S phase R-loop formation is restricted by PrimPol-mediated repriming

During DNA replication, conflicts with ongoing transcription are frequent and require careful management to avoid genetic instability. R-loops, three stranded nucleic acid structures comprising a DNA:RNA hybrid and displaced single stranded DNA, are important drivers of damage arising from such conflicts. How R-loops stall replication and the mechanisms that restrain their formation during S phase are incompletely understood. Here we show in vivo how R-loop formation drives a short purine-rich repeat, (GAA)10, to become a replication impediment that requires the repriming activity of the primase-polymerase PrimPol for its processive replication. Further, we show that loss of PrimPol results in a significant increase in R-loop formation around the repeat during S phase. We extend this observation by showing that PrimPol suppresses R-loop formation in genes harbouring secondary structure-forming sequences, exemplified by G quadruplex and H-DNA motifs, across the genome in both avian and human cells. Thus, R-loops promote the creation of replication blocks at susceptible sequences, while PrimPol-dependent repriming limits the extent of unscheduled R-loop formation at these sequences, mitigating their impact on replication.


Introduction
Tracts of repetitive sequence, known as microsatellites or short tandem repeats, occur frequently in vertebrate genomes (Clark et al., 2006, Tripathi & Brahmachari, 1991, Willems et al., 2014. Many such sequences are capable of forming secondary structures, including hairpins, cruciforms, triplexes (H-DNA) and G-quadruplexes (G4s), that have the potential to impede DNA replication (Mirkin & Mirkin, 2007). However, the factors that determine whether these sequences pose a barrier to DNA synthesis in vivo and the consequences of their doing so are not well understood.
It is well established that long repeats lead to problems with both replication and transcription. For example, a long tract of polypurine-polypyrimidine (GAA) n repeats (in which n can exceed 1500) is linked to the inherited neurodegenerative disorder Friedreich's ataxia (Campuzano et al., 1996). These repeats can form H-DNA (Frank-Kamenetskii & Mirkin, 1995) and are also prone to form R-loops (Groh et al., 2014), three-stranded nucleic acid structures in which nascent RNA hybridises to its complementary DNA template, displacing the non-template DNA strand (Thomas et al., 1976). These secondary structures block replication both in bacterial and yeast models, as well as in human cells (Chandok et al., 2012, Krasilnikova & Mirkin, 2004, Ohshima et al., 1998, which promotes genetic instability of the repeat (Gerhardt et al., 2016). They also perturb transcription by reducing RNA polymerase II (RNAPII) elongation (Bidichandani et al., 1998, Punga & Buhler, 2010 and lead to deposition of repressive chromatin marks (Al-Mahdawi et al., 2008, Saveliev et al., 2003. In the case of Friedreich's ataxia this results in transcriptional silencing of Frataxin (FXN), in which the repeat is located.
Less clear is the impact of the much more common short repetitive tracts found throughout the genome (Clark et al., 2006, Willems et al., 2014. These have generally not been linked to any significant impact on replication or transcription. For example, the (GAA) n repeat in normal alleles of FXN (n < 12) is not at risk of expansion (Schulz et al., 2009), despite the ability of even (GAA) 9 to form a stable H-DNA structure at physiological pH in vitro (Potaman et al., 2004). Further, these 'normal' repeats also induce significantly less R-loop formation than disease-length alleles and are not associated with delay of RNAPII or transcriptional silencing (Groh et al., 2014). However, it remains unclear whether this apparently inert behaviour is due to these sequences being incapable of forming secondary structures or whether it is the result of activities that counter structure formation and its consequences.
In this paper, we address these questions by studying the replication of a short GAA repeat in and G4-motifs throughout the genome. These results provide a direct demonstration that Rloop formation can promote DNA sequences with structure forming potential to become replication impediments and that repriming by PrimPol, in turn, limits excessive R-loop accumulation in the vicinity of these sequences, we suggest by preventing excessive exposure of single stranded DNA during their replication.

Instability of BU-1 expression monitors replication delay at (GAA)n
We have previously shown that expression instability of the BU-1 locus in chicken DT40 cells provides a sensitive readout for replication delay at G4 motifs (Schiavone et al., 2014).
The wild type locus contains a G4 motif 3.5 kb downstream of the TSS (the +3.5 G4) towards the end of the second intron ( Fig 1A), which is responsible for stochastic, replicationdependent BU-1 expression instability under conditions in which G4 replication is impaired (Guilbaud et al., 2017, Sarkies et al., 2012, Schiavone et al., 2014. Failure to maintain processive replication through the +3.5 G4 motif leads to uncoupling of DNA unwinding and DNA synthesis, interrupting normal histone recycling at the fork and the accurate propagation of epigenetic information carried by post-translational modifications on histone proteins ( Fig 1A). This leads to replication-dependent instability of BU-1 expression manifested as stochastic, replication-dependent conversion of the normal 'high' expression state to a lower expression level (Sarkies et al., 2012, Schiavone et al., 2014. This expression instability can be readily monitored by flow cytometric analysis of surface Bu-1 protein (Sarkies et al., 2012) (Fig EV1), providing a simple method to cumulatively 'record' episodes of interrupted DNA synthesis at the secondary structure-forming motif.
To model the replication of (GAA) n repeats we started with DT40 cells in which the +3.5 G4 motif of BU-1 had been deleted on both the A and B alleles (Schiavone et al., 2014). (GAA) n repeats of lengths between n = 10 and n = 75 were constructed either by synthesis for n ≤ 30 or, for the longer tracts, using a cloning strategy for highly repetitive sequences ( Fig EV2).
The reduced expression in cells carrying (GAA) 30-75 affects the entire population and thus appears to be distinct from the stochastic, replication dependent loss of expression we have previously reported to be induced by G4 motifs in cells lacking enzymes involved in G4 replication (Sarkies et al., 2012, Sarkies et al., 2010, Schiavone et al., 2014. Rather, these longer repetitive tracts resulted in the accumulation of chromatin-associated nascent RNA (ChrRNA) (Nojima et al., 2016) within the locus (Fig EV3a), consistent with impaired expression being due to reduced processivity of RNA polymerase II. As the global reduction of BU-1 expression in (GAA) 30-75 alleles precluded the analysis of stochastically generated loss variants, we focussed our subsequent analyses on (GAA) 10 and (GAA) 20 .
Fluctuation analysis for the generation of Bu-1a loss variants confirmed that the presence of (GAA) 10 at the +3.5 kb position did not affect the stability of BU-1 expression in a wild type background (Fig 1c). However, (GAA) 20 induced modest, but significant, formation of Bu-1a loss variants (Fig 1c) suggesting that this repeat is able to impede replication even in wild type conditions. We next examined the effect of deleting PrimPol to examine the extent to which repriming mitigates the replication impediment posed by these sequences. The results were striking: the rate at which Bu-1a loss variants were generated in primpol cells increased significantly, both for (GAA) 10 and (GAA) 20 (Fig 1c). These observations are consistent with repriming preventing significant uncoupling of DNA unwinding and DNA synthesis at these short repeats. They also suggest that these sequences are forming frequent impediments to otherwise unperturbed DNA replication.
The BU-1 instability seen in primpol cells carrying (GAA) 10 is comparable to that observed in primpol cells harbouring the endogenous +3.5 G4 . Similar Bu-1a medium and Bu-1a low expression states, characterised by loss of H3K4me3 and additional DNA methylation respectively, were isolated (Fig EV3b-e). Genetic instability, at a level that could explain the formation of Bu-1a loss variants, was not detected (Fig EV3f). Together, these observations suggest that the (GAA) 10 sequence causes epigenetic instability through the same replication-dependent mechanism that we have previously described for G4s. The effect of the repeat was orientation dependent (Fig 1d), only producing instability when knocked-in such that the purine-rich strand formed the leading strand template for a fork entering from the 3' end of the locus.
The RPA-binding and repriming functions of PrimPol are required to ensure processive replication at the BU-1 (GAA) 10 repeat.
While PrimPol can perform some translesion synthesis, considerable evidence now supports repriming as its main in vivo role (Keen et al., 2014, Kobayashi et al., 2016, Mouron et al., 2013. The repriming function of PrimPol requires the C-terminal zinc finger and RPA binding motif A (RBM-A), which mediates an interaction with the singlestranded binding protein replication protein A (Guilliam et al., 2017, Guilliam et al., 2015, Wan et al., 2013. To confirm that the primary in vivo role played by PrimPol in the replication of (GAA) 10 is indeed repriming, we performed a complementation study by ectopically expressing YFP-tagged human PrimPol in primpol cells (Fig 2). Expression of full-length human PrimPol completely restored the stability of BU-1A expression in primpol cells carrying the (GAA) 10 repeat. Expression of catalytically inactive PrimPol (D114A, E116A or hPrimPol[AxA]) and the DNA-binding zinc finger mutant (C419A, H426A or hPrimPol[ZfKO]) shows that neither is able to prevent instability of BU-1 expression (Fig 2), a result concordant with our previous observation in cells harbouring the natural +3.5 G4.
Importantly, RBM-A, but not RBM-B, of PrimPol is essential for efficient suppression of (GAA) 10 repeat-induced instability of BU-1 expression, consistent with previous work showing that while both RBM-A and RBM-B can bind the same basic cleft in RPA70N in vitro, only RBM-A appears important in vivo (Guilliam et al., 2017). REV1, a Y-family DNA polymerase, is required for maintaining stability of BU-1 expression when the sequence at the +3.5 kb position is a G4 motif (Sarkies et al., 2012, Schiavone et al., 2014, which reflects a direct role for REV1 in G4 replication (Eddy et al., 2014, Sarkies et al., 2010. Replacing the +3.5 G4 in rev1 cells with (GAA) 10 repeats did not result in significant destabilisation of BU-1 expression (Fig EV4a), while (GAA) 20 results in a modest destabilisation of BU-1 expression, comparable to the effect in wild type cells. This suggests that the role REV1 plays in maintaining epigenetic stability of BU-1 is specific to G4 motifs in contrast to PrimPol, which is involved in replicating both types of secondary structure through its ability to reprime.

PrimPol limits R-loop formation around a (GAA) 10 repeat
The orientation dependence of the GAA tract with respect to BU-1 instability is in line with the predicted formation of triplex DNA when a polypurine tract is transcribed as the coding strand, but not as the template strand (Grabczyk & Fishman, 1995, Grabczyk et al., 2007. In vitro studies have shown that the formation of triplexes at (GAA) n occurs concurrently with the formation of a stable DNA:RNA hybrid between the TTC-rich template strand and the nascent GAA-containing RNA strand (Grabczyk et al., 2007). Furthermore, pathological formation of R-loops has been reported at long (GAA) n repeats (n ≥ 650) in immortalised lymphoblasts derived from Friedreich's ataxia patients (Groh et al., 2014). These reports, together with the results presented thus far, prompted us to investigate whether R-loops contribute to replication stalling induced by short GAA tracts in BU-1.
R-loops can be detected using a DNA:RNA hybrid-specific antibody, S9.6 (Boguslawski et al., 1986). We first examined R-loop formation in the (GAA) 10 -containing BU-1 locus of wild type cells using DNA:RNA immunoprecipitation (DRIP) followed by quantitative PCR (Fig   3a). The first striking feature to note is the strong signal in the vicinity of the transcription termination site (Fig 3a), consistent with the previously described R-loops formed in this region in a subset of genes that are mechanistically associated with transcription termination (Skourti-Stathaki et al., 2014). Interestingly, this signal is consistently higher in primpol cells compared with wild type, although we do not have a clear explanation for this observation at present. In the body of BU-1 in wild type, the presence of (GAA) 10 at the + 3.5kb position results in a very modest DRIP signal in the vicinity of the repeat. In contrast, primpol cells exhibit a highly significant increase in R-loop signal around the same repeat. Crucially, this enrichment is abrogated when the repeat is deleted, showing that R-loop accumulation is not simply a direct consequence of loss of PrimPol, but arises due to failure of repriming at the replication impediment created by (GAA) 10 . This is unlikely to reflect a direct role of PrimPol in processing R-loops as primpol cells are not sensitive to camptothecin, a topoisomerase I inhibitor that leads to increased R-loop formation ( Fig EV5) (Kobayashi et al., 2016).

R-loop formation is required for (GAA) 10 to induce expression instability of BU-1 in primpoldeficient cells
We next asked whether formation of a replication block at (GAA) 10 requires an R-loop. To address this, we overexpressed chicken RNaseH1, an enzyme that degrades R-loops (Stein & Hausen, 1969) and that we have previously shown can be stably expressed and active in DT40 cells . YFP-tagged chicken RNaseH1 carrying a disrupted mitochondrial localisation sequence was stably expressed in primpol BU-1A (GAA)10 cells (Fig  3b). Expression of this construct reduced the R-loop signal in the vicinity of the repeat (+3 kb) to a level comparable with the background signal (Fig 3c).
We next performed fluctuation analysis for the formation of Bu-1a loss variants in the RNaseH1-overexpressing primpol BU-1A (GAA)10 cells. Strikingly, RNaseH1 completely prevented the formation of Bu-1a loss variants in three separate clones, an effect comparable to removing the (GAA) 10 repeat itself (Fig 3d). This suggests that DNA:RNA hybrid formation makes a crucial contribution to the ability of (GAA) 10 to induce BU-1 expression instability.
Since the activity of PrimPol is intimately linked with replication, we hypothesised that specifically removing the R-loops in S phase would suppress (GAA) 10 -induced BU-1 expression instability. We therefore expressed chicken RNaseH1 fused to a degron sequence from geminin, which ensures protein expression is restricted to S phase (Sakaue-Sawano et al., 2008) (Fig 3e). Expression of this construct was able to prevent instability of BU-1 expression in primpol BU-1A (GAA)10 (Fig 3f), confirming that R-loops present during S phase are indeed responsible for the (GAA) 10 -dependent destabilisation of BU-1. Together with the DRIP experiments, this result suggests that R-loop formation is not only a consequence of the (GAA) 10 repeat but is necessary for it to become a replication impediment.

R-loop stabilisation converts the (GAA) 10 sequence into a replication impediment
The R-loop dependence of BU-1 expression instability in primpol mutants led us to the prediction that the enforced stabilisation of R-loops might lead the (GAA) 10 repeat to induce BU-1 expression instability even in wild type cells. To achieve this, we overexpressed the 52 amino acid DNA:RNA hybrid binding domain (HBD) of human RNaseH1, a fragment previously shown by the Aguilera lab to co-localise with and stabilise DNA:RNA hybrids in vivo (Bhatia et al., 2014). We cloned the HBD in frame with mCherry separated by a flexible GSGSG linker. The resulting fusion protein could be stably expressed in DT40 cells as monitored by mCherry fluorescence and by Western blotting (Fig 4a & b). Expression of the HBD in cells lacking a structure-forming sequence at the +3.5 kb position of BU-1A (DT40 BU-1A ∆G4 ) did not induce statistically significant destabilisation of BU-1 expression compared with the control (Fig 4c). However, when the (GAA) 10 repeat was present at the +3.5kb position we observed significantly greater expression instability. This observation provides further evidence that R-loops are causal in promoting a (GAA) 10 motif to become a replication block.

PrimPol curtails R-loop formation during S phase
As the activity of PrimPol is linked with replication, we next asked if the accumulation of Rloops at the BU-1 locus of primpol cells (Fig 3a) occurs during S phase. Wild type BU-1A (GAA)10 and primpol BU-1A (GAA)10 cells were synchronised in G1 by double thymidine block, with cells at different phases of S phase sampled upon release ( Fig EV6a) and steady-state Rloop dynamics monitored using DRIP (Fig 5a). In wild type cells, the highest levels of DNA:RNA hybrids occurred in G1 and G2 phases, most likely associated with transcription, while they remained relatively sparce during S phase. In contrast, R-loops in BU-1 of primpol cells were significantly elevated at the early stages of S phase relative to wild type. Using the published replication timing map for BU-1 (Schiavone et al., 2014) we estimate that the locus is replicated within 2 hours of entry into S phase, which corresponds to the time points at which we observe an increase in R-loop signal.
To explore whether PrimPol suppresses R-loop formation in S phase, we used metabolic labelling of nascent RNA with 4-thiouridine (4-sU) to examine active DNA:RNA hybrid formation in different phases of the cell cycle. DNA containing R-loops was gently extracted, and free cytoplasmic and nuclear RNA was degraded with RNaseA in high salt leaving behind the RNA moiety of R-loops. Surviving 4-sU-labelled RNA was biotinylated and its relative abundance analysed by immunoblotting with streptavidin-HRP ( Fig EV6b). Using this analysis, which is independent of the S9.6 antibody, we observed that, in wild type cells, the majority of R-loop formation is seen in the G1 and G2 periods of the cell cycle when cotranscriptional R-loop formation would be expected (Fig 5b). Interestingly, R-loop formation also increases transiently in early S phase, but then diminishes. In contrast, primpol cells exhibit a sustained DNA:RNA hybrid signal throughout S phase. Thus, the increased R-loops we observe in primpol cells result from unscheduled DNA:RNA hybrid formation during S phase. Extending from the experiments showing that R-loop accumulation in BU-1 is potentiated by a repriming defect (Fig 3a), the unscheduled R-loop formation in S phase can be interpreted as the consequence of failure to restrict the length of single stranded DNA gaps produced by interruptions of DNA polymerisation as S phase progresses.

PrimPol suppresses R-loop formation in the vicinity of secondary structure-forming sequences throughout the genome
To explore whether our observations at BU-1 could be extended to the whole genome, we performed quantitative high-throughput sequencing of S9.6 immunoprecipitated DNA (DRIP-Seq) from wild type and primpol DT40 cells. To allow quantitation of the DRIP signal the DT40 cells were spiked with a fixed proportion of Drosophila S2 cells to provide an internal control (Orlando et al., 2014). Following read alignment and peak calling, peaks in the wild type and primpol samples were normalised to the mean number of Drosophila reads.
This revealed a highly significant increase in the height of the DRIP peaks in primpol cells (Fig 6a). Between wild type and primpol, 84% of peaks were shared suggesting that the loss of PrimPol does not result in the appearance of new peaks, but that for any given peak there is a greater DRIP signal, suggesting a higher steady-state level of R-loops (Fig 6b). 41% of DRIP peaks overlapped with genes and 83% of genes with DRIP peaks were shared between wild type and primpol. We next asked if genes with DRIP peaks are enriched for H-DNA motifs. Using the 'Triplex' R package (Hon et al., 2013), sequences with H-DNA-forming potential were identified as overlapping just under 8% of all genes. The subset of genes harbouring DRIP peaks was significantly enriched for these sequences, with c. 15% of these genes overlapping sequences with H-DNA-forming potential (Fig 6c). Within this set of genes there was a significant increase in peak height in the primpol mutant (Fig 6d).
Our previous work has shown that G4s are able to induce similar epigenetic instability to the (GAA) 10 repeat that has been the focus of this study. Further, G4 motifs have been linked to R-loop formation (Duquette et al., 2004). We therefore used the expression [G 3-5 N 1-7 G 3-5 N 1-7 G 3-5 N 1-7 G 3-5 ] (Huppert & Balasubramanian, 2005) to identify G4 motifs in the chicken genome. G4 motifs were identified to overlap with 59% of all genes, while 76% of genes with DRIP peaks overlapped motifs, a significant enrichment (Fig 6e) and, again, the heights of the peaks in the primpol data set were significantly increased (Fig 6f).
The striking increase in steady state R-loop accumulation in genes containing G4 motifs and our previous demonstration that G4 motifs also potently destabilise BU-1 expression in primpol cells  prompted us to ask if R-loops also contribute to promoting G4 motifs to become replication impediments. Forcing RNaseH1 expression resulted in a highly significant reduction in BU-1 instability induced by the natural +3.5 G4 in four separate RNaseH1-expressing clones (Fig 6g), demonstrating that R-loop formation increases the probability of this G4 forming a significant replication impediment, but that it is not essential for it to do so.
Finally, we asked whether loss of PrimPol affected R-loop levels in human cells. PrimPol was disrupted using CRISPR/Cas9 editing in the induced pluripotent stem cell line BOBSC (Yusa et al., 2011). Genome wide R-loops were isolated with S9.6 immunoprecipitation followed by a modified sequencing protocol, RNA-DIP, that monitors the RNA moiety within the DNA:RNA hybrids. The analysis, normalised by spiking with DT40 cells, revealed a very similar picture to that seen in DT40. A highly significant increase in peak height in the primpol mutant (Fig 6h) reflected an increase in the RNA-DIP signal at existing peaks, with 66% of wild type and primpol peaks overlapping (Fig 6i). As in DT40, the genes containing peaks were associated with H-DNA and G4 motifs (Fig 6j & l) but in both cases the mean peak height was significantly higher in primpol cells (Fig 6k & m) demonstrating that loss of PrimPol also increases R-loop formation in the vicinity of DNA secondary structures in human cells.

A requirement for PrimPol reveals that (GAA) 10 forms a replication impediment
The (GAA) 10 repeat upon which this study has focussed is typical of widespread short tandem repeats found throughout vertebrate genomes (Willems et al., 2014). Repeats of this length have not been previously linked to detectable disturbances in replication or transcription in vitro (Bidichandani et al., 1998, Ohshima et al., 1998 despite their potential to form triplex structures at physiological pH (Potaman et al., 2004). Our previous work supports a model in which instability of BU-1 expression induced by G4s results from uncoupling of DNA unwinding from leading strand DNA synthesis (Sarkies et al., 2010, Schiavone et al., 2014, Šviković & Sale, 2017, and have shown that this uncoupling can be reduced by repriming . We now show that the repriming function of PrimPol is frequently deployed at short tandem repeats present throughout the genome as exemplified by the engineered (GAA) 10 repeat. Consequently, loss of PrimPol causes widespread replication defects (Bianchi et al., 2013, Mouron et al., 2013.

The nature of the replication impediment formed by (GAA) 10
(GAA) n repeats, in common with other polypurine-polypyrimidine tracts, are capable of forming triplex secondary structures in which a third strand anneals through Hoogsteen base pairing. This tendency has been linked to the detrimental effect of long GAA repeats on transcription elongation (Bidichandani et al., 1998, Punga & Buhler, 2010 through the trapping of transcribing RNA polymerase II (Grabczyk & Fishman, 1995). However, the very act of transcription also promotes formation of secondary structures (Kouzine et al., 2017, Lilley, 1980, including triplexes (Grabczyk & Fishman, 1995, Kouzine et al., 2004. This is likely driven by structure formation releasing the negative supercoiling generated in the wake of translocating RNA polymerase (Levens et al., 2016, Liu & Wang, 1987. Similar topologyinduced structure formation could also contribute to leading strand secondary structure formation behind the replicative helicase (reviewed in Kurth et al., 2013, Yu & Droge, 2014. We show that (GAA) 10 requires an RNaseH1-sensitive R-loop in order to create a replication impediment that requires PrimPol-dependent repriming. It is well established that formation of DNA:RNA hybrids coincides with the sequences able to adopt non-B DNA (Duquette et al., 2004, Grabczyk et al., 2007 and is favoured in a negatively supercoiled DNA template (Roy et al., 2010). R-loops have been implicated as a major factor in the severity of head-on collisions between the replication and the transcriptional machinery (Hamperl et al., 2017).
However, a direct head-on collision with transcribing RNA polymerase is likely to halt the entire replisome (Pomerantz & O'Donnell, 2010), precluding the displacement of parental nucleosomes caused by the functional uncoupling between the replicative helicase and DNA synthesis. It is difficult to reconcile this type of stall with the involvement of PrimPol.
Specifically, the DNA and RPA binding activities of the enzyme suggests the transient formation of ssDNA, which most likely arises as a result of functional uncoupling of replicative helicase from the replicative polymerases, and which is the basis for BU-1 expression instability.
How then can a (GAA) n repeat generate the uncoupling of DNA unwinding and leading strand DNA synthesis necessary to induce transcriptional instability of BU-1? We propose that transcription of the (GAA) 10 repeat generates an R-loop (Fig 7). During replication, the approaching replicative helicase traverses the transcription complex by displacing the RNA polymerase (Pomerantz & O'Donnell, 2010) or by reorganising the helicase itself , Vijayraghavan et al., 2016. Biophysical calculations show that DNA:RNA hybrids are sufficiently thermodynamically stable to survive the accumulation of positive supercoiling generated ahead of the replicative helicase (Belotserkovskii et al., 2013). Since the eukaryotic replicative helicase tracks on the leading strand (Douglas et al., 2018), we suggest that the DNA:RNA hybrid could remain intact on the lagging strand during passage of the helicase. Behind the replicative helicase, the persistent lagging strand DNA:RNA hybrid may retrap the purine-rich leading strand through triplex formation. The resulting R:R•Y triplex would then block leading strand synthesis (Samadashwily & Mirkin, 1994).
This model is consistent with the observation that the depletion of DNA:RNA hybrids through overexpression of RNaseH1 completely abolishes (GAA) 10 -dependent BU-1 expression instability in PrimPol-deficient cells. An alternative explanation for the creation of a leading strand impediment is the formation of a DNA triplex stabilised by an adjacent DNA:RNA hybrid, of the form proposed by Grabczyk and Fishman (1995). In either event, continued helicase activity would result in exposure of ssDNA ahead of the stalled replicative polymerase, which through being bound by RPA promotes the recruitment of PrimPol.
Repriming then allows DNA synthesis to remain coupled to unwinding leaving the triplex in a small gap to be disassembled post-replicatively (Fig 7).

Loss of PrimPol-mediated repriming at structured DNA promotes S phase R-loop accumulation
In the absence of PrimPol, continued unwinding of the parental duplex by the replicative helicase in the context of a continued stalling of DNA synthesis would create a more extensive region of ssDNA. That this results in a net increase in DNA:RNA hybrids in the vicinity of structure-forming sequences (Fig 3, 5b & 6) suggests that there is greater ssDNA exposure on the lagging, as well as the leading strand, which may explain the increase in nascent transcription during S phase (Fig 5a). These observations imply that RNAPII continues to transcribe despite its template remaining single stranded. This idea is consistent with both biochemical (Kadesch & Chamberlin, 1982) and in vivo reports (Michelini et al., 2017, Ohle et al., 2016 demonstrating that DNA:RNA hybrid formation can occur at ssDNA generated by resection of DNA ends at double-stranded DNA breaks. We suggest that ssDNA generated as a consequence of helicase-polymerase uncoupling, unmitigated by repriming, could also act as a substrate for unscheduled RNAPII transcription and DNA:RNA hybrid formation (Fig 7).
The results we present here establish two important mechanistic points concerning the relationship between R-loops and impeded replication. First, we show that R-loops are able to promote short sequences with structure-forming potential to become replication impediments, requiring the repriming activity of PrimPol to maintain their processive replication. Second, failure to reprime at these sequences increases R-loop formation. We suggest this is due to failure to prevent the exposure of excessive single stranded DNA during S phase. This relationship between structure-forming DNA and repriming is seen at a significant fraction of R-loops detected across the genome, suggesting that repriming plays an important role in allowing cells to manage the complex challenges created by the interaction of transcription and replication.

CRISPR-Cas9 mediated gene disruption in human cells
Guide RNA sequences used for disrupting PRIMPOL in BOBSC iPS cell lines are listed in the Appendix. Each gRNA sequence was cloned into pX458 (Ran et al., 2013). A targeting construct carrying puromycin selection marker was constructed by Gibson assembly using PCR amplified 5' and 3' homology arms (primer sequences in the Appendix). Equimolar amounts of targeting construct, gRNA expression vectors and Cas9 expression vectors were delivered by Amaxa electroporation. Puromycin resistant clones were genotyped by PCR and Sanger sequencing (Shen et al., 2014).

Molecular cloning and transgene constructs
For transgene expression, cDNA was cloned in frame with fluorescent protein (mCherry or YFP) in the polylinker of pXPSN2 (Ross et al., 2005). The expression module was released with SpeI digestion and subcloned into pBluescript based vectors containing a loxP flanked puromycin or blasticidin S selection cassettes (Arakawa et al., 2001), which were transfected into DT40 via electroporation. Primers used for molecular cloning are listed in the Appendix.
Enrichment was normalized to H3 and reported relative to wild-type.

Cell cycle synchronization and 2D cell cycle analysis
G1 phase synchronization of DT40 cells was achieved by double thymidine block. Cells were treated overnight with 2 mM thymidine, released for 9 hours, and again treated with thymidine overnight, after which cells were released into medium containing 0.2 µM nocodazole. Upon release, cells in different cell cycle phases were harvested to be analysed or pulse labeled with BrdU. 5 -10 x10 6 DT40 cells were pulse labelled with 50 μM BrdU for 30 minutes in complete medium at 37 °C. BrdU staining was performed as previously described (Frey et al., 2014).

4-thiouridine (4sU) metabolic labelling of nascent DNA:RNA hybrids
4-thiouridine was used to label nascent RNA during different stages of cell cycle. 75-100 million DT40 cells were resuspended in 10 ml of complete medium supplemented with 100 µM 4sU and incubated for 30 minutes at 37 ºC. 4sU incorporation was terminated by adding ice cold PBS, after which cells were lysed in TE buffer supplemented with 1% SDS and 0.2 mg/ml Proteinase K. Nucleic acids were purified with phenol:chloroform:isoamyl alcohol and digested with KpnI, EcoRI and BamHI overnight at 37°C, followed by treatment with 100 μg RNaseA in 0.5 M NaCl. RNaseA in high-salt inactivates its RNaseH-activity (Halasz et al., 2017). DNA was purified again with phenol:chloroform and resuspended in 10 mM Tris-HCl pH 8.0. 4sU-labelled RNA moiety of DNA:RNA hybrids were biotinylated with thiol-specific reaction using EZ-Link HPDP-Biotin (Thermo Scientific) as described previously (Fuchs et al., 2015). Free biotin was removed by chloroform:isoamyl alcohol (24:1 v/v). Nucleic acids were bound to Hybond-XL membrane using slot blot apparatus, crosslinked and blotted with HRP-Conjugated Streptavidin (N100, Thermo Scientific). As DMF is a known DNA denaturation agent (Cortadas & Subirana, 1977, Herskovits et al., 1961, generated ssDNA was used as an internal loading control. Biotin was stripped for one hour in 100 mM dithiothreitol at room temperature, and the membrane incubated with anti-ssDNA antibody.

DRIP-Seq
Sample preparation for DRIP-Seq was essentially performed as described above, but with some minor changes. All the samples were spiked (Orlando et al., 2014) with the same batch of Drosophila S2 cells in 1:4.2 ratio to DT40 cells. Digested DNA was treated with 100 μg RNase A in the presence of 0.5 M NaCl for 2 h at 37ºC. Elution from magnetic beads was performed for 1 h at 37 ºC in 300 μl elution buffer supplemented with 0.1 mg/ml RNase A.
To ensure complete elution, 10 μg Proteinase K was added and incubated further 90 minutes at 37 ºC. DNA was purified by phenol:chloroform:isoamyl alcohol extraction, quantified with Qubit dsDNA HS Assay Kit (Invitrogen), diluted with ultra-pure water to 55 μl and sheared with Covaris M220 Focused-ultrasonicator and Holder XTU to average size of 300 bp in microTUBE-50 AFA. DNA libraries were built using NEBNext Ultra II DNA Library Prep Kit (New England Biolabs) as per the manufacturer's instructions.

RNA DIP-Seq
Between 60-100 million human cells were spiked in with DT40 cells (1/10 ratio), harvested, washed with cold PBS and nuclei isolated by lysing cells in HLB+N and passing it through a 10% sucrose cushion. Collected nuclei were lysed overnight in with NLB (25 mM Tris-HCl (pH 7.5), 1% SDS, 5 mM EDTA, 0.125 mg/ml Proteinase K) with agitation at 37°C. Nucleic acids were purified with 1 M potassium acetate (pH 5.5), precipitated and treated with of RNaseI (Ambion, AM2294) per 90 μg of DNA (15 minutes at 37°C) to degrade soluble RNA molecules. DNA was purified with phenol:chloroform, diluted with the IP dilution buffer and sheared with a Bioruptor plus (Diagenode) to average size of 300 bp. DNA:RNA hybrids were immunoprecipitated with S9.6 mAb (1 μg antibody for each 2 μg of DNA) overnight.
Immunocomplexes were captured with Protein G beads and washed as for ChIP and DRIP preparation. DNA:RNA hybrids were eluted by incubating the sample with Proteinase K for 2 hours at 42°C. Nucleic acids were cleaned up with phenol:chloroform:isoamylalchohol, precipitated with glycogen and resuspended in water, denatured for 5 minutes at 90°C and immediately placed on ice. DNA moiety of DNA:RNA hybrids were removed with 4U DNase I for 30 minutes at 37°C. RNA was extracted with QIAzol, precipitated overnight and dissolved in RNase-free water. Strand-specific Illumina-compatible libraries were prepared with NEBNext Ultra II Directional RNA Library Prep Kit (NEB, E7760) with 100 ng input.
Libraries were quality checked as before and sequenced on a NextSeq500 (Illumina).

Quantification and statistical analysis of deep sequencing data
DRIPseq libraries were sequenced on an Illumina HiSeq 4000, RNA-DIP libraries were sequenced on an Ilumina NextSeq. Reads were trimmed and quality filtered using trim galore (version 0.4.4) (https://www.bioinformatics.babraham.ac.uk/projects/trim_galore/), then aligned to genomes with bowtie2 (version 2.26) (Langmead & Salzberg, 2012) using default settings. DT40 reads were aligned to Ggal 5.0 and Dmel r6.18, BOBSC reads were aligned to GRCh38 and Ggal 5.0. Alignments were filtered for uniquely matching reads and separated into sample and spike-in. Peaks were called on filtered alignments using MACS2 (version 2.1.1.20160309) with the default settings and -g 1.87e9 (or -g hs for human) --broad (Feng et al., 2012). Peak heights were normalised to the read number of the spike-ins and compared using the Mann Whitney U test. Overlaps between peaks were calculated using bedtools2 closest (version 2.27.1) with default settings (Quinlan & Hall, 2010). Peaks were considered to be overlapping if at least 1bp overlapped. Sequences with H-DNA forming potential were identified with the Triplex R package (Hon et al., 2013). G4 motifs were identified using the Quadparser algorithm (Huppert & Balasubramanian, 2005) with the regex [G 3-5 N 1-7 G 3-5 N 1-7 G 3-5 N 1-7 G 3-5 ]. Enrichment testing for secondary structures was performed using the hypergeometric test.

Data Availability
Deep sequencing data has been deposited in the GEO repository

Declarations of interest
The authors declare that they have no conflicting interests with respect to this work.   For all panels, mean ± SD reported. **** p ≤ 0.0001, *** p ≤ 0.001, ** p ≤ 0.01, ns = not significant.