING-dependent signaling (Guerra et al., 2020).

Whether hybrids directly activate the innate immune response or whether the activation is a result of the damage produced by hybrids is still not clear. However, there are hints they might be in a position to directly activate innate immune responses. First, hybrids have been detected in the cytoplasm of virus-free cells by immunofluorescence (Koo et al., 2015). Recent work, however, has demonstrated that the majority of signal from the S9.6 antibody, which recognizes dsRNA and hybrid structures, derives from dsRNA and not hybrid recognition (Crossley et al., 2021; Smolka et al., 2021). As such, further characterization is needed to verify these species as hybrids. Second, the transfection of in vitro generated synthetic RNA-DNA hybrids has been shown to induce a cGAS- or TLR9-dependent response, depending on the cell type (Mankan et al., 2014; Rigby et al., 2014). Synthetic hybrids can also activate cGAS in vitro (Mankan et al., 2014). Although these observations suggest hybrids could directly interact with pattern recognition receptors in cells, additional work will be needed to verify this idea and to understand the mechanism of this activation.

While the accumulation of R-loops is associated with the induction of an inflammatory response, there is also evidence that increased inflammatory signaling induces R-loop formation. NF-kB activation is a downstream effector of the inflammatory response that can be activated by the HTLV-1 transactivator/oncoprotein Tax to promote cellular senescence. In adult T leukemia (ATL) cells, NF-kB activation by TAX leads to R-loop dependent DNA damage and senescence in a pathway mediated by XPF and XPG. Many ATL cells are deficient for these endonuclease activities, which may help them proliferate (He et al., 2021). Similarly, NF-kB-mediated innate immune signaling activated by Helicobacter pylori infection causes R-loop-induced replication stress, and cancers associated with this infection demonstrate higher mutational burden (Bauer et al., 2020).

These findings illustrate how R-loops could contribute to immune signaling, but whether R-loop formation is causative for or consequential to innate immune signaling is still unclear. One can envision an R-loop-dependent positive feedback loop, whereby hybrids activate cGAS/STING to elicit an immune response, which further drives R-loop formation during the change in transcription.

R-Loop Pathology

Deregulation of R-loop levels has also been associated with multiple human diseases, including cancers, neurological disorders and autoimmune diseases (Table 1). In these situations, R-loops may drive genome instability, alter gene expression or alter cellular signaling pathways. Understanding the mechanisms behind R-loop-driven disease could reveal novel therapeutic opportunities. Below, we provide an update on R-loop regulation and disease connections, though we also refer the reader to other in-depth reviews (Groh and Gromak, 2014; Richard and Manley, 2017).

Table 1.

R-loop resolution and suppression factors and their links to disease. Bold denotes a gain-of-function mutation for that factor associated with disease, while italics signifies a loss-of-function mutation.

Gene	Description	Disease	Reference
ADAR1	Edits mismatches to promote RNase H2 R-loop resolution	Telomerase-Reactivated Cancers	(Shiromoto et al., 2021)
BRCA1	Suppresses satellite RNA production to suppress R-loops; also sequestered by satellite RNAs preventing its proper functioning	Breast and Ovarian Cancers	(Zhu et al., 2018)
	Resolves R-loops at transcription termination sites (with SETX)		(Hatchi et al., 2015)
	Limits RNAPII proximal pausing, preventing instability		(Zhang et al., 2017)
	Limits the ESR1 R-loop to maintain neighboring gene transcription		(Chiang et al., 2019)
BRCA2	Resolves R-loops and limits genomic instability	Breast and Ovarian Cancer	(Bhatia et al., 2014)
	Depletion by aldehydes increases replication stress and instability		(Tan et al., 2017)
	Promotes the transition of RNAPII from an initiating to an elongating complex		(Shivji et al., 2018)
C9orf72	R-loops formed over this region interfere with transcription and promote disease pathologies	ALS/Frontotempor al Dementia (FTD)	(Haeusler et al., 2014; Reddy et al., 2014)
CDCA7/HELLS	Promotes methylation at pericentromeres to suppress R-loops	ICF Syndrome	(Unoki et al., 2020)
CSB	Processes R-loops to limit levels	Cockayne Syndrome (CS)	(Sollier et al., 2014)
Dicer	Prevents R-loop associated genome instability	Embryonal Tumors with Multilayered Rosettes (ETMR)	(Lambo et al., 2019)
DDX41	Recognizes viral hybrids to initiate immune signaling	Human Immunodeficiency Virus (HIV)	(Stavrou et al., 2018)
	Regulates alternative splicing	Cervical Cancer	(Qin et al., 2021)
	Suppresses R-loops and R-loop-induced immune signaling	Myelodysplastic Syndromes	(Weinreb et al., 2021)
Estrogen	Induces transcription and R-loops, leading to instability	Breast/Ovarian Cancer	(Stork et al., 2016)
EWS-FLI	Promotes R-loop formation and prevents BRCA1 localization to damage sites	Ewing’s Sarcoma	(Gorthi et al., 2018)
FANCM FANCD2	Resolves TERRA R-loops to limit genome instability and regulate ALT activity	ALT cancers	(Pan et al., 2019; Silva et al., 2019)
FANCM FANCD2	Suppresses R-loops and TRCs to maintain genomic stability	Fanconi Anemia	(García-Rubio et al., 2015; Schwab et al., 2015)
FXN FMR1	R-loops in these genes lead to transcriptional silencing	Fragile X Syndrome Friedreich Ataxia	(Colak et al., 2014; Groh et al., 2014; Loomis et al., 2014)
H. pylori	Activates Nf-kB-driven expression, leading to TRCs and damage	Gastric Cancer	(Bauer et al., 2020)
HRAS	Increases RNA synthesis and R-loops leading to increased damage and replication stress	Many Cancers	(Kotsantis et al., 2016)
INO80	Remodels chromatin to suppress R-loops and prevent TRCs	Multiple Cancers	(Prendergast et al., 2020)
MDM2/RNF2	Ubiquitinates H2A to increase replication and prevent R-loops	Sarcomas	(Klusmann et al., 2018)
MYCN	MYCN recruits BRCA1 to release RNAPII from proximal pause to prevent R-loops	Neuronal and Neuroendocrine Cancers (eg. Neuroblastoma)	(Herold et al., 2019)
RECQ4	Resolves mitochondrial R-loops to promote mtDNA replication	Rothmund-Thomson Syndrome, RAPADILINO, Baller-Gerold Syndrome	(Chang et al., 2020)
RNase H2	Suppress R-loop formation and genomic instability	Aicardi-Goutières Syndrome (AGS)	(Giordano et al., 2022; Mackenzie et al., 2016)
SAMHD1	Suppress R-loop accumulation and R-loop induced TRCs	AGS	(Lim et al., 2015; Park et al., 2021)
SAN1	Interacts with SETX to resolve R-loops	Cardiomyopathies	(Liu et al., 2021c)
SETX	Resolves R-loops to prevent instability	AOA2	(Becherel et al., 2015)
	Resolves R-loops to prevent aberrant TGF-β signaling	Amyotrophic Lateral Sclerosis (ALS4)	(Grunseich et al., 2018)
	Interacts with ZPR1 to resolve R-loops	Amyotrophic Lateral Sclerosis (ALS4)	(Kannan et al., 2022)
	Promotes meiosis	Infertility	(Becherel et al., 2013)
SF3B1 U2AF1 SRSF2	Maintain splicing to prevent R-loops	Myelodysplastic Syndromes	(Cheruiyot et al., 2021; Sorrells et al., 2018)
	Promotes RNAPII release from promoter pause sites		(Chen et al., 2018)
	When lost, R-loops activate ATR to promote survival		(Nguyen et al., 2018)
SMN1	Promotes splicing to prevent R-loops and instability	SMA	(Jangi et al., 2017)
Snord116	Loss of R-loops in this region leads to altered imprinting and chromatin condensation	Prader-Willi Syndrome	(Powell et al., 2013)
SS18-SSX	Increases R-loop formation	Synovial Sarcoma	(Jones et al., 2017)
TDP-43	Suppress hybrid formation and replication stress	ALS/FTD	(Giannini et al., 2020; Wood et al., 2020)
TDRD3/TOP3B	Induces negative supercoiling and increases R-loop resolution via DHX9 to prevent c-MYC/IgH translocations	Multiple Myeloma Burkhitt’s Lymphoma	(Yang et al., 2014; Yuan et al., 2021)
TERRA	DNMT3B mutations in ICF cells elevate TERRA, increasing R-loops and causing telomere shortening	Immunodeficiency, Centromere instability and facial anomalies syndrome (ICF)	(Sagie et al., 2017)
TERRA	Increased production leads to increased R-loops and recombination to lengthen telomeres, mediated by RNase H1	Alternative-Lengthening of Telomeres (ALT)-driven cancers	(Arora et al., 2014)
TonEBP	Recruits METTL3 to resolve R-loops and maintain stability	Hepatocellular carcinoma	(Kang et al., 2021; Ye et al., 2021)
TPR	Prevents R-loop formation and maintains fork progression	Ovarian cancers Others	(Kosar et al., 2021)
TREX (Transcription-export) complex	Prevents R-loop formation and DNA breaks; sequestered and inhibited by KSHV ORF57	Kaposi’s Sarcoma	(Jackson et al., 2014)
TREX1	Suppress R-loop formation and maintain genome stability; maintain DNA methylation	AGS	(Giordano et al., 2022; Lim et al., 2015)
XPF	Processes R-loops to limit levels	XP	(Cristini et al., 2019; Goulielmaki et al., 2021; Makharashvili et al., 2018; Sollier et al., 2014)
XPF	Processes R-loops to limit cytosolic ssDNA accumulation and inflammation	Chronic Pancreatitis	(Chatzidoukaki et al., 2021)
XPG	Processes R-loops to limit their levels	Xeroderma Pigmentosum (XP)	(Cristini et al., 2019; Goulielmaki et al., 2021; Makharashvili et al., 2018; Sollier et al., 2014).
ZPR1	Interacts with SETX to resolve R-loops	ALS4	(Kannan et al., 2022)
ZPR1	Regulates SETX and SMN2 levels to promote R-loop resolution	Spinal Muscular Atrophy (SMA)	(Kannan et al., 2020, 2022)

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Cancers

The ability of R-loops to cause replication stress and DNA damage, two hallmarks of cancer, suggest these structures could drive tumorigenesis. Indeed, several oncogenes and proliferation factors promote R-loop formation and R-loop associated DNA damage, as do loss-of-function mutations in tumor suppressors and other genes that suppress and resolve R-loops or TRCs (Table 1). For example, estrogen and HRAS both drive R-loop-dependent DNA damage, and mutations are found in genes where estrogen-responsive R-loops form (Stork et al. 2016; Kotsantis et al. 2016). Dicer mutations are also associated with R-loop induced chromosomal instability in pediatric brain tumors (Lambo et al., 2019). Similarly, the Cyclin E and MYC oncogenes promote early S phase entry, leading to intra-genic activation of origins that are normally deactivated by G1-phase transcription. This leads to an increase in HO TRCs and chromosomal rearrangements at sites altered in human cancers (Macheret and Halazonetis, 2018). As HO TRCs and some oncogenes cause R-loop accumulation (Table 1), R-loops may drive the genome instability occurring at these sites. R-loop-driven instability may promote both specific mutational signatures that are associated with specific cancers and the evolution of tumorigenic traits.

R-loops may also promote tumorigenesis by supporting telomere maintenance. In ALT-driven cancers, upregulation of the TERRA RNA and subsequent R-loop formation promotes telomeric recombination to lengthen telomeres (Table 1). In ALT-driven cancers, R-loop and G4 formation is also interdependent, and these structures appear to act synergistically in driving ALT (Yang et al., 2021). Chromatin remodeling at TERRA may also link R-loops to telomere maintenance. For example, mutation of the histone chaperone ATRX, which normally suppresses TERRA expression, is common in ALT cancers (Clynes et al., 2015; Flynn et al., 2015; Goldberg et al., 2010).

How R-loop deregulation contributes to tumorigenesis is complex. At one extreme, low R-loops levels could drive mutation, but high R-loop levels may cause excessive damage and cell death. As discussed, increased DNA damage activates immune signaling and elicits an innate immune response (Coquel et al., 2018; Harding et al., 2017), leading to chronic inflammation, which can drive tumorigenesis. If left unchecked, inflammation may also lead to apoptosis or elimination of the tumor cell by the immune system. Cancer cells may adapt to this state, downregulating the immune system or promoting a tolerance to R-loops. In patients with BRCA1/2-deficient breast cancer, high levels of RNF168, a factor that recruits the R-loop resolving helicase DHX9 to R-loop prone loci, are associated with poor prognosis. Moreover, RNF168 loss in a BRCA1/2 mouse prevents tumorigenesis (Patel et al., 2021). R-loop levels may need to be regulated even when pathologic in nature, and understanding how R-loops activate and avoid immune responses could provide insight into ways to improve immunotherapy.

Importantly, elevated R-loop levels and R-loop associated DNA damage proffer additional avenues by which therapies can capitalize upon heightened instability in cancer cells. For instance, mutations in splicing factors causative for myelodysplastic syndromes and acute myeloid leukemia initiate R-loop dependent ATR signaling (Nguyen et al., 2018). ATR inhibitor treatment exacerbated DNA damage and led to cell death, making ATR inhibition an attractive therapy for diseases harboring such mutations. As ATR is activated by R-loops, its inhibition may represent a broadly generalizable therapeutic strategy for many diseases that augment R-loop levels. Splicing factor mutations also increase cellular sensitivity to Chk1 and PARP inhibition, providing alternative therapies (Lappin et al., 2022; Singh et al., 2020). Interestingly, some of these mutations result in a BRCA1-like phenotype, suggesting there may be overlap in therapeutic strategies between BRCA1-driven diseases and R-loop diseases (Lappin et al., 2022). In addition, inhibiting the RNA degradation machinery, which also results in excessive R-loop formation and damage, may be effective in cells already bearing a high R-loop burden (Cheruiyot et al., 2021).

Neurological Disorders

Alterations in R-loop levels are also strongly associated with several neurological disorders (reviewed in Perego et al., 2019). One such factor mutated in disease on which a number of studies have focused is SETX. Activating and loss-of-function mutations in SETX, found in amyotrophic lateral sclerosis type 4 (ALS4) and ataxia with oculomotor apraxia2 (AOA2), respectively, are thought to drive disease progression (Table 1). Hyperactive SETX reduces R-loop levels, leading to increased gene silencing in ALS4 patients, including at one gene that suppresses TGF-β expression. Increased TGF-β signaling is thought to ultimately cause neuronal death (Grunseich et al., 2018). Conversely, SETX loss in AOA2 results in elevated R-loops at promoter-proximal regions, changes in transcription and chromosomal instability in gene bodies and at common fragile sites (Becherel et al., 2015; Kanagaraj et al., 2022). R-loops in AOA2 cells are processed via an auxiliary pathway involving CSB, XPG and RAD52, which may contribute to the observed genomic instability (Kanagaraj et al., 2022). Other studies have focused on regulators of SETX demonstrating that SETX protein levels are regulated by ubiquitin-mediated degradation, controlled by the E3 ligase KEAP1 and the deubiquitinase USP11 (Jurga et al., 2021). Targeting USP11 in ALS4 patients to limit the activity of mutated SETX may present a viable therapy, and applications in cancer cells can also be envisioned to overload cells with R-loops.

Emerging evidence also supports a role for R-loops in neurodegenerative diseases associated with repeat instability, including trinucleotide repeat (TNR) expansion or deletion. This includes Huntington’s Disease, Fragile X syndrome, Friedreich’s ataxia, myotonic dystrophy and several motor neuron disorders and ataxias (reviewed in Groh and Gromak, 2014; Malik et al., 2021). TNRs are hotspots for R-loop formation, as they are prone to form secondary structures in the ssDNA which stabilize hybrids on the template strand (Loomis et al., 2014). Interestingly, R-loops contribute to repeat instability by promoting both repeat expansion and deletion. Transcription through GAA repeat regions results in repeat expansion upon RNase H1/2 loss in budding yeast. Although the mechanism is not fully understood, it is likely that R-loops formed at these repeats impede the replisome, causing fork stalling, collapse and then BIR, which leads to TNR expansion (Neil et al., 2018).

Conversely, R-loops drive TNR contraction at CAG repeats. In this case, it is thought that hairpin structures which form in the displaced ssDNA of the R-loop are targeted by cytosine deaminases and downstream repair factors generating abasic sites or DNA nicks. Subsequent removal of the RNA strand of the R-loop allows for additional hairpin formation in the template strand, which allows for repeat deletions when bypassed by Polβ during break repair (Laverde et al., 2020). Hybrid removal can also lead to the formation of mismatches, which are subject to processing by the MutLγ complex, driving break formation and repeat loss (Su and Freudenreich, 2017). It will be interesting to see how R-loops differentially regulate changes in TNR tracts depending upon the type of repeat.

R-loops in repeat regions can also alter the chromatin landscape leading to disease-associated transcriptional silencing. For example, in Friedreich’s ataxia and Fragile X syndrome, stable R-loops in the repeat region of the frataxin (FXN) and FMR genes promote H3K9 dimethylation, a marker of transcriptional repression (Groh et al., 2014). Loss of expression of these genes may be an important driver of neurodegenerative disorders like Fragile X Syndrome. Hence, induction of chromatin repression represents an additional mechanism by which R-loops can drive neurodegeneration.

The extent to which R-loops are responsible for the progression of these different neurodegenerative disorders remains unresolved. Many animal models that recapitulate these disease mutations do not demonstrate elevated R-loop levels and fail to result in similar phenotypic abnormalities (Becherel et al., 2013). Deciphering these discrepancies will be important to further our understanding of these diseases and illuminate the propensity for R-loops to be targeted as a therapy. Furthermore, as these are diseases of post-mitotic cells, the pathological effects of R-loops in these situations should be independent of replication. However, it remains possible that events occurring in dividing cells during development could affect the progeny, by altering their epigenetic state or some other feature. Clearly, further work will be required to address this issue.

Autoimmune Disease

Aicardi Goutières Syndrome (AGS) is an inflammatory/autoimmune disorder associated with the accumulation of nucleic acid fragments in the brain that can be caused by a number of genes involved in processing RNA and DNA, including SAMHD1, RNaseH2, TREX1, and ADAR1 (Table 1). Interestingly, mutations in these genes also lead to increased innate immune signaling, through a mechanism that is not well understood. For instance, loss of SAMHD1 leads to aberrant replication fork processing and the release of ssDNA into the cytosol, activating cGAS-STING signaling (Coquel et al., 2018). Similarly, loss of RNase H2 activates cGAS and is associated with increased micronuclei formation (Giordano et al., 2022; Mackenzie et al., 2016). Perturbation of ADAR1 also activates immune signaling, though this is thought to be through increased dsRNA (Guo et al., 2021; Inoue et al., 2021). However, perturbation of SAMHD1, RNase H2, TREX1, or ADAR1 also elevates R-loop levels (Giordano et al., 2022; Park et al., 2021). As emerging evidence demonstrates that R-loops can activate immune signaling, it will be interesting to assess the extent to which R-loop deregulation is associated with the inflammation observed in AGS patients.

Concluding Remarks

The works described herein have begun to decipher the dichotomous role for R-loops in maintaining genomic integrity. In the context of replication, whether R-loops directly impede fork progression and/or whether they also affect forks post-replicatively requires further investigation. How forks can bypass R-loops is also of great interest. Although our knowledge of this process is increasing, many questions remain about whether RNAPII is bypassed or removed and whether this is context-dependent. Which R-loops threaten the fork is also an outstanding question. The pervasive nature of R-loops and their physiological roles suggest the cell can tolerate them under most circumstances, and recent studies indicate only a small subset may cause replication stress under normal circumstances. Further work will be needed to understand R-loop tolerance, and could be important for understanding how cancer cells may tolerate high levels of R-loops. It seems likely that potential positive roles for hybrids at the fork may also emerge in the years to come, given the similarities in processing stalled forks and DSBs. Consistent with this idea, it was proposed that hybrids may form de novo at reversed replication forks (Adeyemi et al., 2021). Protexin, a complex of SCA1 and the Rev3 polymerase, recruits RNAPII to the reversed fork to protect the fork from degradation, similar to what has been observed at DSBs.

Whether R-loops interfere with or actually mediate DNA repair also necessitates further investigation. Indeed, it is likely that both are true. They may be crucial intermediates in the DDR and in controlling DNA repair, but their removal may also be necessary to complete repair. The extent in which R-loops play a role in pathway choice is still unclear. Does an R-loop pre-dispose a cell to repair damage via one pathway over the other, or do they inhibit certain forms of repair as a consequence of being in the wrong place at the wrong time? As most studies focus on how R-loops affect repair when their formation or turnover is deregulated, the problems observed are likely to result from being unable to remove unscheduled R-loops. As studies are beginning to implicate R-loops as intermediates in both NHEJ and HR, it will be interesting to see in what contexts R-loops favor one pathway over another, and what determines which damage-induced RNA species is formed.

Another important question is why so many factors contribute to R-loop prevention and resolution, and to what extent these factors cooperate to regulate R-loop levels? The field also needs to move past simple analysis of hybrid levels at a few common genes or by imaging. Though a herculean effort, using quantitative approaches to analyze the genomic distribution of unscheduled R-loops could provide insight into this question (Crossley et al., 2020; Šviković et al., 2019). Such large-scale studies could reveal the differential contexts in which these factors might act, such as specifically at long genes, only when a certain chromatin mark is present, or during a specific stage of the cell cycle. This may be more challenging than it appears, however, as many R-loop processing proteins have roles in transcription, which also alters R-loop levels. It is also important to discern whether the ability of so many different cellular perturbations, ranging from distinct forms of DNA damage to disruption of RNA processing and replication factors, may cause R-loops to accumulate for indirect reasons. Many of these stressors cause replication and transcription stress, both of which may exacerbate the other. Thus, R-loop accumulation could reflect replication stress itself and even be a marker of DNA damage or genome instability. For instance, R-loops could arise as a result of replication stress due to the firing of dormant origins, which could increase HO conflicts by disrupting the normal replication program, or they could arise due to the increased ssDNA formation associated with replication stress. The high tolerance of cells to physiological R-loops in general indicates they are not normally a problem.

In this context, it will also be important to determine which unscheduled R-loops present a problem and contribute to genomic instability. One interesting possibility is that the analysis of sites of DNA damage will reveal the different contexts in which R-loop resolution factors are most important. Will the same subset pose a threat following different perturbations or will different R-loops contribute to damage and genome instability under different conditions? As methods for capturing and sequencing R-loops and hybrids have improved, it is more apparent that hybrids are not uniform structures but are rather heterogeneous. Recent insights suggest that R-loops exist in at least two distinct subclasses: pausing-associated and elongating-associated R-loops (Castillo-Guzman and Chédin, 2021). A subclass of R-loops that cause genomic instability may also exist, and if so, what intrinsic properties of those R-loops lead to their pathological nature? A class of RNase H-resistant R-loops was recently identified, which may provide hints into what makes some R-loops harmful (Crossley et al., 2020). R-loops that drive damage may be more intrinsically stable or more likely to contain secondary structures, such as G4s, in the displaced ssDNA, RNA, or DNA near the R-loop that confers resistance to certain resolution pathways. Alternatively, they could be associated with specific chromatin states that make them less accessible to non-damaging forms of R-loop resolution. Along these lines, are the R-loops that cause damage more likely to be processed by nucleases, and does this processing represent an auxiliary, last resort pathway to reduce R-loop levels? Investigation into each of these questions could significantly advance our understanding of R-loop biology.

Finally, the strong link between different diseases and R-loop formation suggests that manipulating and targeting R-loop pathways could lead to novel therapeutic approaches. Additionally, as products of transcription, the distinct perturbations that elevate R-loops and drive cancer could have different profiles, rendering the characterization of R-loops from a tumor a useful diagnostic tool.

Acknowledgements

We thank Stephan Hamperl, Madzia Crossley and Theresa Endres for critically reading this manuscript. We apologize to all authors whose work could not be discussed due to space constraints. This work was supported by the Jane Coffin Childs Memorial Fund for Medical Research [61-1755 to JRB] and the National Institutes of Health [GM119334 to KAC]. KAC is an American Cancer Society Research Professor. All figures were created with BioRender.com.

Footnotes

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Declaration of Interests

Karlene Cimprich is a member of the Advisory Board for Molecular Cell.

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PERMALINK

Walking a tightrope: the complex balancing act of R-loops in genome stability

Joshua R Brickner

Jada L Garzon

Karlene A Cimprich

Summary

Introduction

Figure 1.

R-Loops and the Replication Stress Response

R-loop-induced fork stalling: the nature of the block

Figure 2.

Mechanisms for restart

i. Fork bypass of R-loops and replication restart

Figure 3.

ii. Resolution of the R-loop or transcription block

Figure 4.

iii. Origin firing and genome organization

iv. TRC avoidance by fork pausing

v. TRC avoidance by chromatin remodeling

R-loops and fork progression

Replication-Independent Mechanisms for R-Loop Induced DNA Damage and Genome Stability

Figure 5.

R-loops and Mitochondrial Genomic Stability

R-loops and DNA Break Repair

R-loops hinder DNA repair and augment genomic instability

Figure 6.

R-loops mediate the DNA damage response

i. Non-homologous end joining

Figure 7.

ii. Homologous recombination

iii. Single-strand break repair

iv. RNA-templated DNA repair

RNA-DNA hybrid formation at DSB sites

R-loops and DSB crosstalk: in trans or in cis?

R-Loops and Immune Signaling

R-Loop Pathology

Table 1.

Cancers

Neurological Disorders

Autoimmune Disease

Concluding Remarks

Acknowledgements

Footnotes

References

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