The Interplay of RNA:DNA Hybrid Structure and G-Quadruplexes Determines The

bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 The interplay of RNA:DNA hybrid structure and G-quadruplexes determines the

2 outcome of R-loop-replisome collisions

3 Charanya Kumar1, Sahil Batra1, Jack D. Griffith2, Dirk Remus1*

5 1Molecular Biology Program, Memorial Sloan Kettering Cancer Center, 1275 York Avenue, New York, NY 10065,

6 USA.

7 2Lineberger Comprehensive Cancer Center and Departments of Microbiology and Immunology, and

8 Biochemistry and Biophysics, University of North Carolina at Chapel Hill, Chapel Hill, NC, 27599, USA.

11 *Corresponding author: Molecular Biology Program, Memorial Sloan Kettering Cancer Center, 1275 York

12 Avenue, New York, NY 10065, USA. Tel: 212-639-5263; E-mail: [email protected]. ORCID:

13 https://orcid.org/0000-0002-5155-181X

1 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 ABSTRACT

2 R-loops are a major source of genome instability associated with transcription-induced replication

3 stress. However, how R-loops inherently impact replication fork progression is not understood. Here, we

4 characterize R-loop-replisome collisions using a fully reconstituted eukaryotic DNA replication system. We find

5 that RNA:DNA hybrids and G-quadruplexes at both co-directional and head-on R-loops can impact fork

6 progression by inducing fork stalling, uncoupling of leading strand synthesis from replisome progression, and

7 nascent strand gaps. RNase H1 and Pif1 suppress replication defects by resolving RNA:DNA hybrids and G-

8 quadruplexes, respectively. We also identify an intrinsic capacity of replisomes to maintain fork progression at

9 certain R-loops by unwinding RNA:DNA hybrids, repriming leading strand synthesis downstream of G-

10 quadruplexes, or utilizing R-loop transcripts to prime leading strand restart during co-directional R-loop-

11 replisome collisions. Collectively, the data demonstrates that the outcome of R-loop-replisome collisions is

12 modulated by R-loop structure, providing a mechanistic basis for the distinction of deleterious from non-

13 deleterious R-loops.

2 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 INTRODUCTION

2 Genome maintenance is dependent on the complete and accurate replication of the chromosomal DNA

3 prior to cell division. However, chromosomes present diverse obstacles to normal replication fork progression,

4 including protein-DNA complexes, DNA damage, and non-B-form DNA secondary structures. Among these, R-

5 loops have emerged as critical determinants of transcription-replication conflict (TRC) linked to genome

6 instability in human developmental disorders and disease 1,2. How R-loops inherently impact fork progression is

7 not understood.

8 R-loops are three-stranded nucleic acid structures that are formed co-transcriptionally when nascent

9 RNA anneals to the DNA template, displacing the non-template strand as single-stranded DNA (ssDNA). Some

10 R-loops serve important physiological roles by promoting chromosome segregation 3, transcription termination

11 4, Ig class switch recombination 5, telomere homeostasis 6, or gene expression control 7. In contrast,

12 unscheduled R-loops can induce DNA breaks that threaten genome stability 1,2. R-loop-induced DNA damage

13 occurs predominantly in S phase, consistent with R-loops exacerbating TRC 8-12. However, for reasons that are

14 unknown, only a subset of unscheduled R-loops appears to cause DNA damage 13. In part, this may be due to

15 head-on (HO) TRC being more detrimental than co-directional (CD) TRC. Yet, collisions in either orientation

16 induce a DNA damage response in human cells 14. This raises the question what distinguishes harmful from

17 harmless R-loops.

18 R-loops are detected at 5 % and 8 % of the human and yeast genomes, respectively 15,16. However,

19 under normal conditions R-loops form transiently, limiting their potential to interfere with DNA replication 15 17.

20 This dynamism is a consequence of the action of R-loop resolving enzymes, such as RNase H that degrades

21 RNA in RNA:DNA hybrids and various helicases that may unwind RNA:DNA hybrids 1,2. Consequently, while

22 R-loops form also under normal conditions 18-20, mutations in R-loop resolving enzymes dramatically increase

23 cellular R-loop levels and aggravate genome instability 1,2. In addition, co-transcriptional RNA processing and

24 export factors, and topoisomerases, limit R-loop formation 10,21,22. Thus, cells employ a host of strategies to

25 protect the genome from deleterious R-loops.

26 Cellular R-loops range in size from hundreds to thousands of base pairs 16,23,24. In general, R-loop

27 formation correlates with high gene activity, GC-richness, and G/C skew 1,2. The prevalence of G/C skew at R-

28 loops is partly explained by the enhanced stability of RNA:DNA hybrids featuring a G-rich RNA strand 25. In 3 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 addition, stretches of G on the displaced non-template strand promote the formation of G-quadruplexes (G4s)

2 that stabilize R-loops 26,27. G4s are DNA secondary structures formed by stacks of G-quartets in which four

3 guanine bases form Hoogsteen base pairs in a planar ring configuration 28. G4 sequences can interfere with

4 DNA replication, causing genetic or epigenetic instability 29-31. The mechanisms involved are not clear, but G4

5 sequences on either the leading or the lagging strand template have been reported to interfere with normal

6 DNA replication 31-33. As for RNA:DNA hybrids, a host of helicases has been implicated in promoting DNA

7 replication through G4 sequences 34. How G4s may modulate the impact of R-loops on DNA replication is not

8 known.

9 Cellular studies are limited in their ability to differentiate direct and indirect effects of R-loops. The latter

10 may include enhanced RNAP stalling 35, establishment of repressive chromatin structures 23,36,37, or induction

11 of DNA breaks by nucleases 38,39. To overcome this limitation, here we employ the reconstituted budding yeast

12 DNA replication system in combination with purified R-loop templates to study orientation-specific R-loop-

13 replisome collisions 40-42. We demonstrate that both CD and HO R-loops can adversely affect fork progression,

14 dependent on the specific configuration of RNA:DNA hybrids and G4s. RNA:DNA hybrid- and G4-induced

15 defects can be resolved by RNase H1 and Pif1, respectively. Unexpectedly, we find that leading strand

16 synthesis can be reprimed by distinct mechanisms downstream of G4s and RNA:DNA hybrids, promoting

17 continued fork progression at R-loops. Collectively, our data reveals how the specific structure of R-loops

18 determines the outcome of R-loop-dependent TRC.

20 RESULTS

22 Preparation and characterization of R-loop-containing DNA templates

23 To generate DNA templates harboring R-loops of characteristic length and nucleotide composition we

24 inserted a 1.4 kbp fragment derived from the R-loop-forming Airn locus under control of a T7 promoter into

25 yeast replication origin-containing plasmids (Figure 1A) 20,43. Studies in human cells have demonstrated that

26 R-loops formed at this sequence impede normal fork progression 14. Both the template and non-template

27 strand harbor several G4 sequences, but due to the G/C skew the G4 potential is increased on the non-

28 template strand (Figure 1A and Figure S1A). To reconstitute orientation-specific collisions of replisomes with 4 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 R-loops we inserted the R-loop element in both orientations relative to the replication origin, ARS305. In either

2 orientation, the C-rich strand serves as the template for transcription. Linearization of the plasmid template by

3 restriction digestion prior to origin firing ensures that only one of the two replication forks emanating from the

4 origin encounters the R-loop, while the other fork runs off the opposite template end (Figure 1B).

5 In order to promote R-loop formation in vitro, transcription is carried out on negatively supercoiled

6 plasmid templates 44,45. Moreover, to avoid RNA discontinuities due to repeated transcription initiation, purified

7 T7 lysozyme, which inhibits transcription initiation but not elongation by T7 RNAP 46,47, is added late in

8 transcription reactions (Figure 1C). Subsequently, RNase A is used to digest free RNA, which facilitates the

9 separation of the template from excess free RNA by size-exclusion chromatography (Figure S1B+C). Reaction

10 mixtures are deproteinized to remove T7 RNAP and plasmid templates are purified by gel-filtration (Figure

11 S1C). R-loop formation under these conditions is efficient as demonstrated by the RNase H-sensitive gel-

12 electrophoretic mobility shift of the template DNA (Figure 1D).

13 Analysis of the R-loop-associated RNA by denaturing formaldehyde agarose gel-electrophoresis

14 reveals that most RNA molecules are significantly shorter than 1.4 kb, ranging in size between ~200-600 nt

15 (Figure 1E). Thus, only sections of the Airn sequence form R-loops in vitro, which is consistent with previous

16 single-molecule R-loop sequencing and footprinting data 43. To characterize the R-loop-containing templates

17 we employed electron-microscopy (EM) after rotary shadow casting (Figure 1F). In this approach, R-loop

18 positions can be mapped relative to the template ends. We employed three approaches to image R-loops by

19 EM (Figure 1F). To visualize the displaced non-template strand, DNA templates were incubated with single-

20 stranded DNA binding protein, T4 gp32. This approach reveals bubble-like R-loop structures of variable sizes

21 specifically at the Airn sequence. Relevant to the analyses below, R-loops form preferentially in the 5’ half of

22 the transcribed sequence, resulting in their center of distribution being closer to the replication origin in the CD

23 than in the HO orientation (Figure 1G). To probe for RNA:DNA hybrids, templates were incubated with the

24 RNA:DNA hybrid-specific antibody, S9.6 (Figure 1F) 48. This analysis reveals the formation of unique electron-

25 dense structures at the position of the Airn sequence, confirming the presence of RNA:DNA hybrids at this site.

26 Finally, DNA templates were spread on EM grids under mildly denaturing conditions in the presence of

27 formamide (Figure 1F). This analysis reveals a characteristic difference in the thickness of the two arms of the

28 R-loops, consistent with one arm corresponding to the single-stranded non-template strand and the other 5 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 corresponding to the RNA:DNA duplex. As in the analysis with T4 gp32, and consistent with the gel analysis in

2 Figure 1E, R-loops observed in the presence of formamide exhibit a range of sizes, demonstrating that the

3 size variations are not due to the sample preparation method. This is further confirmed by similar observations

4 using atomic force microscopy (AFM) (Figure S1D). In conclusion, the data demonstrates efficient formation of

5 canonical R-loop structures of variable size and position at the Airn sequence in vitro.

7 Both CD and HO R-loops perturb normal fork progression

8 To test the impact of R-loops on fork progression, DNA templates are linearized with NsiI prior to origin

9 firing. Replication products are digested with ClaI prior to gel analysis to reduce heterogeneity in their gel-

10 mobility caused by the distributive initiation of strand synthesis at the origin (Figure 2A+B)49. In the absence of

11 R-loops, the Airn sequence does not present a notable obstacle to fork progression in either orientation. In

12 contrast, CD or HO R-loops induce a striking loss of full-length replication products and the appearance of new

13 replication intermediates. Native gel analysis reveals reduced levels of full-length linear daughter molecules in

14 the presence of either CD or HO R-loops, as well as the appearance of stalled forks and partially replicated

15 daughter molecules resulting from the uncoupling of DNA unwinding from leading strand synthesis (Figure

16 2C+D). Consistent with the native gel data, nascent strand analysis on denaturing gels demonstrates that CD

17 and HO R-loops cause a decrease in full-length rightward leading strands by ~ 75 % and ~ 55 %, respectively

18 (Figure 2C). The levels of leftward leading strands are not affected, confirming that the loss of full-length

19 rightward leading strands is not due to reduced origin activity. Instead, the loss of full-length rightward leading

20 strands correlates with the generation of prominent ~2.3 kb or ~2.8 kb stall products at CD and HO R-loops,

21 respectively. The heterogeneity of the stalled leading strand products is reduced after ClaI digest, confirming

22 that they originate near ARS305 (Figure S2A). Accordingly, leading strand stalling coincides with the Airn

23 sequence, which spans the region 1.8 kb – 3.2 kb downstream of the origin (Figure 2A). The difference in the

24 position of the leading strand stall sites at CD and HO R-loops is, therefore, attributable to the asymmetric

25 distribution of R-loops in the Airn sequence (Figure 1G). The heterogeneity in stall products remaining after

26 ClaI digest is likely a consequence of variations in R-loop sizes and positions (Figure 1).

27 Two-dimensional gel analysis demonstrates that leading strand stalling at CD and HO R-loops results

28 from both fork stalling and helicase uncoupling (Figure 2E). Time course analyses demonstrate that leading 6 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 strands stall within minutes of origin firing and remain stable for at least two hours (Figure S2B+C). Similarly,

2 native gel analysis reveals that a fraction of forks remains stably stalled specifically in the presence of R-loops

3 (Figure S2B+C). Fork uncoupling occurs after leading strand stalling, as expected due to the reduced rate of

4 DNA unwinding by CMG upon uncoupling from leading strand synthesis 42,50.

5 In addition to stalled leading strands, denaturing gel analysis reveals the formation of novel ~5.5 kb and

6 ~5.0 kb products indicative of leading strand restart downstream of CD and HO R-loops, respectively (Figure

7 2C+D). The identity of these products is confirmed by multiple lines of evidence: First, these products are

8 sensitive to cleavage by the leading strand-specific nicking enzyme Nb.BbvCI, but not lagging strand-specific

9 Nt.BbvCI, demonstrating that they are nascent leading strand products (Figure S2D). Second, these leading

10 strand products are not sensitive to ClaI cleavage, indicating that they do not originate near ARS305 (Figure

11 S2A). Third, in regular time course experiments, these products accumulate with slower kinetics than stalled

12 leading strands, as expected for restart following leading strand stalling (Figure S2B). Fourth, in experiments

13 in which nascent strands are pulse-labeled with [32P]-dATP for 2.5 minutes following origin activation before

14 being chased with excess cold dATP restart products are not detectable, as expected if they form after origin

15 activation (Figure S2C). Moreover, below we will show that this leading strand restart occurs at G4s in the

16 leading strand template.

17 We conclude that both CD and HO R-loops can perturb replication fork progression, which is consistent

18 with observations in human cells 14. Replication abnormalities in either orientation include fork stalling,

19 uncoupling of leading strand synthesis from fork progression, and discontinuous leading strand synthesis

20 involving leading strand restart. A fraction of R-loops in either orientation is also bypassed by replisomes

21 without disruption. This diversity in outcomes is explained by the heterogeneity of R-loops in our templates

22 (Figure 1). Because the replicative DNA helicase, CMG, can efficiently bypass steric blocks on the displaced

23 lagging strand 51,52, fork stalling is likely induced by obstacles on the leading strand. This raises questions

24 about the fork stalling mechanism as forks will encounter either RNA:DNA hybrids or the displaced non-

25 template strand on the leading strand during CD and HO R-loop-replisome collisions, respectively. We,

26 therefore, investigated the molecular basis for replication aberrations at R-loops.

28 Fork stalling at R-loops is not dependent on Tof1 7 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 Csm3, Tof1, and Mrc1 (CTM) form a fork protection complex (FPC) that associates with replisomes to

2 promote normal fork progression, prevent the uncoupling of replisomes from DNA synthesis after nucleotide

3 depletion by hydroxyurea (HU), and maintain fork integrity during the replication of structure-forming

4 trinucleotide repeats 53-58. In addition, Tof1 mediates fork pausing at protein-DNA complexes, such as the rDNA

5 replication fork barrier (RFB), tRNA genes, and centromeres 57,59,60. We find that fork stalling, uncoupling, and

6 restart at CD and HO R-loops are not affected by CTM, demonstrating that the mechanism of fork stalling at R-

7 loops is distinct from that at protein-DNA barriers (Figure S2E).

9 PCNA suppresses helicase-polymerase uncoupling at Airn sequence

10 A previous study found that RNA at 5’ primer-template junctions promotes strand-displacement by Pol 

11 61. We, therefore, tested if Pol  may promote fork progression through R-loops (Figure 2F). On R-loop-

12 containing templates, replication intermediates obtained in the absence of Pol  or its processivity factor PCNA

13 (together with its loader RFC) were indistinguishable from those obtained with complete replisomes. Thus,

14 strand-displacement by Pol  does not promote fork progression at R-loops. The data also demonstrates that

15 Pol  is not required for leading strand restart under these conditions. Intriguingly, however, we note that the

16 absence of PCNA increases leading strand stalling and fork uncoupling at the Airn sequence even in the

17 absence of R-loops (Figure 2F, lanes 3+9), indicating a role for PCNA in maintaining the coupling of

18 replisomes to leading strand synthesis at G4 sequences. We speculate that PCNA mediates this function

19 through stabilization of the leading strand polymerase, Pol , on the template.

21 RNase H1 promotes fork passage specifically at CD R-loops

22 RNase H1 overexpression is commonly used to assess the contribution of R-loops to TRC. We,

23 therefore, tested how purified yeast RNase H1 (Figure 3A) affects fork progression at R-loops in vitro. At CD

24 R-loops, RNase H1 decreased the levels of stalled and uncoupled replication intermediates while increasing

25 the formation of full-length replication products, suggesting that RNA:DNA hybrids on the leading strand can

26 impede fork progression (Figure 3B+C, lanes 1-4; Figure S3A). In contrast, leading strand restart at CD R-

27 loops was not affected by RNase H1, indicating that restart is not a direct consequence of RNA:DNA hybrids.

8 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 Instead, this suggests a role for G4s in leading strand restart at CD R-loops (Figure S3B), which is also

2 supported by data below. While not being dependent on RNA:DNA hybrid persistence, we note that formation

3 of the restart-inducing structures is dependent on transcription (Figure 3B, compare lanes 1+2 to 3+4).

4 In contrast, fork stalling and uncoupling were largely unaffected by RNase H1 at HO R-loops, indicating

5 that RNA:DNA hybrids on the lagging strand are not the direct cause for these events (Figure 3B, lanes 5-8).

6 Instead, this data again suggests that G4s on the leading strand template, formed by the G-rich displaced

7 strand in this orientation, cause fork stalling or uncoupling. Moreover, the insensitivity of the fork block at HO

8 R-loops to RNase H1 demonstrates that G4s on the leading strand template, while dependent on transcription

9 for formation, are not dependent on the persistence of RNA:DNA hybrids on the lagging strand template

10 (Figure S3C). In contrast, leading strand restart at HO R-loops is sensitive to RNase H1, indicating that the

11 restart-inducing G4s in this case are stable only in single-stranded DNA, i.e. in the presence of the RNA:DNA

12 hybrid on the opposite strand, and therefore differ from those G4s inducing fork stalling at some HO R-loops

13 (Figure S3D).

15 Pif1 promotes fork passage specifically at HO R-loops

16 The implication of G4s in the fork block at R-loops led us to examine the effect of the Pif1 helicase on

17 fork progression at R-loops. In vitro studies have demonstrated that Pif1 exhibits an evolutionarily conserved

18 specificity for binding and unwinding G4 DNA 29,62-67. Moreover, Pif1 has been demonstrated to promote fork

19 progression through G4 sequences in vivo and to suppress genomic instability at such sites 29,30,32,68. In

20 addition, Pif1 exhibits enhanced activity on RNA:DNA hybrids in vitro 64,69,70.

21 In contrast to RNase H1, we find that Pif1 does not promote fork progression through CD R-loops,

22 supporting the notion that its limited processivity is insufficient to resolve long RNA:DNA hybrids (Figure 3C,

23 lanes 1-3) 71. However, RNase H1 and Pif1 together effectively eliminate fork stalling, uncoupling, and restart

24 events CD R-loops (Figure 3C, lane 4). This suggests that RNase H1 and Pif1 coordinately promote fork

25 progression at CD R-loops by eliminating both RNA:DNA hybrids and G4s from the leading strand template.

26 Conversely, at HO R-loops Pif1 was markedly more efficient than RNase H1 in promoting fork progression

27 (Figure 3C, lanes 5-7), which is consistent with the notion that G4s on the leading strand template cause fork

28 stalling at HO R-loops. Accordingly, addition of RNase H1 did not further promote fork progression at HO R- 9 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 loops in the presence of Pif1 (Figure 3C, lane 8). Together, the data suggests that fork stalling at R-loops can

2 be induced by RNA:DNA hybrids and G4s on the leading strand template, while neither structure presents a

3 fork block when formed on the lagging strand.

5 G4s and RNA:DNA hybrids on the leading strand template can pose fork blocks

6 To directly test the impact of G4s on fork progression we performed reactions in the presence of the

7 G4-stabilizer, pyridostatin (PDS). On DNA templates that lack the Airn sequence PDS does not impede DNA

8 replication (Figure 3D, lanes 1+2). In contrast, PDS induces a strong block to fork progression specifically at

9 the Airn sequence in the HO orientation, i.e. when the G-rich strand forms the leading strand template, even in

10 the absence of R-loops (Figure 3D, lanes 3-6). Thus, G4s forming on the G-rich strand of the Airn sequence in

11 the absence of transcription impede replication forks only when stabilized by PDS (Figure 3D, lanes 3+5),

12 demonstrating that G4 stability determines the fork block potential of G4s. In contrast, fork stalling is induced at

13 HO R-loops even in the absence of PDS, and this effect is enhanced by PDS (Figure 3D, lanes 7-10). Thus,

14 R-loop formation modulates the G4 composition on the displaced strand, which is consistent with studies

15 demonstrating a cooperative relationship between R-loops and G4s 26,27,72. Since RNase H1 is unable to

16 prevent fork stalling at HO R-loops, R-loop formation induces the formation of fork-stalling G4s on the

17 displaced strand but is not required for their maintenance.

18 In the CD orientation, PDS also induces fork stalling at the Airn sequence in the absence of R-loops,

19 but the extent of stalling is less pronounced than in the HO orientation, consistent with the lower G4 potential

20 on the C-rich strand (Figure 3E, lanes 1-4; Figure S1A). In addition, in the absence of R-loops, PDS induces

21 leading strand restart at the Airn sequence, demonstrating that replisomes can pass some PDS-stabilized G4s

22 and reprime leading strand synthesis downstream. Deletion of G4810-828 (Figure 1A; Figure S1A) eliminates

23 the major PDS-induced fork block in the CD orientation, identifying this G4 as a fork block (Figure 3E, lanes 9-

24 12). Deletion of G4810-828 exposes a novel downstream leading strand stall site that correlates with greatly

25 increased leading strand restart in the presence of PDS (Figure 3E, lanes 11+12). We conclude that G4s can

26 induce fork stalling or uncoupling depending on G4 stability, which is consistent with the correlation of genetic

27 instability with G4 stability in vivo 73.

10 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 Replisome uncoupling from leading strand synthesis at PDS-stabilized G4s indicates that some G4s

2 pose a block to the leading strand polymerase, Pol , but not the CMG helicase (Figure 3E, native gel, lanes

3 2+4). Importantly, while a fraction of uncoupling events at G4s leads to persistent unwinding in the absence of

4 DNA synthesis, we also observe significant leading strand restart downstream of G4s. We note that the leading

5 strand restart efficiency observed at G4s appears to be greater than that observed previously at leading strand

6 DNA damage 74, which we will discuss further below. Strikingly, unlike PDS-induced fork stall events in the CD

7 orientation, CD R-loop-induced fork stall events are sensitive to RNase H1, occur at a position closer to the

8 origin-proximal side of the Airn sequence, and are not exacerbated by PDS (Figure 3E, compare lanes 5+6 to

9 1+2). This indicates that fork stalling at R-loops in the CD orientation is caused by RNA:DNA hybrids on the

10 leading strand template. Accordingly, stall sites are shifted downstream at CD R-loops in the presence of both

11 RNase H1 and PDS (Figure 3E, lanes 6+8).

12 In summary, the data demonstrates that both RNA:DNA hybrids and G4s on the leading strand can

13 induce fork stalling during R-loop-replisome collisions. Moreover, both RNA:DNA hybrids and G4s have the

14 potential to induce uncoupling of leading strand synthesis from fork progression. At G4s transient uncoupling of

15 leading strand synthesis from fork progression is frequently followed by leading strand restart. Notably, a

16 fraction of R-loops in either orientation is also bypassed by forks without disruption. This diversity in outcomes

17 is likely a consequence of the structural heterogeneity of R-loops.

19 CMG can unwind or translocate on RNA:DNA hybrids

20 To determine the molecular basis for the diversity in outcomes of R-loop-replisome collisions, we

21 examined the helicase activity of purified CMG (Figure 4A) on RNA:DNA hybrid- and G4-containing

22 substrates. During DNA unwinding at replication forks, CMG translocates on the single-stranded leading strand

23 template in 3’-5’ direction while sterically displacing the lagging strand. Additionally, purified yeast and human

24 CMG have been demonstrated to be able to translocate on duplex DNA, which is supported by studies in

25 Xenopus suggesting that CMG translocates on dsDNA after replication termination or upon encounter of nicks

26 in the lagging strand template 75-78. To determine if CMG can translocate on RNA:DNA hybrids we constructed

27 oligonucleotide substrates composed of a 40 bp forked duplex DNA preceded by a 40 bp DNA or RNA:DNA

28 duplex that forms a flush 5’ ss/dsDNA junction downstream of a 40 nt poly(dT) 3’ tail, which serves as the 11 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 loading site for CMG. A 5’ hairpin-like secondary structure promotes the steric unwinding of the downstream

2 oligonucleotide by an approaching CMG 52. Using this approach, we find that the downstream oligonucleotide

3 is unwound with similar efficiencies by CMG if preceded by a DNA or RNA:DNA duplex (Figure 4B i+ii).

4 Importantly, unwinding of the downstream oligonucleotide is not accompanied by the unwinding of the

5 upstream DNA or RNA oligonucleotide (Figure 4B iii+iv). Thus, CMG can translocate on both DNA and

6 RNA:DNA duplexes. Next, to test the ability of CMG to unwind RNA:DNA hybrids, we constructed

7 oligonucleotide substrates comprising a forked 60 bp duplex in which the template strand is composed of DNA

8 and the non-template strand is composed of either DNA or RNA. Time course analyses demonstrate that CMG

9 unwinds RNA:DNA and DNA:DNA duplexes with similar efficiencies (Figure 4C).

10 Thus, RNA:DNA hybrids on the leading strand at CD R-loops are not automatic blocks to fork

11 progression. In the presence of a 5’ flap, the RNA:DNA hybrid can be unwound by CMG, which may account

12 for the generation of full-length replication products observed here in the presence of CD R-loops, and which

13 could also explain the reduction in R-loop levels during CD transcription-replication collisions in human cells 14.

14 In contrast, in the absence of a 5’ flap, CMG can translocate on RNA:DNA hybrids without displacing the RNA.

15 Such events may occur at R-loops harboring a flush 5’ RNA end or at nicks in the RNA, analogous to

16 replisome encounters with nicks in the lagging strand DNA template 78. Since the RNA:DNA hybrid will form a

17 block to Pol , which lacks strand-displacement activity, CMG translocation across RNA:DNA hybrids would

18 induce uncoupling of leading strand synthesis from fork progression, thus providing a molecular basis for the

19 RNase H1-sensitive uncoupling observed at CD R-loops (e.g. native gel, Figure 3C, lanes 1+2).

21 G4s on the template strand can block DNA unwinding by CMG

22 Next, we tested the impact of G4s on the DNA unwinding activity of CMG. For this, we incorporated a

23 G4 sequence of predicted high stability (G4-wt: 5’-[GGGT]3GGG-3’) or a mutant derivative (G4-mut: 5’-

24 GGGTCCCTGGGTGGG-3’) in the template or non-template strand of 60 bp forked duplex oligonucleotide

25 substrates. Strikingly, the wildtype G4 sequence, but not its mutant derivative, strongly attenuates DNA

26 unwinding by CMG when placed in the template strand (Figure 4D). In contrast, substrate unwinding

27 efficiencies were equivalent in the presence of either wildtype or mutant G4 sequences on the non-template

28 strand (Figure 4E). Thus, G4s on the leading strand template have the potential to block DNA unwinding by 12 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 CMG, explaining the fork stalling at the Airn sequence in the presence of HO R-loops or PDS. Consistent with

2 the steric exclusion model for DNA unwinding by CMG 52, G4s on the lagging strand do not pose an obstacle to

3 fork progression, enabling RNase H1 to promote efficient fork progression at CD R-loops.

5 G4s at CD R-loops can induce lagging strand gaps that can be resolved by Pif1

6 Previous primer extension studies have demonstrated that G4s can impede DNA synthesis by the

7 lagging strand polymerase, Pol  79,80. If G4s also inhibit progression of the lagging strand polymerase in the

8 context of replisomes is not known. Since the experiments so far were performed in the absence of Fen1 and

9 Cdc9 to allow examination of leading strands, lagging strand blocks were not detectable in the experiments

10 above. Therefore, to investigate the impact of R-loops on lagging strand synthesis, we performed DNA

11 replication reactions in the presence of Fen1 and Cdc9 40,42.

12 In the CD orientation and in the absence of R-loops, replication products obtained in the presence of

13 Fen1/Cdc9 correspond primarily to full-length replication products, indicating that the G-rich strand of the Airn

14 sequence does not pose an intrinsic obstacle to lagging strand synthesis (Figure 5A, lanes 1-4). In the

15 presence of CD R-loops, leading strand stall and restart products are detected as before in the absence of

16 Fen1/Cdc9 (Figure 5A, lanes 5+7). However, while RNase H1 is sufficient to prevent fork stalling at CD R-

17 loops (Figure 5A, lanes 5+6; Figure 3), in the presence of Fen1/Cdc9 and RNase H1 new replication products

18 are observed at CD R-loops that are identical in size to leading strand stall and restart products (2.3 kb and 5.5

19 kb, respectively), but exhibit reversed band intensities (Figure 5A, lane 8). These products correspond to

20 lagging strand products generated upstream (LagUS) or downstream (LagDS) of the R-loop, respectively (Figure

21 5B). Two lines of evidence are consistent with this interpretation. First, formation of these products is

22 dependent on Okazaki fragment ligation (Figure 5A, lanes 6+8). Second, LagUS, but not the stalled leading

23 strand product of same length, is sensitive to cleavage by the lagging strand specific nicking enzyme Nt.BbvCI

24 (Figure 5B+C): In the absence of RNase H1, Nt.BbvCI digestion separates the 2.3 kb replication products into

25 cleavage-resistant stalled leading strands (2.3 kb) and truncated LagUS (1.9 kb; Figure 5C, lanes 1+3). In

26 contrast, in the presence of RNase H1, i.e. in the absence of fork stalling, the entire population of 2.3 kb

27 products is sensitive to Nt.BbvCI cleavage and thus corresponds to LagUS (Figure 5C, lanes 2+4).

13 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 This data is consistent with the notion that R-loop formation induces G4s on the displaced non-template

2 strand, blocking the progression of the lagging strand polymerase during CD replisome collisions, thereby

3 causing a gap in the nascent lagging strand. We, therefore, tested the ability of Pif1 to overcome the lagging

4 strand block at CD R-loops. Indeed, addition of Pif1 greatly reduced the formation of lagging strand gaps at CD

5 R-loops in the presence of RNase H1 (Figure 5D, lanes 2+4). Thus, Pif1 promotes lagging strand synthesis at

6 G4s in vitro, which is supported by observations in vivo 32.

8 Both G4s and RNA:DNA hybrids cause lagging strand gaps at HO R-loops

9 While the G-rich displaced strand forms the lagging strand template at CD R-loops, RNA:DNA hybrids

10 form on the lagging strand template at HO R-loops, presenting a potentially distinct challenge to lagging strand

11 synthesis. Intriguingly, even in the absence of R-loops a prominent lagging strand gap is observed at the Airn

12 sequence in the HO orientation (Figure 6A, lanes 1-4; Figure 6E i). Restriction analysis confirms the position

13 of LagUS and LagDS (Figure S4). The lagging strand gap is caused by G4810-828, as deletion of this G4 sequence

14 is sufficient to eliminate the gap (Figure 6B, lanes 1-4).

15 In the presence of R-loops, the length of LagDS is slightly reduced by treatment with RNase H1 (Figure

16 6A, lanes 7+8), indicating that LagDS stalls at RNA:DNA hybrids during HO R-loop-replisome collisions, but at

17 G4810-828 after resolution of the RNA:DNA hybrid by RNase H1 (Figure 6E ii). Consistent with RNA:DNA

18 hybrids impeding lagging strand synthesis, HO R-loops cause an RNase H1-sensitive lagging strand gap also

19 on templates in which G4810-828 has been deleted (Figure 6B, lanes 5-8). As shown above (Figure 3), leading

20 strand stalling persists at HO R-loops even in the presence of RNase H1 due to G4s on the G-rich leading

21 strand template, giving rise to stalled leading strand products that are similar in size to LagUS (Figure 6B, lane

22 8; Figure 6E iii).

23 Consistent with the observations above, we find that Pif1 promotes lagging strand synthesis at G4810-828

24 in the absence of R-loops (Figure 6C). This Pif1 function is specifically dependent on the Pif1 helicase activity,

25 as mutation of the Pif1 Walker B motif, which has been shown to disrupt the helicase but not DNA binding

26 activity of the human Pif1 orthologue 81, abrogates the ability of Pif1 to promote lagging strand synthesis at

27 G4810-828 (Figure 6C). In contrast, Pif1 alone was unable to promote lagging strand synthesis at RNA:DNA

28 hybrids (Figure 6D, lanes 1+3) but promoted completion of lagging strand synthesis in conjunction with RNase 14 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 H1 (Figure 6D, lanes 2+4). We conclude that both RNA:DNA hybrids and G4s can inhibit lagging strand

2 synthesis at R-loops, and this inhibition is reversed by RNase H1 and Pif1, respectively.

4 R-loop transcripts can prime leading strand restart after CD R-loop-replisome collisions

5 Studies in E. coli have demonstrated that transcripts can prime leading strand synthesis after CD

6 collisions of replisomes with RNAP 82. Whether a similar mechanism exists in eukaryotes, is unknown. This

7 question is particularly intriguing as CMG tracks along the leading strand template, while the bacterial

8 replicative DNA helicase, DnaB, tracks along the lagging strand template. Our purification protocol for R-loop

9 templates involves digestion of free RNA with RNase A. However, RNase A cleavage leaves a 3’ phosphate

10 (3’-P), which prevents extension by DNA polymerase. We, therefore, converted RNA 3’-P ends into 3’-hydroxyl

11 (3’-OH) ends using T4 polynucleotide kinase (T4 PNK). Strikingly, T4 PNK treatment significantly increased the

12 levels of leading strand restart products at CD R-loops (Figure 7A, lanes 1+3). Unlike leading strand restart

13 products obtained at CD R-loops in the absence of T4 PNK treatment, these novel restart products are

14 sensitive to RNase H1 treatment (Figure 7A). Moreover, formation of these products is dependent on DNA

15 synthesis at replisomes (Figure 7B) and is not observed at HO R-loops, in which the RNA:DNA hybrid is on

16 the lagging strand (Figure 7C). Together, this data indicates the repriming of leading strand synthesis at the

17 RNA 3’-OH of RNA:DNA hybrids (Figure 7D).

18 We find that under the conditions used here Pol , but not Pol  or Pol  is capable of efficiently

19 extending R-loop-associated RNA with DNA (Figure S5A). Extension by Pol  is limited to ~ 200 – 700 nt of

20 DNA and is further increased to up to ~ 2.5 kb in the presence of all three polymerases, well below the length

21 of complete restart products observed in the context of replisomes. This suggests that leading strand restart at

22 R-loop RNA during CD R-loop-replisome collisions is initiated by Pol  and extended by Pol  and/or Pol .

23 Although Pol  is not required for leading strand restart at R-loop RNA (Figure S5B), we do not rule out its

24 involvement, as discussed below.

25 How may replisomes utilize R-loop transcripts to prime leading strand restart? It is possible that CMG

26 slides over RNA:DNA duplexes at CD R-loops harboring nicks in the RNA, analogous to replisome encounters

27 with nicks in the lagging strand template 78, which is supported by our demonstration that CMG can translocate

28 on RNA:DNA duplexes (Figure 4). To test this model, we treated CD R-loops with sub-saturating 15 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 concentrations of RNase H1 to introduce nicks in the RNA of CD R-loops. While the formation of full-length

2 leading strand products increases with the concentration of RNase H1 due to the resolution of the RNA:DNA

3 hybrids, leading strand restart correlates inversely with the concentration of RNase H1 (Figure 7E).

4 Concomitantly, RNase H1 treatment reduces the formation of uncoupled products in the native gel analysis.

5 Similarly, we find that a sub-saturating concentration of purified RNase H2 promotes leading strand restart

6 specifically at CD R-loops (Figure S5C). We note that RNase H generates a 3’-OH at the cleavage site,

7 eliminating the requirement for T4 PNK to convert RNA 3’-P ends into 3’-OH ends for restart here 83. We

8 conclude that R-loop-associated RNAs can prime leading strand restart when CMG translocates over

9 RNA:DNA hybrids during CD R-loop-replisome collisions.

11 DISCUSSION

12 R-loops perform important physiological roles in the cell but can also be harmful if generated in an

13 unscheduled manner. Accordingly, R-loops are considered ‘good’ or ‘bad’. What distinguishes ‘good’ R-loops

14 from ‘bad’ ones is unclear. Moreover, although R-loops have been widely recognized as a major determinant of

15 genome instability caused by TRC, the mechanism by which R-loops impede DNA replication has remained

16 obscure. By reconstituting eukaryotic R-loop-replisome collisions in vitro we demonstrate here that the specific

17 structural configuration of R-loops determines the outcome of R-loop-replisome collisions, providing a

18 mechanistic explanation for ‘good’ and ‘bad’ R-loops.

19 We uncover several ways in which replisomes can continue progression at R-loops. 5’ RNA flaps

20 promote the unwinding of RNA:DNA hybrids by CMG on the leading strand, allowing unhindered fork

21 progression. This mechanism is consistent with the observation that CD transcription-replication collisions

22 reduce R-loop levels in cells 14. In the HO orientation forks may also progress continuously, provided the

23 displaced R-loop strand, which forms the leading strand template for the replisome, is devoid of inhibitory

24 secondary DNA structure. We demonstrate here that G4s can inhibit DNA unwinding by CMG, but it is

25 conceivable that other DNA secondary structures forming on the displaced non-template strand similarly

26 impede fork progression 84. The ability of replisomes to pass through G4s is likely modulated by G4 stability,

27 which has been shown to influence the impact of G4s on genome stability 73.

16 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 In addition, we demonstrate that forks can progress through R-loops harboring RNA:DNA hybrids or

2 G4s with polymerase-stalling potential on the leading strand. This results in the uncoupling of leading strand

3 synthesis from replisome progression. Persistent uncoupling would be expected to induce the S phase

4 checkpoint in cells, which may facilitate the clearance of polymerase blocks before S phase completion 85,86.

5 Importantly, we also identify distinct mechanisms by which leading strand synthesis can be restarted at G4s

6 and RNA:DNA hybrids.

7 In the case of G4s, leading strand synthesis is reprimed downstream of the block, causing a gap in the

8 nascent leading strand. Transient uncoupling of leading strand synthesis from fork progression at G4s has

9 been suggested to cause epigenetic instability in vertebrate cells, where repriming is promoted by PrimPol 87.

10 Notably, the leading strand restart observed at G4s here appears to be significantly more efficient than that

11 reported previously at DNA damage sites 74. We hypothesize that transient stalling of CMG, which may be less

12 pronounced at DNA damage that does not pose a physical obstacle to the CMG, may promote leading strand

13 repriming at G4s.

14 Our data demonstrates that leading strand restart at RNA:DNA hybrids occurs by an ‘on-the-fly’

15 mechanism in which Pol  extends the RNA 3’ end with DNA before handover to the leading strand

16 polymerase at the replisome (Figure 7F). To a degree this restart mechanism is analogous to that at DNA

17 damage sites, where restart is initiated by TLS polymerases or Pol  88,89. The involvement of Pol  can be

18 rationalized by its inherent preference for RNA/DNA junctions 90. Whether the handover of DNA synthesis from

19 Pol  to Pol  involves Pol  at RNA:DNA hybrids remains to be determined. The ability of transcripts to

20 reprime leading strand synthesis was previously observed E. coli 82. However, while the replicative DNA

21 helicase in E. coli translocates on the lagging strand, eukaryotic replisomes are assembled around CMG on

22 the leading strand, indicating that the respective mechanisms are fundamentally distinct. We demonstrate that

23 CMG can translocate across RNA:DNA duplexes, allowing the replisome to access RNA 3’ ends of RNA:DNA

24 duplexes. This mechanism requires the RNA 5’ end of RNA:DNA hybrids to be annealed to the template strand

25 in order to prevent unwinding by the replisome. The structure of R-loops in vivo is not known and is likely

26 variable. However, we note that R-loop formation may be promoted by invasion of the DNA duplex by the RNA

27 5’ end in order to limit topological clashes during the winding of the RNA around the template strand 91.

28 Alternatively, flush RNA 5’ ends may be encountered by the replisome at nicks in the RNA, analogous to 17 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 lagging strand nicks 78. Such nicks may be generated by RNase H1 or RNase H2, whose activities are spatially

2 and temporally limited, respectively 92,93.

3 We find that both RNA:DNA hybrids and G4s can also induce gaps in nascent lagging strands. This is

4 consistent with lagging strand G4s impeding DNA replication in vivo and with G4s posing a block to Pol  in

5 primer extension assays 32,80. However, the block to lagging strand synthesis by RNA:DNA hybrids is surprising

6 given the proficiency of the lagging strand machinery in removing RNA:DNA hybrids at Okazaki fragment 5’

7 ends 94. The reasons for this are currently unknown, but it is possible that long RNA 5’ flaps or G4s in the RNA

8 strand impinge on the ability of Pol  and Fen1 to process RNA:DNA hybrids.

9 We show that RNA:DNA hybrid- and G4-induced blocks to fork progression, as well as leading and

10 lagging strand synthesis, are mitigated by RNase H1 and Pif1, respectively. This data is consistent with in vivo

11 studies demonstrating that both RNase H1 and Pif1 suppress genome instability at R-loops 21,22,95 and promote

12 replication fork progression at R-loops and G4 sequences, respectively 8,68,96. We note that the coincidence of

13 RNA:DNA hybrids and G4s at R-loops may limit the efficiency of RNase H1 overexpression to resolve R-loops,

14 which is commonly used to assess the contribution of R-loops to TRC.

15 R-loops in both the CD and HO orientation can induce persistent fork stalling, which requires the

16 RNA:DNA hybrid- and/or G4-resolving activities of RNase H1 and Pif1 for continued fork progression.

17 However, we also find that replisomes have an intrinsic capacity to progress through R-loops by bypassing or

18 unwinding G4s or RNA:DNA hybrids. Despite the potential to induce nascent strand gaps, continued fork

19 progression may be beneficial in order to establish homology-directed repair-competent chromatin on sister

20 chromatids 97,98. Nascent strand discontinuities at R-loops may also be processed post-replicatively 99, which

21 could allow the timely progression of S phase. Alternatively, replisome bypass of R-loops may promote the

22 resolution of R-loops analogous to the repair of DNA-protein crosslinks 100.

23 The molecular characterization of R-loop-replisome collision presented here enhances our

24 understanding of the mechanism(s) by which G4 ligands can affect the growth of cancer cells 27. Moreover, we

25 expect that the system developed here will aid future studies directed at characterizing the molecular functions

26 of the multitude of additional factors implicated in the processing of RNA:DNA hybrids and G4s 2,34.

28 18 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 ACKNOWLEDGMENTS

2 This work was supported by a MSKCC Functional Genomics Initiative grant (DR), NIGMS grants R01-

3 GM127428 and R01-GM107239 (DR), NIH grant ES031635 (JDG), and NIH/NCI Cancer Center Support Grant

4 P30 CA008748. We thank Sujan Devbhandari for help with CMG helicase assays, Ademola Adegbemigun for

5 help with protein purification, and Iestyn Whitehouse for helpful discussions.

7 AUTHOR CONTRIBUTIONS

8 C.K., S.B., and J.D.G. conducted the experiments. D.R. and C.K. designed the experiments. D.R. wrote

9 the paper with input from C.K.

11 DECLARATION OF INTERESTS

12 The authors declare no competing interests.

19 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 FIGURE LEGENDS

3 Figure 1: Preparation and characterization of R-loop-containing templates. (A) Schematic of R-

4 loop plasmid template. Plot shows G/C skew at Airn sequence. Graphic shows positions of potential G-

5 quadruplexes composed of stacks of three G-quartets in non-template (top) and template (bottom) strand. 3x

6 T: T7 terminator tandem repeat. (B) Schematic of CD and HO R-loop-replisome collisions in experimental

7 setup. Template strands: black; leading strand: red; lagging strand: blue; RNA: green. (C) Reaction scheme for

8 preparation of R-loop-containing template. (D) Native agarose gel analysis of purified plasmid template. The

9 gel was stained with ethidium bromide. (E) R-loop-containing template harboring 32P-labeled RNA was mock-

10 treated or digested with RNase H and analyzed by denaturing formaldehyde agarose gel-electrophoresis and

11 autoradiography. (F) EM analysis of R-loop templates. White arrows in center panels indicate S9.6-specific

12 density. Thin and thick arrows in right panels indicate displaced non-template or RNA:DNA duplex,

13 respectively. (G) Frequency distribution of R-loop distances from ClaI site in CD and HO orientation.

15 Figure 2: Both CD and HO R-loops perturb normal fork progression. (A) Schematic illustrating

16 expected sizes of replication products. (B) Denaturing agarose gel analysis of replication products obtained on

17 R-loop-free template. Left lead: Leftward leading strands; Right lead: Rightward leading strands. (C) Native

18 (top) and denaturing (bottom) agarose gel analysis of replication products obtained on templates harboring

19 Airn sequence in CD or HO orientation. Stall: Stalled rightward leading strands; Restart: Rightward leading

20 strand restart product; Full-length: Full-length rightward leading strand; RI: Replication Intermediates. (D)

21 Schematic illustrating replication products observed in (C). (E) Two-dimensional gel analysis of replication

22 products obtained in presence of R-loops (corresponding to lanes 4 and 8 in (C)). Products were digested with

23 ClaI. (F) Replication products obtained in the absence or presence of RFC/PCNA or Pol .

25 Figure 3: G4s and RNA:DNA hybrids pose impediment to leading strand synthesis that can be

26 resolved by Pif1 or RNase H1, respectively. (A) Purified proteins analyzed by SDS-PAGE and Coomassie

27 stain. (B) Replication products obtained on CD or HO templates in the absence or presence of RNase H1. (C)

28 Replication products obtained on CD or HO templates in the absence or presence of RNase H1 and Pif1. (D) 20 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 Replication products obtained on HO templates in the absence or presence of RNase H1 and pyridostatin. (E)

2 Replication products obtained on CD templates in the absence or presence of RNase H1 and pyridostatin.

3 G4810-828: Airn sequence containing deletion of G4 sequence at position 810-828.

5 Figure 4: CMG can unwind or translocate on RNA:DNA hybrids, while G4s can block DNA

6 unwinding by CMG. (A) Purified CMG. (B) Helicase assays with 40 bp forked DNA duplex preceded by 40 bp

7 DNA (i + iii) or RNA:DNA (ii + iv) duplex. indicates position of 5’-32P label. Products were analyzed by native

8 PAGE and autoradiography. (C) CMG helicase activity on 60 bp forked DNA (left) or RNA:DNA duplex (right).

9 Plot shows average of two replicates each. (D) CMG helicase activity on 60 bp substrate harboring wildtype

10 (left) or mutant (right) G4 sequence on the template strand (‘lead’). (E) As (D), with wildtype (left) or mutant

11 (right) G4 sequence on the non-template strand (‘lag’).

13 Figure 5: G4s at CD R-loops can induce lagging strand gaps that can be resolved by Pif1. (A)

14 Replication products obtained on CD template in the absence or presence of Fen1/Cdc9 and RNase H1 (B)

15 Schematic illustrating replication products observed in (A). (C) Replication products obtained on CD template

16 in the presence of Fen1/Cdc9. RNase H1 was included and products were digested with Nt.BbvCI as

17 indicated. (D) Replication products obtained on CD template in the presence of Fen1/Cdc9. Pif1 and RNase

18 H1 were included as indicated.

20 Figure 6: Both G4s and RNA:DNA hybrids cause lagging strand gaps at HO R-loops. (A)

21 Replication products obtained on HO template in the absence or presence of Fen1/Cdc9 and RNase H1. (B)

22 Replication products obtained on HO template harboring G4810-828 deletion in the absence or presence of

23 Fen1/Cdc9 and RNase H1. (C) HO template lacking R-loop was replicated with Fen1/Cdc9 in the absence or

24 presence of Pif1 or Pif1-E303Q. Signal intensities of LagUS and LagDS were quantified and normalized to

25 reactions without Pif1. (D) HO template replicated with Fen1/Cdc9 in the absence or presence of Pif1. (E)

26 Schematic illustrating replication products observed in A-D.

21 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 Figure 7: R-loop transcripts can prime leading strand restart after CD R-loop-replisome

2 collisions. (A) Replication products obtained on mock- or T4 PNK-treated CD R-loop-containing templates in

3 absence or presence RNase H1. (B) Products obtained on CD T4 PNK-treated R-loop templates in absence or

4 presence of DDK. Relative signal intensity for restart product is quantified on the right. (C) Replication products

5 obtained on mock- or T4 PNK-treated CD R-loop-containing templates in absence or presence RNase H1. (D)

6 Schematic illustrating replication products observed in A. (E) RNase H1 titration into reactions with CD R-loop

7 template. (F) Model for leading strand restart at R-loop transcript after replisome encounter with CD R-loop

8 harboring 5’ RNA flap and RNA nick.

11 Supplementary figures:

13 Figure S1: R-loop template preparation and characterization by AFM. (A) Sequence of Airn

14 sequence element. Top strand: non-template strand; bottom strand: template strand. QGRS Mapper

15 (https://bioinformatics.ramapo.edu/QGRS/index.php) was used to identify sequences with G-quadruplex

16 forming potential in non-template and template strand, highlighted in orange and green, respectively. Numbers

17 in brackets indicate G-quadruplex forming potential. (B) pARSR-loop was transcribed, the salt adjusted to 0.4 M

18 NaCl, and either mock-treated or treated with RNase A. Reactions were de-proteinated, phenol/chloroform

19 extracted, filtered through Illustra MicroSpin G25 Spin column, and 2 L of each sample analyzed by native

20 agarose gel-electrophoresis and ethidium bromide staining. (C) S1000 gel-filtration profiles of R-loop plasmid

21 templates. RNA is 32P-labeled. Fractions were analyzed by ethidium bromide staining (top) or autoradiography

22 (bottom) after native agarose gel-electrophoresis. (D) Sample AFM images of linearized R-loop templates.

23 Templates were incubated with yeast RPA prior to deposition on mica.

25 Figure S2: Both CD and HO R-loops perturb normal fork progression. (A) Replication products

26 obtained on CD and HO templates with and without ClaI digestion post replication. (B) Conventional time

27 course analysis of replication reactions on CD and HO templates. Time indicates minutes after addition of

28 Mcm10 (origin firing). (C) Pulse-chase analysis of replication reactions on CD (top) and HO (bottom) templates. 22 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 Signal intensities of replication products were quantified and plotted as percentage of total signal. (D) Top:

2 Schematic of products expected after digestion of full-length or restart replication products with Nt.BbvCI or

3 Nb.BbvCI. Bottom: Denaturing gel analysis of replication products digested as indicated. (E) Replication

4 products obtained on CD and HO templates in the absence or presence of Csm3-Tof1 and Mrc1 (CTM).

6 Figure S3: RNase H1 promotes fork passage specifically at CD R-loops. (A) Schematic of

7 replication products observed after replisome encounters with CD R-loops lacking G4s in the leading strand

8 template. Treatment with RNase H1 removes the RNA:DNA hybrid, allowing completion of DNA replication.

9 (B) Schematic of replication products observed after replisome bypass of CD R-loops harboring G4s in the

10 leading strand template. Bypass of CD R-loops or fork passage in the presence of RNase H1 results in the

11 uncoupling of leading strand synthesis at G4(s), which promotes restart of leading strand synthesis by

12 repriming. (C) Schematic of replication products observed after replisome encounters with HO R-loops

13 harboring G4s in the displaced strand that block CMG progression. Stalling persists upon RNase H1 treatment.

14 (D) Schematic of replication products observed after replisome encounters with HO R-loops harboring G4s in

15 the displaced strand that are stabilized in single-stranded DNA, i.e. when an RNA:DNA hybrid is present on the

16 template strand, and that block leading strand polymerase but not CMG progression.

18 Figure S4: Both G4s and RNA:DNA hybrids cause lagging strand gaps at HO R-loops. (A)

19 Denaturing gel analysis of replication products obtained in the presence of Fen1/Cdc9 on HO templates

20 lacking R-loops were digested with ClaI (lanes 1+2), ClaI and MscI (lanes 3+4), or ClaI and BseRI (lanes 5+6).

21 LagUS and LagDS are sensitive to Msc I and BseR I, respectively. (B) Schematic of expected products

22 generated by digestion of full-length replication products. (C) Schematic of expected products generated by

23 digestion of replication products containing a lagging strand gap.

25 Figure S5: R-loop transcripts can prime leading strand restart after CD R-loop-replisome

26 collisions. (A) Polymerase assay with T4 PNK-treated CD R-loop template in presence of Pol , Pol , or Pol

27 , as indicated. All reactions include RFC, PCNA, RPA, dNTPs, NTPs and -[32P]-dATP. The concentrations

28 of reaction components are equivalent to those used in the replication assay. (B) Replication products obtained 23 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 on RNase H1-treated CD R-loop templates with or without Pol . Reactions were performed either in the

2 presence of 1 nM (lanes 1+2) or 0.06 nM (lanes 3+4) RNase H1 to induce resolution or nicking of the

3 RNA:DNA hybrid, respectively. (C) Left: Purified RNase H2. Right: Replication products obtained on CD and

4 HO templates in the presence of sub-saturating levels of RNase H2.

24 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 METHODS

2 Table 1: List of plasmids used in this study Name of plasmid Features p1216 pCtrl (no AIRN) p1214 CD, AIRN p1215 HO, AIRN p1290 p1214 with BbvCI site 2075 bp downstream of AIRN p1285 p1215 with BbvCI site 2075 bp downstream of AIRN p1291 p1214 BbvCI site 11 bp upstream of AIRN p1292 p1215 BbvCI site 11 bp upstream of AIRN p1293 CD, AIRN 810-828 p1284 HO, AIRN 810-828 p1134 pET15b-6x-His-RNH1 p1254 pET15b-T7 Lysozyme-6x-His p1102 pET15b-PIF1-N p1130 pET15b-PIF1-N-E303Q 3 4 5 Table 2: Yeast strains Strain name Protein Genotype YSD35 CMG MAT a/ ade2-1 ura3-1 his3- 11,15 trp1-1 leu2-3,112 can1- 100 pep4::kanMX bar1::hphNAT1 Gal-GAL4 (HIS3) Gal-MCM5/MCM4 (TRP1) Gal-MCM2/FLAG- MCM3 (URA3) Gal- MCM7/MCM6 (LEU2) Gal1- 10 CDC45 (TRP1) Gal1-10 PSF2++/PSF3++ (URA3) Gal1-10 PSF1++/CBP- SLD5++ (LEU2)

YDR154 RNase H2 MATa ade2-1 ura3-1 his3- 11,15 trp1-1 leu2-3,112 can1-100 pep4::kanMX bar::hphNAT1 Gal-Gal4 (HIS3) GAL-CBP- RNH201++ (LEU2) GAL- RNH202++ / RNH203++ (URA3) 25 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 2 3 4 Table 3: List of oligonucleotides used to prepare templates for helicase assays Name Sequence 5’ to 3’ A GGCTCGTTTTACAACGTCGTGCTGAGGTGATATCTGCTGAGGCAATGGGAATTC GCCAACCTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTT B GGCAGGCAGGCAGGCAGGCAGGCAGGCAGGCAGGCAGGCAGGTTGGCGAATT CCCATTGCCTCAGCAGATATCACCTCAGCACGACGTTGTAAAACGAG C GGCAGGCAGGCAGGCAGGCAGGCAGGCAGGCAGGCAGGCAGGUUGGCGAAU UCCCAUUGCCUCAGCAGAUAUCACCUCAGCACGACGUUGUAAAACGAG D GGCAGGCAGGCAGGCAGGCAGGCAGGCAGGCAGGCAGGCAGATTAAGTTGGG TAACGCCAGGGTTTTCCCAGTCACGAC E GTCGTGACTGGGAAAACCCTGGCGTTACCCAACTTAATCGCCTTGCAGCACATC CCCCTTTCGCCAGCTGGCGTAATAGTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTT TTTTTTT F CTATTACGCCAGCTGGCGAAAGGGGGATGTGCTGCAAGGC G CUAUUACGCCAGCUGGCGAAAGGGGGAUGUGCUGCAAGGC H GGCAGGCAGGCAGGCAGGCAGGCAGGCAGGCAGGCAGGCAGGTTGGCGAATT CCCTTTTTTTTTTTTTTTTTTTCCTCAGCACGACGTTGTAAAACGAG I GGCTCGTTTTACAACGTCGTGCTGAGGTTGGGTGGGTGGGTGGGTTGGGAATT CGCCAACCTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTT J GGCTCGTTTTACAACGTCGTGCTGAGGTTGGGTGGGTCCCTGGGTTGGGAATT CGCCAACCTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTT K GGCTCGTTTTACAACGTCGTGCTGAGGTTTTTTTTTTTTTTTTTTTGGGAATTCGC CAACCTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTT L GGCAGGCAGGCAGGCAGGCAGGCAGGCAGGCAGGCAGGCAGGTTGGCGAATT CCCTTGGGTGGGTGGGTGGGTTCCTCAGCACGACGTTGTAAAACGAG M GGCAGGCAGGCAGGCAGGCAGGCAGGCAGGCAGGCAGGCAGGTTGGCGAATT CCCTTGGGTGGGTCCCTGGGTTCCTCAGCACGACGTTGTAAAACGAG 5 6 Table 4: List of templates used in helicase assays Template Oligonucleotides Figure 32P-labeled oligonucleotide Forked DNA duplex A+B 4C, left A Forked RNA-DNA A+C 4C, right A duplex DNA-DNA 1 D+E+F 4B(i) D DNA-DNA 2 D+E+F 4B(iii) F RNA-DNA 1 D+E+G 4B(ii) D RNA-DNA 2 D+E+G 4B(iv) G G-quad wt, lead H+I 4D, left H G-quad mut, lead H+J 4D, right H G-quad wt, lag K+L 4E, left K G-quad mut, lag K+M 4E, right K 7 8 9 10 DNA templates for replication assays

26 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 p1214 and p1215 were generated from pARS305 101. Cryptic origins were eliminated by excising the

2 region flanked by ClaI and the region flanked by BaeI and BsaAI. The Airn sequence was amplified from

3 pFC53 (kind gift from Fred Chedin) 20. T7 promoter, 3 x T7 terminators and the Airn sequence were inserted at

4 the BsaAI site by Gibson assembly. p1290 and p1285 were generated from p1214 and p1215, respectively, by

5 inserting a BbvCI recognition sequence between BglII and SwaI sites, 2075 bp downstream of the Airn

6 sequence. p1291 was generated from p1214 by inserting a BbvCI recognition sequence at the BstZ17I site.

7 p1292 was generated from p1215 by inserting a BbvCI recognition sequence at the ApaI site. To generate

8 p1293 and p1284, the Airn sequence at position 810-828 was deleted by PCR mutagenesis and the mutant

9 Airn sequence inserted at the BsaAI site of p1214 and p1215, respectively.

11 R-loop templates

12 Transcription reactions were carried out for 20 minutes at 37 oC in a reaction volume of 250 L

13 containing 20 g of plasmid template in 1x transcription buffer (Promega; 40 mM Tris-HCl pH 7.9 / 6 mM MgCl2

14 / 10 mM spermidine / 50 mM NaCl) / 25 U T7 RNA Polymerase (Promega) / 20 mM DTT / 0.05 % Tween 20 /

15 0.5 mM each ATP, GTP, CTP and UTP. Subsequently, 0.3 M T7 lysozyme were added to the reaction and

16 incubation continued for 10 minutes at 37 oC. Then, the salt concentration was adjusted to 400 mM NaCl, 10

17 g / ml RNaseA (ThermoScientific) were added to the reaction, and incubation continued for 30 minutes at 37

18 oC. Finally, 4 U of Proteinase K (NEB) were added to the reaction and incubation continued for 30 minutes at

19 37 oC. The reaction was concentrated to a volume of 50 L using Amicon Ultra 0.5 mL centrifugal filter and

20 fractionated through a custom-made 2 mL S1000 column equilibrated in 10 mM Tris-HCl pH 7.5. Template-

21 containing peak fractions were pooled, concentrated and stored at -20 oC. 32P-RNA-labeled R-loop templates

22 were generated as above, with the exception that transcription reactions were performed in the presence of

23 0.25 mM GTP and 60 Ci of -[32P]-GTP (Perkin Elmer).

25 Calculation of G/C skew:

27 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 GC skew at every nucleotide of the 1374 bp Airn sequence was calculated using a 100-nucleotide

2 sliding window as described 20. Briefly, the number of Gs and Cs was counted in each given window. The G/C

3 Skew was calculated as (G-C) / (G+C).

5 Templates for helicase assays

6 Individual oligonucleotides for helicase template preparation (Table 3) were gel-isolated using the crush

7 and soak method: Oligonucleotides were electrophoresed on 8 % denaturing polyacrylamide gels at 120 V for

8 1.5 h in 1 x TBE buffer. The bands were excised, crushed into a fine paste, and soaked overnight in an

9 Eppendorf ThermoMixer at 37 oC and 1,400 rpm in 500 mM ammonium acetate / 10 mM magnesium acetate /

10 1 mM EDTA. The extracted oligonucleotides were ethanol-precipitated twice and stored in TE pH 8.0.

11 Radiolabeling of oligonucleotides was carried out for 1 h at 37 oC using T4 polynucleotide kinase

12 (NEB), 0.25 μM oligonucleotide, and 0.5 μM -[32P]-ATP. Reactions were terminated by incubation for 20 min

13 at 80 oC. Following the radiolabeling, oligonucleotide annealing was carried out in a thermocycler in T4

14 polynucleotide kinase reaction buffer (NEB) / 50 mM potassium acetate by heating oligonucleotide mixtures to

15 95 oC for 5 minutes, followed by cooling at a rate of 1 oC / minute until the temperature reached 10 oC.

16 Annealed products were fractionated by 10 % native PAGE at 150 V for 45 min in 0.5 x TBE, corresponding

17 bands excised from the gel, and templates eluted by soaking the excised gel slices overnight in buffer

18 containing 10 mM Tris-HCl pH 7.2 (for templates with RNA) or pH 8.0 (for templates with DNA) / 50 mM

19 potassium acetate / 1 mM EDTA.

21 Gel-analysis of R-loop templates

22 For Figure 1D, 0.5 g of R-loop-containing plasmid was incubated with 0.5 U RNase H (NEB) in 75 mM

o 23 KCl / 50 mM Tris-HCl pH 7.5 / 3 mM MgCl2 / 10 mM DTT in a 20 L reaction volume and incubated at 37 C for

24 30 minutes. The reaction was quenched with 40 mM EDTA. Half the reaction was added to loading dye (10

25 mM Tris-HCl pH 7.6 / 0.15% orange G / 60% glycerol / 60 mM EDTA) and analyzed by 1 % agarose gel-

26 electrophoresis in TAE. The gel was run at 40 V for 5 hours and stained with Ethidium Bromide.

27 For Figure 1E, 0.5 g of 32P-labelled R-loop-containing plasmid was either mock-treated or incubated

28 with 0.5 U RNase H (NEB) in 75 mM KCl / 50 mM Tris-HCl pH 7.5 / 3 mM MgCl2 / 10 mM DTT in a 20 L 28 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 reaction volume for 30 minutes at 37 oC. The reaction was quenched with 40 mM EDTA. Half the reaction was

2 added to denaturing loading buffer (50 % formamide / 6 % formaldehyde / 10 % glycerol / 0.01 % Bromophenol

3 Blue) and incubated for 2 minutes at 90 oC, cooled on ice, and fractionated on a 1 % agarose gel containing

4 0.7 % formaldehyde. The gel was washed in 1x SSC, dried on Whatman paper, and analyzed by phosphor

5 imaging.

7 Electron Microscopy

8 Plasmid templates for all EM analyses were linearized with Msc I (NEB).

10 T4 gene 32 protein (gp32) staining.

11 To stain the displaced single strand (ss) DNA of the R-loop with gp32 (gift of Nancy Nossal, NIH,

12 Bethesda MD), linearized template plasmid was mixed with gp32 at a ratio of 1 g of gp32 per g of DNA in a

13 binding buffer of 10 mM Hepes pH 7.5 / 1 mM EDTA / 50 mM NaCl on ice for 30 minutes. The sample was

14 then fixed with 0.6% glutaraldehyde for 5 minutes on ice followed by surface spreading the DNA on a droplet of

15 0.25 mM ammonium acetate, with 7 g /mL of cytochrome C protein and allowed to develop for 3 min followed

16 by picking up the surface film with a Parlodion coated 400 mesh copper grids. The grids were then rotary

17 shadow cast with 80 % platinum – 20 % palladium (EMS sciences inc) followed by an overlayer of carbon.

19 S9.6 antibody staining.

20 Linearized template plasmid DNA was mixed with the S9.6 antibody (Santi Cruz Biotechnology inc.) at

21 a ratio of 1 g of antibody per g of DNA in the binding buffer above, fixed, and prepared for EM as described

22 for gp32 staining.

24 Formamide spreading

25 DNA spreads were prepared according to Lopes 102. Typically, 5 µl of DNA corresponding to 5–20 ng

26 were used for each spread. The DNA was mixed with 5 µl of formamide (Thermo Scientific, 17899) and 0.4 µl

27 of 0.02 % benzalkonium chloride (BAC, Sigma B6285) in 10 mM Tris, 1 mM EDTA pH 7.5. After mixing, the

28 drop was immediately spread on a water surface in a 15 cm dish containing 50 ml of distilled water, using a 29 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 freshly cleaved mica sheet (Ted Pella Inc.) as a ramp. The monomolecular layer was gently touched with a

2 carbon-covered 400-mesh copper grid, activated just before use by floating it on a drop of ethidium bromide

3 solution (33.3 µg / ml in H2O), for 30–45 min at RT. Grids with adsorbed DNA molecules were immediately

4 stained with a solution of 0.2 µg / µl uranyl acetate in ethanol and transferred to a Denton evaporator where the

5 sample was rotary shadow cast with platinum at a vacuum of 2 x 10-6 torr and at an angle of 3 between the

6 sample and the platinum wire.

8 Distance measurements.

9 ImageJ was used to measure the distance from the short end of the DNA to the start of the loop. At

10 least 100 images each for CD and HO templates were used for distance measurements. Distances were

11 plotted and frequency distribution analyses were performed using GraphPad Prism.

13 Atomic Force Microscopy

14 A 50 L reaction containing 10 nM linearized R-loop-containing plasmid / 22 nM RPA / 5 % glycerol /

15 0.3 mM ATP / 10 mM Mg(OAc)2 / 100 mM KOAc / 25 mM Hepes-KOH pH 7.5 was incubated on ice for 3

16 minutes. 40 l of this reaction were deposited onto freshly cleaved mica for 2 minutes. The sample was rinsed

17 with 10 ml ultrapure deionized water and the surface was dried using a stream of nitrogen. AFM images were

18 captured using an Asylum Research MFP-3D-BIO (Oxford Instruments) microscope in tapping mode at room

19 temperature. An Olympus AC240TS-R3 AFM probe with resonance frequencies of approximately 70 kHz and

20 spring constant of approximately 1.7 Nm−1 was used for imaging. Images were collected at a speed of 0.5–1Hz

21 with an image size of 2-3 μm at a 2048 × 2048 pixel resolution.

23 Proteins

24 T7 lysozyme

25 T7 lysozyme was expressed as C-terminally His-tagged fusion protein in BL21 DE3 RIL cells. 1L of

26 cells was grown to log phase, induced with 1 mM IPTG at 37 oC for 3 hours. Cells were harvested by

27 centrifugation for 30 minutes at 4,000 rpm and resuspended in buffer L (50 mM Tris-HCl pH 7.6 / 10% glycerol

28 / 0.02% NP-40) / 300 mM NaCl / protease inhibitor cocktail / 1 mM DTT. Cells were lysed by incubation with 30 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 0.1 mg / ml lysozyme for 30 minutes on ice followed by sonication. Cleared lysate was obtained by

2 centrifugation at 15,000 rpm in an SS34 rotor for 30 minutes at 4 oC. The clarified extract was supplemented

3 with 10 mM imidazole and rotated for 2 hours at 4 oC with 1 ml of Ni-NTA beads (Qiagen). Bound protein was

4 eluted with 10 CV of buffer L / 200 mM imidazole. Peak fractions were pooled, diluted with Buffer L to lower the

5 salt concentration to 10 mM NaCl, and fractionated over a 1 ml MonoS column using a 20 CV gradient of 10 -

6 125 mM NaCl in buffer L. Peak fractions were pooled, concentrated using Amicon Ultra 4 ml centrifugal filters

7 (Millipore) and fractionated over 24 ml S200 gel filtration column equilibrated in buffer L / 125 mM NaCl. Peak

8 fractions were pooled, aliquoted, flash frozen and stored at -80 oC.

10 RNase H1

11 RNase H1 was expressed as an N-terminal 6x-His tag fusion protein in BL21 DE3 RIL cells. 1L of cells

12 was grown to log phase and induced with 1 mM IPTG at 16 oC overnight. Cells were harvested by

13 centrifugation and resuspended in buffer A (25 mM Potassium phosphate pH 7.5 / 10% glycerol / 0.02% NP-

14 40) / 500 mM KCl / 1 mM DTT / protease inhibitors. Cells were lysed by addition of 0.1 mg / ml lysozyme,

15 incubation for 30 minutes on ice, followed by sonication. Insoluble material was removed by centrifugation at

16 15,000 rpm in an SS35 rotor for 30 minutes at 4 oC. Clarified extract was supplemented with 10 mM imidazole,

17 passed over 1 mL Ni-NTA resin (Qiagen), and bound protein eluted with 10 CV of buffer A / 200 mM imidazole.

18 Peak fractions were pooled, dialyzed against buffer B (25mM Hepes-KOH pH 7.5 / 10% glycerol / 1 mM EDTA

19 / 1 mM EGTA / 0.02% NP-40) / 0.1 M KCl / 1 mM DTT, and fractionated over a 1 mL MonoS column using a

20 salt gradient of 0.1 - 1 M KCl over 20 CV. Peak fractions were pooled, concentrated in an Amicon Ultra 4 mL

21 centrifugal filter, and fractionated over a 24 mL S200 gel-filtration column equilibrated in buffer B / 0.3 M KOAc

22 / 1 mM DTT. Peak fractions were pooled, aliquoted, flash frozen and stored at -80 oC.

24 RNase H2

25 Strain YDR154 was grown at 30 oC in YP-GL (YP + 2 % glycerol / 2 % lactic acid) to a density of 5 x

26 107 cells / ml, and protein expression was induced by addition of 2 % galactose for 4 hours. Cells were

27 harvested by centrifugation and washed once with cold 25 mM Hepes-KOH pH 7.6 / 1 M sorbitol, and once

28 with with buffer C (25 mM Hepes-KOH pH 7.6 / 0.02 % NP-40 / 10 % glycerol / 1 mM DTT) / 100 mM KCl. The 31 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 cell pellet was resuspended in 0.5 volumes of buffer C / 100 mM KCl / protease inhibitor, and frozen dropwise

2 in liquid nitrogen. The frozen cell popcorn was crushed in a freezer mill (SPEX CertiPrep 6850 Freezer/Mill) for

3 10 cycles of 2 minutes at a rate of 15 impacts per second. The crushed cell powder was thawed on ice and

4 mixed with 1 volume of buffer C / 100 mM KCl / protease inhibitor. The KCl concentration was adjusted to 0.4

5 M and the cell lysate rotated for 30 min at 4 oC. Insoluble material was precipitated by ultracentrifugation for 1

6 hour at 40,000 rpm in a T647.5 rotor (Thermo Scientific). The clarified cell lysate was supplemented with 2 mM

o 7 CaCl2 and rotated for 2-3 hours at 4 C with 1 ml of calmodulin affinity resin. The resin was collected in a

8 disposable column, washed with 10 CV of buffer C / 2 mM CaCl2 / 500 mM KCl, and bound protein eluted in 6

9 CV of buffer C / 500 mM KCl / 1 mM EDTA / 2 mM EGTA. Peak fractions were pooled, dialyzed against buffer

10 C / 100 mM KCl / 1 mM EDTA / 1 mM EGTA, and fractionated on a 1 ml Mono S column using a gradient of

11 0.1 M – 1 M KCl in buffer C / 1 mM EDTA / 1 mM EGTA. Peak fractions were pooled and gel-filtered on a 24

12 ml S200 column equilibrated in buffer C / 0.3 M KOAc / 1 mM EDTA / 1 mM EGTA / 1 mM DTT. Peak fractions

13 were pooled and stored in aliquots at -80 oC.

15 Pif1

16 The nuclear form (amino acids 40-859) of Pif1 was expressed as an N-terminally 6x-His tagged fusion

17 protein in BL21 DE3 RIL cells. 2 L of cells were grown to log phase and induced with 0.1 mM IPTG for 18

18 hours at 16 oC. Cells were harvested by centrifugation, resuspended in 50 ml Buffer A / 100 mM KCl / 1 mM

19 DTT / protease inhibitors, and cells lysed by addition of 0.1 mg/ml lysozyme, incubation for 30 minutes on ice,

20 and subsequent sonication. Lysate was clarified by centrifugation in an SS34 rotor at 15,000 rpm for 30

21 minutes at 4 oC. Clarified extract was passed over a 5 mL Q sepharose column, the flow-through applied to a 5

22 mL SP-Sepharose column, and bound protein eluted with a salt gradient of 150 - 660 mM KCl in buffer A over

23 10 CV. Subsequent purification steps were as described before 103. Pif1-N-E303Q was purified as described

24 above.

26 CMG

27 Strain YSD35 was grown at 30 oC in YP-GL (YP + 2 % glycerol / 2 % lactic acid) to a density of 5 x 107

28 cells / ml, and protein expression was induced by addition of 2 % galactose for 6 hours. Cells were harvested 32 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 by centrifugation and washed once with 25 mM HEPES-KOH pH-7.6 / 1 M sorbitol and once with buffer D (40

2 mM HEPES-NaOH pH-7.6 / 150 mM sodium acetate / 10 % glycerol / 0.005% Tween 20). The cell pellet was

3 resuspended in 0.5 volumes of buffer D / 1 mM DTT / protease inhibitor, and frozen dropwise in liquid nitrogen.

4 The frozen cell popcorn was crushed in a freezer mill (SPEX CertiPrep 6850 Freezer/Mill) for 10 cycles of 2

5 minutes at a rate of 15 impacts per second. The crushed cell powder was thawed on ice and mixed with 0.5

6 volumes of buffer D / 1 mM DTT. The cell lysate was centrifuged for 1 h at 40,000 rpm in T647.5 rotor (Thermo

7 Scientific) to sediment insoluble material. The clarified extract was passed over 6 ml of FLAG M2-affinity beads

8 (Sigma) using the Biorad Econo pumping system. The beads were washed with 10 CV of buffer D / 1 mM DTT,

9 followed by washes with 20 ml of buffer D / 1 mM DTT / 5 mM magnesium acetate / 0.5 mM ATP and 10 CV of

10 buffer D / 1 mM DTT. Bound protein was eluted with 1 CV of buffer D / 1 mM DTT / 2 mM CaCl2 / 0. 2 mg / ml

11 3xFLAG peptide, followed by 2 CV of buffer D / 1 mM DTT / 2 mM calcium chloride / 0. 1 mg / ml 3xFLAG

12 peptide. The eluates were pooled and applied to 1 ml of calmodulin affinity resin (Agilent Technologies). Beads

13 were washed with 20 CV of buffer D / 1 mM DTT / 2 mM CaCl2 and bound protein eluted with 8 CV of buffer D

14 / 1 mM DTT / 2 mM EGTA / 2 mM EDTA. The eluates from the calmodulin beads were further fractionated on a

15 MiniQ PC 3.2/3 anion exchange column equilibrated in 25 mM Tris pH-7.2 / 10% glycerol / 0.005% Tween 20 /

16 1 mM DTT / 150 mM KCl. Bound protein was eluted with a 30 CV gradient from 150 – 1000 mM KCl in 25 mM

17 Tris pH-7.2 / 10% glycerol / 0.005% Tween 20 / 1 mM DTT. CMG-containing peak fractions were pooled,

18 dialyzed against 2 L of 25 mM HEPES-KOH pH-7.6 / 200 mM potassium acetate / 2 mM magnesium acetate /

19 10 % glycerol / 1 mM DTT, aliquoted, flash frozen in liquid nitrogen, and stored at -80 oC.

21 DNA replication assays

22 All replication reactions were performed on linearized template with or without R-loop. 4 nM template

23 DNA was incubated with 5 U NsiI in an 8 uL reaction containing 50 mM KOAc, 25 mM Hepes-KOH pH 7.5, 5%

24 glycerol, 2.5 mM DTT, 0.02 % NP-40, and incubated for 30 minutes at 37 oC.

25 The salt concentration was increased to 100 mM KOAc. 16 nM ORC, 50 nM Cdc6, 100 nM Cdt1Mcm2-

26 7, and 5 nM ATP were added to the reaction and incubation continued for 20 minutes at 30 oC. 150 nM of DDK

27 was added and incubation continued for 20 minutes at 30 oC. Subsequently a mastermix was added containing

28 50 g of BSA, 20 nM Sld3-7, 125 nM Cdc45, 80 nM each dNTP, 16 nM CDK, 100 nM GINS, 20 nM Pol , 8 nM 33 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 Dpb11, 30 nM Sld2, 150 nM RPA, 75 nM Pol , 150 nM Ctf4, 25 nM RFC, 75 nM PCNA, 2 nM Pol , 0.2 mM

2 each rNTP, 30 nM Top1, 20 nM Mrc1, 20 nM Csm3-Tof1. The salt concentration was increased to 180 mM

3 KOAc and origin firing was induced by the addition of 14 nM Mcm10 in the presence of 5 Ci -[32P]-dATP and

4 incubation continued for one hour at 30 oC. Reactions were stopped by addition of 40 mM EDTA, 0.8 U

5 Proteinase K ,and 0.8 % SDS, followed by incubation for 30 minutes at 37oC. Reactions were

6 phenol/chloroform extracted and filtered through Illustra MicroSpin G-25 spin columns (GE).

7 For denaturing gels, 10 L of each sample was digested with ClaI (Neb) in 1x CutSmart buffer and

8 subsequently quenched with 40 mM EDTA and loading dye. Digested samples were fractionated on 0.8 %

9 agarose gels in 30 mM NaOH / 2 mM EDTA. Denaturing gels were neutralized and fixed in 5% TCA. For

10 native gels, 10 L of undigested sample were supplemented with loading dye and fractionated on 0.8% native

11 agarose gels in TAE. Denaturing and native gels were dried on Whatman paper and analyzed by phosphor

12 imaging. Signal intensities were quantified using ImageJ.

13 For conventional time course experiments, aliquots were removed at desired times after the addition of

14 Mcm10 and reactions stopped with 40 mM EDTA. For pulse-chase experiments, the concentration of dATP

15 was reduced to 2 nM. Reactions were pulsed for 2.5 minutes by adding 5 Ci of -[32P]-dATP along with

16 Mcm10 and chased by addition of 1 mM cold dATP. Aliquots were removed and quenched with 40 mM EDTA

17 at desired time points.

18 Unless indicated otherwise, Pif1 (0.7 nM), RNase H1 (1 nM), pyridostatin (0.5 M; Sigma), Fen1 (15

19 nM), and Cdc9 (15 nM) were added to the reactions alongside Mcm10. For treatment of templates with T4

20 PNK, 5 U of T4 PNK (NEB) were included in the initial NsiI digestion reaction and incubated at 37 oC for one

21 hour.

22 For two-dimensional gel analyses, a 20 L standard replication reaction was digested with ClaI,

23 fractionated in the first dimension by 0.85 % native agarose gel-electrophoresis, the sample lane excised from

24 the native gel and fractionated on a denaturing agarose gel overnight with buffer circulation in 30 mM NaOH / 2

25 mM EDTA. The gel was neutralized by shaking in 5% TCA, dried on Whatman paper, and analyzed by

26 phosphor imaging.

34 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 Polymerase assay

2 4 nM template was incubated in a 16 L reaction volume with 0.5 U T4 PNK and 5 U NsiI in 50 mM

3 KOAc / 25 mM Hepes-KOH pH 7.5 / 5% glycerol / 2.5 mM DTT / 0.02 % NP-40 and incubated for one hour at

4 37 oC. Subsequently, the reaction was supplemented with 5 nM ATP, 50 g of BSA, 80 nM each dNTP, 150

5 nM RPA, 25 nM RFC, 75 nM PCNA, 0.2 mM each rNTP. Reactions were supplemented with 5 Ci -[32P]-

6 dATP, followed by addition of Pol  (20 nM), Pol  (75 nM), and Pol  (2 nM), as indicated, and incubated for

7 one hour at 30 oC. Reactions were stopped by addition of 40 mM EDTA, 0.8 U Proteinase K ,and 0.8 % SDS,

8 followed by incubation for 30 minutes at 37oC. Reactions were phenol/chloroform extracted and filtered through

9 Illustra MicroSpin G-25 spin columns (GE). 10 L of each reaction sample was fractionated by denaturing

10 agarose gel-electrophoresis.

12 CMG helicase assays

13 Assays were performed in helicase assay buffer (20 mM HEPES-KOH pH-7.6 / 100 mM potassium

14 acetate / 10 mM magnesium acetate / 0.1 mg / ml BSA / 2.5 mM DTT). For time course analyses, 25 l

15 reactions containing 4 nM templates and 20 nM CMG were supplemented with 100 M ATPS and incubated

16 at 30 oC for 1 h. Unwinding was initiated by addition of 25 l of helicase assay buffer / 10 mM ATP to the

17 reaction mixture. Incubation was continued at 30 oC and 6 l aliquots were withdrawn at indicated time points.

18 The reactions were stopped by adding 20 mM EDTA and 0.12 % SDS. Reaction products were fractionated by

19 10 % native PAGE in a Biorad Mini-PROTEAN system at 75 V for 2 hours in 0.5x TAE buffer. Gels were dried

20 on a Whatman paper, and exposed to phosphorimager screen, and scanned on a Typhoon 7000 imager (GE

21 Healthcare). The images were quantified using ImageJ and plotted using GraphPad Prism software.

22 For assays in Figure 4B, 5 l reactions containing 4 nM template, 20 nM CMG, and 100 M ATPS

23 were incubated for 1 hour at 30 oC. Unwinding was initiated by adding 5 l of helicase assay buffer / 10 mM

24 ATP. Reaction were stopped after 15 minutes by adding 10 l of 40 mM EDTA and 0.24 % SDS.

35 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 REFERENCES

2 1. Crossley, M.P., Bocek, M. & Cimprich, K.A. R-Loops as Cellular Regulators and Genomic Threats. Mol 3 Cell 73, 398-411 (2019). 4 2. Garcia-Muse, T. & Aguilera, A. R Loops: From Physiological to Pathological Roles. Cell 179, 604-618 5 (2019). 6 3. Kabeche, L., Nguyen, H.D., Buisson, R. & Zou, L. A mitosis-specific and R loop-driven ATR pathway 7 promotes faithful chromosome segregation. Science 359, 108-114 (2018). 8 4. Skourti-Stathaki, K., Proudfoot, N.J. & Gromak, N. Human senataxin resolves RNA/DNA hybrids formed 9 at transcriptional pause sites to promote Xrn2-dependent termination. Mol Cell 42, 794-805 (2011). 10 5. Yu, K., Chedin, F., Hsieh, C.L., Wilson, T.E. & Lieber, M.R. R-loops at immunoglobulin class switch 11 regions in the chromosomes of stimulated B cells. Nat Immunol 4, 442-51 (2003). 12 6. Graf, M. et al. Telomere Length Determines TERRA and R-Loop Regulation through the Cell Cycle. Cell 13 170, 72-85 e14 (2017). 14 7. Niehrs, C. & Luke, B. Regulatory R-loops as facilitators of gene expression and genome stability. Nat 15 Rev Mol Cell Biol 21, 167-178 (2020). 16 8. Gan, W. et al. R-loop-mediated genomic instability is caused by impairment of replication fork 17 progression. Genes Dev 25, 2041-56 (2011). 18 9. Gomez-Gonzalez, B. et al. Genome-wide function of THO/TREX in active genes prevents R-loop- 19 dependent replication obstacles. EMBO J 30, 3106-19 (2011). 20 10. Tuduri, S. et al. Topoisomerase I suppresses genomic instability by preventing interference between 21 replication and transcription. Nat Cell Biol 11, 1315-24 (2009). 22 11. Wellinger, R.E., Prado, F. & Aguilera, A. Replication fork progression is impaired by transcription in 23 hyperrecombinant yeast cells lacking a functional THO complex. Mol Cell Biol 26, 3327-34 (2006). 24 12. Stork, C.T. et al. Co-transcriptional R-loops are the main cause of estrogen-induced DNA damage. Elife 25 5(2016). 26 13. Costantino, L. & Koshland, D. Genome-wide Map of R-Loop-Induced Damage Reveals How a Subset of 27 R-Loops Contributes to Genomic Instability. Mol Cell 71, 487-497 e3 (2018). 28 14. Hamperl, S., Bocek, M.J., Saldivar, J.C., Swigut, T. & Cimprich, K.A. Transcription-Replication Conflict 29 Orientation Modulates R-Loop Levels and Activates Distinct DNA Damage Responses. Cell 170, 774-786 30 e19 (2017). 31 15. Sanz, L.A. et al. Prevalent, Dynamic, and Conserved R-Loop Structures Associate with Specific 32 Epigenomic Signatures in Mammals. Mol Cell 63, 167-78 (2016). 33 16. Wahba, L., Costantino, L., Tan, F.J., Zimmer, A. & Koshland, D. S1-DRIP-seq identifies high expression 34 and polyA tracts as major contributors to R-loop formation. Genes Dev 30, 1327-38 (2016). 35 17. Wahba, L., Amon, J.D., Koshland, D. & Vuica-Ross, M. RNase H and multiple RNA biogenesis factors 36 cooperate to prevent RNA:DNA hybrids from generating genome instability. Mol Cell 44, 978-88 (2011). 37 18. Chan, Y.A. et al. Genome-wide profiling of yeast DNA:RNA hybrid prone sites with DRIP-chip. PLoS 38 Genet 10, e1004288 (2014). 39 19. El Hage, A., Webb, S., Kerr, A. & Tollervey, D. Genome-wide distribution of RNA-DNA hybrids identifies 40 RNase H targets in tRNA genes, retrotransposons and mitochondria. PLoS Genet 10, e1004716 (2014). 41 20. Ginno, P.A., Lott, P.L., Christensen, H.C., Korf, I. & Chedin, F. R-loop formation is a distinctive 42 characteristic of unmethylated human CpG island promoters. Mol Cell 45, 814-25 (2012). 43 21. Huertas, P. & Aguilera, A. Cotranscriptionally formed DNA:RNA hybrids mediate transcription 44 elongation impairment and transcription-associated recombination. Mol Cell 12, 711-21 (2003). 45 22. Li, X. & Manley, J.L. Inactivation of the SR protein splicing factor ASF/SF2 results in genomic instability. 46 Cell 122, 365-78 (2005).

36 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 23. Garcia-Pichardo, D. et al. Histone Mutants Separate R Loop Formation from Genome Instability 2 Induction. Mol Cell 66, 597-609 e5 (2017). 3 24. Malig, M., Hartono, S.R., Giafaglione, J.M., Sanz, L.A. & Chedin, F. Ultra-deep Coverage Single-molecule 4 R-loop Footprinting Reveals Principles of R-loop Formation. J Mol Biol 432, 2271-2288 (2020). 5 25. Roberts, R.W. & Crothers, D.M. Stability and properties of double and triple helices: dramatic effects of 6 RNA or DNA backbone composition. Science 258, 1463-6 (1992). 7 26. Duquette, M.L., Handa, P., Vincent, J.A., Taylor, A.F. & Maizels, N. Intracellular transcription of G-rich 8 DNAs induces formation of G-loops, novel structures containing G4 DNA. Genes Dev 18, 1618-29 9 (2004). 10 27. De Magis, A. et al. DNA damage and genome instability by G-quadruplex ligands are mediated by R 11 loops in human cancer cells. Proc Natl Acad Sci U S A 116, 816-825 (2019). 12 28. Burge, S., Parkinson, G.N., Hazel, P., Todd, A.K. & Neidle, S. Quadruplex DNA: sequence, topology and 13 structure. Nucleic Acids Res 34, 5402-15 (2006). 14 29. Paeschke, K. et al. Pif1 family helicases suppress genome instability at G-quadruplex motifs. Nature 15 497, 458-62 (2013). 16 30. Ribeyre, C. et al. The yeast Pif1 helicase prevents genomic instability caused by G-quadruplex-forming 17 CEB1 sequences in vivo. PLoS Genet 5, e1000475 (2009). 18 31. Sarkies, P., Reams, C., Simpson, L.J. & Sale, J.E. Epigenetic instability due to defective replication of 19 structured DNA. Mol Cell 40, 703-13 (2010). 20 32. Dahan, D. et al. Pif1 is essential for efficient replisome progression through lagging strand G- 21 quadruplex DNA secondary structures. Nucleic Acids Res 46, 11847-11857 (2018). 22 33. Lopes, J. et al. G-quadruplex-induced instability during leading-strand replication. EMBO J 30, 4033-46 23 (2011). 24 34. Lerner, L.K. & Sale, J.E. Replication of G Quadruplex DNA. Genes (Basel) 10(2019). 25 35. Belotserkovskii, B.P. et al. Mechanisms and implications of transcription blockage by guanine-rich DNA 26 sequences. Proc Natl Acad Sci U S A 107, 12816-21 (2010). 27 36. Castellano-Pozo, M. et al. R loops are linked to histone H3 S10 phosphorylation and chromatin 28 condensation. Mol Cell 52, 583-90 (2013). 29 37. Garcia-Rubio, M. et al. Yra1-bound RNA-DNA hybrids cause orientation-independent transcription- 30 replication collisions and telomere instability. Genes Dev 32, 965-977 (2018). 31 38. Sollier, J. et al. Transcription-coupled nucleotide excision repair factors promote R-loop-induced 32 genome instability. Mol Cell 56, 777-85 (2014). 33 39. Su, X.A. & Freudenreich, C.H. Cytosine deamination and base excision repair cause R-loop-induced CAG 34 repeat fragility and instability in Saccharomyces cerevisiae. Proc Natl Acad Sci U S A 114, E8392-E8401 35 (2017). 36 40. Devbhandari, S., Jiang, J., Kumar, C., Whitehouse, I. & Remus, D. Chromatin Constrains the Initiation 37 and Elongation of DNA Replication. Mol Cell 65, 131-141 (2017). 38 41. Yeeles, J.T., Deegan, T.D., Janska, A., Early, A. & Diffley, J.F. Regulated eukaryotic DNA replication origin 39 firing with purified proteins. Nature 519, 431-5 (2015). 40 42. Devbhandari, S. & Remus, D. Rad53 limits CMG helicase uncoupling from DNA synthesis at replication 41 forks. Nat Struct Mol Biol 27, 461-471 (2020). 42 43. Carrasco-Salas, Y. et al. The extruded non-template strand determines the architecture of R-loops. 43 Nucleic Acids Res 47, 6783-6795 (2019). 44 44. Drolet, M., Bi, X. & Liu, L.F. Hypernegative supercoiling of the DNA template during transcription 45 elongation in vitro. J Biol Chem 269, 2068-74 (1994). 46 45. Stolz, R. et al. Interplay between DNA sequence and negative superhelicity drives R-loop structures. 47 Proc Natl Acad Sci U S A 116, 6260-6269 (2019).

37 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 46. Kumar, A. & Patel, S.S. Inhibition of T7 RNA polymerase: transcription initiation and transition from 2 initiation to elongation are inhibited by T7 lysozyme via a ternary complex with RNA polymerase and 3 promoter DNA. Biochemistry 36, 13954-62 (1997). 4 47. Zhang, X. & Studier, F.W. Mechanism of inhibition of bacteriophage T7 RNA polymerase by T7 5 lysozyme. J Mol Biol 269, 10-27 (1997). 6 48. Boguslawski, S.J. et al. Characterization of monoclonal antibody to DNA.RNA and its application to 7 immunodetection of hybrids. J Immunol Methods 89, 123-30 (1986). 8 49. Aria, V. & Yeeles, J.T.P. Mechanism of Bidirectional Leading-Strand Synthesis Establishment at 9 Eukaryotic DNA Replication Origins. Mol Cell (2018). 10 50. Taylor, M.R.G. & Yeeles, J.T.P. Dynamics of Replication Fork Progression Following Helicase-Polymerase 11 Uncoupling in Eukaryotes. J Mol Biol 431, 2040-2049 (2019). 12 51. Fu, Y.V. et al. Selective bypass of a lagging strand roadblock by the eukaryotic replicative DNA helicase. 13 Cell 146, 931-41 (2011). 14 52. Kose, H.B., Larsen, N.B., Duxin, J.P. & Yardimci, H. Dynamics of the Eukaryotic Replicative Helicase at 15 Lagging-Strand Protein Barriers Support the Steric Exclusion Model. Cell Rep 26, 2113-2125 e6 (2019). 16 53. Bando, M. et al. Csm3, Tof1, and Mrc1 form a heterotrimeric mediator complex that associates with 17 DNA replication forks. J Biol Chem 284, 34355-65 (2009). 18 54. Gellon, L. et al. Mrc1 and Tof1 prevent fragility and instability at long CAG repeats by their fork 19 stabilizing function. Nucleic Acids Res 47, 794-805 (2019). 20 55. Katou, Y. et al. S-phase checkpoint proteins Tof1 and Mrc1 form a stable replication-pausing complex. 21 Nature 424, 1078-83 (2003). 22 56. Szyjka, S.J., Viggiani, C.J. & Aparicio, O.M. Mrc1 is required for normal progression of replication forks 23 throughout chromatin in S. cerevisiae. Mol Cell 19, 691-7 (2005). 24 57. Tourriere, H., Versini, G., Cordon-Preciado, V., Alabert, C. & Pasero, P. Mrc1 and Tof1 promote 25 replication fork progression and recovery independently of Rad53. Mol Cell 19, 699-706 (2005). 26 58. Yeeles, J.T.P., Janska, A., Early, A. & Diffley, J.F.X. How the Eukaryotic Replisome Achieves Rapid and 27 Efficient DNA Replication. Mol Cell 65, 105-116 (2017). 28 59. Calzada, A., Hodgson, B., Kanemaki, M., Bueno, A. & Labib, K. Molecular anatomy and regulation of a 29 stable replisome at a paused eukaryotic DNA replication fork. Genes Dev 19, 1905-19 (2005). 30 60. Hodgson, B., Calzada, A. & Labib, K. Mrc1 and Tof1 regulate DNA replication forks in different ways 31 during normal S phase. Mol Biol Cell 18, 3894-902 (2007). 32 61. Stith, C.M., Sterling, J., Resnick, M.A., Gordenin, D.A. & Burgers, P.M. Flexibility of eukaryotic Okazaki 33 fragment maturation through regulated strand displacement synthesis. J Biol Chem 283, 34129-40 34 (2008). 35 62. Duan, X.L. et al. G-quadruplexes significantly stimulate Pif1 helicase-catalyzed duplex DNA unwinding. J 36 Biol Chem 290, 7722-35 (2015). 37 63. Sanders, C.M. Human Pif1 helicase is a G-quadruplex DNA-binding protein with G-quadruplex DNA- 38 unwinding activity. Biochem J 430, 119-28 (2010). 39 64. Zhou, R., Zhang, J., Bochman, M.L., Zakian, V.A. & Ha, T. Periodic DNA patrolling underlies diverse 40 functions of Pif1 on R-loops and G-rich DNA. Elife 3, e02190 (2014). 41 65. Byrd, A.K. & Raney, K.D. A parallel quadruplex DNA is bound tightly but unfolded slowly by pif1 42 helicase. J Biol Chem 290, 6482-94 (2015). 43 66. Hou, X.M. et al. Molecular mechanism of G-quadruplex unwinding helicase: sequential and repetitive 44 unfolding of G-quadruplex by Pif1 helicase. Biochem J 466, 189-99 (2015). 45 67. Wallgren, M. et al. G-rich telomeric and ribosomal DNA sequences from the fission yeast genome form 46 stable G-quadruplex DNA structures in vitro and are unwound by the Pfh1 DNA helicase. Nucleic Acids 47 Res 44, 6213-31 (2016).

38 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 68. Paeschke, K., Capra, J.A. & Zakian, V.A. DNA replication through G-quadruplex motifs is promoted by 2 the Saccharomyces cerevisiae Pif1 DNA helicase. Cell 145, 678-91 (2011). 3 69. Boule, J.B. & Zakian, V.A. The yeast Pif1p DNA helicase preferentially unwinds RNA DNA substrates. 4 Nucleic Acids Res 35, 5809-18 (2007). 5 70. Chib, S., Byrd, A.K. & Raney, K.D. Yeast Helicase Pif1 Unwinds RNA:DNA Hybrids with Higher 6 Processivity than DNA:DNA Duplexes. J Biol Chem 291, 5889-5901 (2016). 7 71. Pohl, T.J. & Zakian, V.A. Pif1 family DNA helicases: A helpmate to RNase H? DNA Repair (Amst) 84, 8 102633 (2019). 9 72. Lee, C.Y. et al. R-loop induced G-quadruplex in non-template promotes transcription by successive R- 10 loop formation. Nat Commun 11, 3392 (2020). 11 73. Piazza, A. et al. Short loop length and high thermal stability determine genomic instability induced by 12 G-quadruplex-forming minisatellites. EMBO J 34, 1718-34 (2015). 13 74. Taylor, M.R.G. & Yeeles, J.T.P. The Initial Response of a Eukaryotic Replisome to DNA Damage. Mol Cell 14 70, 1067-1080 e12 (2018). 15 75. Kang, Y.H., Galal, W.C., Farina, A., Tappin, I. & Hurwitz, J. Properties of the human Cdc45/Mcm2-7/GINS 16 helicase complex and its action with DNA polymerase epsilon in rolling circle DNA synthesis. Proc Natl 17 Acad Sci U S A 109, 6042-7 (2012). 18 76. Langston, L. & O'Donnell, M. Action of CMG with strand-specific DNA blocks supports an internal 19 unwinding mode for the eukaryotic replicative helicase. Elife 6(2017). 20 77. Low, E., Chistol, G., Zaher, M.S., Kochenova, O.V. & Walter, J.C. The DNA replication fork suppresses 21 CMG unloading from chromatin before termination. Genes Dev 34, 1534-1545 (2020). 22 78. Vrtis, K.B. et al. Single-strand DNA breaks cause replisome disassembly. Mol Cell 81, 1309-1318 e6 23 (2021). 24 79. Edwards, D.N., Machwe, A., Wang, Z. & Orren, D.K. Intramolecular telomeric G-quadruplexes 25 dramatically inhibit DNA synthesis by replicative and translesion polymerases, revealing their potential 26 to lead to genetic change. PLoS One 9, e80664 (2014). 27 80. Sparks, M.A., Singh, S.P., Burgers, P.M. & Galletto, R. Complementary roles of Pif1 helicase and single 28 stranded DNA binding proteins in stimulating DNA replication through G-quadruplexes. Nucleic Acids 29 Res 47, 8595-8605 (2019). 30 81. Dehghani-Tafti, S., Levdikov, V., Antson, A.A., Bax, B. & Sanders, C.M. Structural and functional analysis 31 of the nucleotide and DNA binding activities of the human PIF1 helicase. Nucleic Acids Res 47, 3208- 32 3222 (2019). 33 82. Pomerantz, R.T. & O'Donnell, M. The replisome uses mRNA as a primer after colliding with RNA 34 polymerase. Nature 456, 762-6 (2008). 35 83. Hyjek, M., Figiel, M. & Nowotny, M. RNases H: Structure and mechanism. DNA Repair (Amst) 84, 36 102672 (2019). 37 84. Neil, A.J., Liang, M.U., Khristich, A.N., Shah, K.A. & Mirkin, S.M. RNA-DNA hybrids promote the 38 expansion of Friedreich's ataxia (GAA)n repeats via break-induced replication. Nucleic Acids Res 46, 39 3487-3497 (2018). 40 85. Pardo, B., Crabbe, L. & Pasero, P. Signaling pathways of replication stress in yeast. FEMS Yeast Res 41 17(2017). 42 86. Saldivar, J.C., Cortez, D. & Cimprich, K.A. The essential kinase ATR: ensuring faithful duplication of a 43 challenging genome. Nat Rev Mol Cell Biol 18, 622-636 (2017). 44 87. Schiavone, D. et al. PrimPol Is Required for Replicative Tolerance of G Quadruplexes in Vertebrate Cells. 45 Mol Cell 61, 161-9 (2016). 46 88. Guilliam, T.A. & Yeeles, J.T. The eukaryotic replisome tolerates leading-strand base damage by 47 replicase switching. EMBO J 40, e107037 (2021).

39 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

1 89. Guilliam, T.A. & Yeeles, J.T.P. Reconstitution of translesion synthesis reveals a mechanism of eukaryotic 2 DNA replication restart. Nat Struct Mol Biol 27, 450-460 (2020). 3 90. Perera, R.L. et al. Mechanism for priming DNA synthesis by yeast DNA polymerase alpha. Elife 2, 4 e00482 (2013). 5 91. Belotserkovskii, B.P., Tornaletti, S., D'Souza, A.D. & Hanawalt, P.C. R-loop generation during 6 transcription: Formation, processing and cellular outcomes. DNA Repair (Amst) 71, 69-81 (2018). 7 92. Lockhart, A. et al. RNase H1 and H2 Are Differentially Regulated to Process RNA-DNA Hybrids. Cell Rep 8 29, 2890-2900 e5 (2019). 9 93. Zimmer, A.D. & Koshland, D. Differential roles of the RNases H in preventing chromosome instability. 10 Proc Natl Acad Sci U S A 113, 12220-12225 (2016). 11 94. Stodola, J.L. & Burgers, P.M. Mechanism of Lagging-Strand DNA Replication in Eukaryotes. Adv Exp Med 12 Biol 1042, 117-133 (2017). 13 95. Tran, P.L.T. et al. PIF1 family DNA helicases suppress R-loop mediated genome instability at tRNA 14 genes. Nat Commun 8, 15025 (2017). 15 96. Prado, F. & Aguilera, A. Impairment of replication fork progression mediates RNA polII transcription- 16 associated recombination. EMBO J 24, 1267-76 (2005). 17 97. Nakamura, K. et al. H4K20me0 recognition by BRCA1-BARD1 directs homologous recombination to 18 sister chromatids. Nat Cell Biol 21, 311-318 (2019). 19 98. Saredi, G. et al. H4K20me0 marks post-replicative chromatin and recruits the TONSL-MMS22L DNA 20 repair complex. Nature 534, 714-718 (2016). 21 99. Wu, W. et al. RTEL1 suppresses G-quadruplex-associated R-loops at difficult-to-replicate loci in the 22 human genome. Nat Struct Mol Biol 27, 424-437 (2020). 23 100. Sparks, J.L. et al. The CMG Helicase Bypasses DNA-Protein Cross-Links to Facilitate Their Repair. Cell 24 176, 167-181 e21 (2019). 25 101. Gros, J., Devbhandari, S. & Remus, D. Origin plasticity during budding yeast DNA replication in vitro. 26 EMBO J 33, 621-36 (2014). 27 102. Lopes, M. Electron microscopy methods for studying in vivo DNA replication intermediates. Methods 28 Mol Biol 521, 605-31 (2009). 29 103. Wilson, M.A. et al. Pif1 helicase and Poldelta promote recombination-coupled DNA synthesis via 30 bubble migration. Nature 502, 393-6 (2013). 31

40 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission. Figure 1

A 0.6 B CD 0.4 (i) ARS305

0.2 replication 0 G/C skew - 0.2

- 0.4 473-495 510-523 1030-1056 3xT 3’ mAirn, 1.4 kbp HO 5’ (ii) ARS305 810-828

replication

R-loop ARS305 pARS Cla I (8.6 kbp) Nsi I

- + RNase H C D RNase H E pARSR-loop - - + - + + transcription T7 RNAP (transcribe)

+ R-loop Autorad T7 lysozyme nicked nt (stop re-initiation) supercoiled 1000 600 RNase A 400 (digest free RNA) 1 2 3 200 100 Proteinase K EtBr (deproteinize) 1 2 F Gel-filtration T4 gp32 S9.6 Formamide (isolate plasmid template) G

20 CD HO 15

10 requen cy f 5

0 0 1000 2000 3000 4000 Distance from Cla I (bp) mAirn ARS305

Cla I bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission. Figure 2 A B Cla I - + Cla I ARS305 CD kb mAirn 8.5 Right lead 803 925 2728 4102 8624 6.4 4.8 3.7

Cla I HO 2.3 ARS305 mAirn 1.4 Left lead 803 925 2728 8624 4102 Left lead, cut

denaturing 0.7 replicate Okazaki Cla I 0.2 fragments

0.8 kb 1 2 7.8 kb E CD HO

0.8 kb 1.9 kb 5.9 kb

0.8 kb 3.3 kb 4.5 kb Linear Uncoupled Left fork Linear Uncoupled Left fork Stalled fork / RI Stalled fork / RI C Native CD HO kb R-loop 8.5 Full-length +- +- 6.4

- + - + - + - + DDK Denaturing 4.8 3.7 Stalled fork / Stall 2.3 kbp RI 8.5 Linear native 6.4 Uncoupled 1.4 4.8 Left lead kb 0.7 8.5 Full-length Okazaki 6.4 Restart 0.2 4.8 fragments 3.7 2.3 Stall

1.4 CD Left lead F HO 0.7

denaturing - + - + R-loop + -- + -- + -- + -- Pol δ 0.2 Okazaki ++ - ++ - ++ - ++ - RFC/PCNA fragments Stalled fork / kbp RI 1 2 3 4 5 6 7 8 8.5 Linear native 6.4 Uncoupled D 4.8 kb mAirn 8.5 6.4 Full-length Fork 4.8 Restart 0.8 kb 3.7 stalling ~ 2.3 - 2.8 kb 2.3 Stall 1.4 Fork 0.8 kb uncoupling ~ 2.3 - 2.8 kb 0.7 Left lead denaturing 0.2 Fork Okazaki 0.8 kb fragments restart ~ 2.3 - 2.8 kb ~ 5.5 - 5.0 kb

1 2 3 4 5 6 7 8 9 10 11 12 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

Figure 3

AB CD HO C CD HO RNase H1 +- +- R-loop + +-- --+ + -- ++ - - ++ Pif1 - + - + - + - + RNase H1 Stalled fork /

RNase H1 Pif1-wt Pif1-E303Q Stalled fork / kbp RI kbp RI native native 8.5 Linear 8.5 Linear 6.4 Uncoupled Uncoupled kb Right lead FL kb 8.5 8.5 6.4 6.4 Full-length 4.8 Restart 4.8 3.7 3.7 Right lead stall 2.3 Stall 2.3

1.4 1.4 Left lead 0.7 0.7 Left lead denaturing

denaturing Okazaki 0.2 fragments 0.2 Okazaki fragments

1 2 3 4 5 6 7 8 1 2 3 4 5 6 7 8 D E CD

810-828 pCtrl HO wt wt ΔG4 AIRN - - + R-loop - + -- R-loop - +- +- RNase H1 +- +- -- RNase H1 - + - +-+ - +-+ pyridostatin - +-+ - +-+ - +-+ pyridostatin Stalled fork / Stalled fork / RI kbp RI 8.5 Linear kbp Linear native 6.4 Uncoupled 8.5 native 6.4 Uncoupled kb kb 8.5 Full-length 8.5 Full-length 6.4 6.4 4.8 Restart 4.8 Restart 3.7 3.7 2.3 Stall 2.3 Stall 1.4 Left lead 1.4 0.7 Left lead denaturing 0.7 Okazaki 0.2 fragments Okazaki denaturing 0.2 fragments 1 2 3 4 5 6 7 8 9 10

Figure 4

A CMG B (i) (ii) (iii) (iv) Boiled - CMG Boiled - CMG Boiled - CMG + CMG Boiled - CMG + CMG + CMG + CMG RNA Mcm2-7 DNA RNA DNA Cdc45

Sld5

Psf1-3 1 2 3 1 2 3 1 2 3 1 2 3

C DNA RNA DNA:DNA 80 RNA:DNA

- CMG - ATP - CMG - ATP 60

Boiled 60 0.5 1 2 3 5 15 60 60 min Boiled 60 0.5 1 2 3 5 15 60 60 min 40

20 % unwoun d

0 1 2 3 4 5 6 7 8 9 10 1 2 3 4 5 6 7 8 9 10 0 10 20 30 40 50 60 Time (min) D G4-wt, G4-mut, lead lead G4-wt, lead 80 G4-mut, lead - CMG - CMG Boiled 60 0.5 1 2 3 5 15 60 min Boiled 60 0.5 1 2 3 5 15 60 min 60

20 % unwoun d

0 10 20 30 40 50 60 1 2 3 4 5 6 7 8 9 1 2 3 4 5 6 7 8 9 Time (min)

G4-wt, lag E G4-wt, G4-mut, 80 G4-mut, lag lag lag - CMG 60 - CMG

Boiled 60 0.5 1 2 3 5 15 60 min Boiled 60 0.5 1 2 3 5 15 60 min 40

20 % unwoun d

0 10 20 30 40 50 60 1 2 3 4 5 6 7 8 9 1 2 3 4 5 6 7 8 9 Time (min) bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

Figure 5

A C D CD CD - + R-loop + R-loop +- +- Fen1 / Cdc9 +- Pif1 - + - + - + - + RNase H1 CD - + - + RNase H1 Stalled fork / + R-loop Stalled fork / kbp RI +- Nt.BbvcI kbp RI

native 8.5 Linear native 8.5 Linear kb - + - + RNase H1 kb 8.5 kb Full-length 8.5 Full-length 8.5 Full-length 6.4 Restart / LagDS 6.4 Full-length, cut 6.4 Restart / LagDS 4.8 Restart / LagDS 4.8 3.7 4.8 3.7 3.7 US 2.3 Stall / LagUS Stall / LagUS 2.3 Stall / Lag 2.3 US Lag , cut 1.4 1.4 1.4 0.7 Left lead / lag 0.7 Left lead / lag

denaturing Left lead / lag

0.7 denaturing denaturing 0.2 0.2 Okazaki 0.2 fragments

1 2 3 4 5 6 7 8 1 2 3 4 1 2 3 4 B - Fen1 / Cdc9 + Fen1 / Cdc9 0.8 kb 1.9 kb 0.4 kb

Cla I Cla I LagUS 0.8 kb 0.8 kb 2.3 kb 2.3 kb Nt.BbvCI

RNase H1 RNase H1 0.8 kb 1.9 kb 0.4 kb 5.5 kb Cla I Cla I LagUS LagDS

0.8 kb 0.8 kb Nt.BbvCI 7.8 kb 7.8 kb

Pif1 Pif1 0.8 kb 1.9 kb 5.9 kb Cla I Cla I

0.8 kb 0.8 kb Nt.BbvCI 7.8 kb 7.8 kb bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission. Figure 6

A B HO HO, ΔG4810-828 - + R-loop - + R-loop +- +- Fen1 / Cdc9 +- +- Fen1 / Cdc9 - + - + - + - + RNaseH1 - + - + - + - + RNaseH1 Stalled fork / Stalled fork / kbp RI kbp RI native 8.5 Linear native 8.5 Linear kb kb 8.5 8.5 6.4 Full-length 6.4 Full-length 4.8 Restart / LagDS 4.8 Restart / LagDS 3.7 3.7 Stall US US Stall / Lag 2.3 Lag 2.3

1.4 1.4

0.7 Left lead / lag 0.7 Left lead / lag denaturing denaturing

0.2 Okazaki 0.2 Okazaki fragments fragments

1 2 3 4 5 6 7 8 1 2 3 4 5 6 7 8 C D HO HO (- R-loop) +- Pif1 - wt EQ Pif1 kb - + - + RNase H1 kb 1.5 US 8.5 Full-length 8.5 Full-length Lag 6.4 DS 6.4 DS 4.8 Restart / Lag 4.8 Lag LagDS 3.7 3.7 Stall / LagUS US 2.3 2.3 Lag 1.0 1.4 1.4 Left lead / lag 0.7 Left lead / lag 0.7

0.5 denaturing denaturing

0.2 normalized signal 0.2 0.0 Pif1- Pif1- 1 2 3 wt E303Q 1 2 3 4 E (i) No R-loop (ii) Bypass / Restart G4810-828 G4810-828 Cla I Cla I LagUS LagDS LagUS LagDS

0.8 kb ~ 2.5 kb ~ 5.3 kb 0.8 kb ~ 2.5 kb ~ 5.1 kb 0.8 kb 7.8 kb 0.8 kb

RNase H1 (iii) Fork stall G4810-828 G4810-828 Cla I LagUS LagDS Cla I 0.8 kb ~ 2.5 kb ~ 5.3 kb 0.8 kb ~ 2.5 kb 0.8 kb ~ 2.5 kb 0.8 kb bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

Figure 7 A B C CD CD HO +- T4 PNK + T4 PNK +- T4 PNK 1.0 - + - + RNase H1 - + DDK - + - + RNase H1 Stalled fork / Stalled fork / RI Stalled fork / RI kbp RI kbp native

native kbp Linear native 8.5 Linear 8.5 8.5 Linear Uncoupled Uncoupled kb kb kb 8.5 Full-length 8.5 Full-length 8.5 Full-length 6.4 6.4 Restart 0.5 6.4 4.8 Restart 4.8 4.8 Restart 3.7 3.7 3.7 2.3 2.3 Stall Stall 2.3 Stall 1.4 1.4 1.4

0.7 Left lead relative restart signal 0.7 Left lead 0.7 Left lead denaturing denaturing 0.0 denaturing 0.2 Okazaki 0.2 Okazaki Okazaki (-) (+) 0.2 fragments fragments fragments DDK 1 2 3 4 1 2 1 2 3 4

(i) Cla I D F CMG Pol α Ctf4 Pol δ PCNA Pol ε 0.8 kb ~ 2.3 kb 5’ 3’ P

Cla I Uncoupling Flap displacement 0.8 kb ~ 2.3 kb 5’ 3’ P

(ii) Cla I

0.8 kb ~ 2.3 kb 5’ 3’ OH Restart Translocation on RNA:DNA duplex Cla I

0.8 kb ~ 2.3 kb 5’ ~ 5.5 kb

E CD

RNase H1 (nM) - 1 0.25 0.06 RNA extension by Pol α Stalled fork / RI kbp native 8.5 Linear Uncoupled kb 8.5 Full-length 6.4 Restart 4.8 3.7 2.3 Stall DNA extension by Pol ε or Pol δ 1.4

0.7 Left lead Okazaki

denaturing fragments 0.2

1 2 3 4 ? bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

Figure S1

A [14] [13] 5’- CGAGGCTTCAACATTATATCTATCCCTCCTTGCTCTAGCGGGCAGCAAGAAGCACAGCACCGCCAGTTACCACGCAGACATCCTGGGGAACTGAGGTAAGCTAGGGTTCCCCGCGTGAAGCGCGGCAATGCGAGGGGAGG 3’- GCTCCGAAGTTGTAATATAGATAGGGAGGAACGAGATCGCCCGTCGTTCTTCGTGTCGTGGCGGTCAATGGTGCGTCTGTAGGACCCCTTGACTCCATTCGATCCCAAGGGGCGCACTTCGCGCCGTTACGCTCCCCTCC [12] [12] [17] ATTCTGCAGATGAGGGTAGGATTCCGTTGCAAGGGGAGGATTCCACGCGTTAGAGGATTCCGCAAAGGAAGGGTTCCACAGGAGGGAAGGGCTCTACGCGAGGTGAGGGTTCCACTGATCCGGCAGCTCGAGGGCTCCGA TAAGACGTCTACTCCCATCCTAAGGCAACGTTCCCCTCCTAAGGTGCGCAATCTCCTAAGGCGTTTCCTTCCCAAGGTGTCCTCCCTTCCCGAGATGCGCTCCACTCCCAAGGTGACTAGGCCGTCGAGCTCCCGAGGCT

CCAAGAGTTCCAGGCCGCTCAAAGGTTCCCCTACGCGAGATTCAGCGCTAGGGTGAAGATTTCTGGGCTACAAGAAAACTCAGCACTAGGTGCCCCACTGCTCACCAGTGTTCTGAACTACACGAGGGCCGATAGGGCCG GGTTCTCAAGGTCCGGCGAGTTTCCAAGGGGATGCGCTCTAAGTCGCGATCCCACTTCTAAAGACCCGATGTTCTTTTGAGTCGTGATCCACGGGGTGACGAGTGGTCACAAGACTTGATGTGCTCCCGGCTATCCCGGC [16] [32] [41] GTAGAAAGCCGAAAGCCGCCGCTGCCGCCATGCCGCGTCAGCAAGAGGAAAAGGGAGGGGGGCGTCCAGCGCGGGCCGACTGCTGCACTGGGGGGGGGGGGGGTTTACGGGCGATTTAGAGCACGAGGGTGCCACGC CATCTTTCGGCTTTCGGCGGCGACGGCGGTACGGCGCAGTCGTTCTCCTTTTCCCTCCCCCCGCAGGTCGCGCCCGGCTGACGACGTGACCCCCCCCCCCCCCAAATGCCCGCTAAATCTCGTGCTCCCACGGTGCG [12] [19] [21] TGCGCAAGGGGAGCAGGGGTTCGCGCTAGGGCACCACGCTGCTGGAGAGTTGGGGGGGGGGGTGGAGATCGGAGGATTTCTACACGACCTGATCCGCGGTTTGCGGGGCAGGGGGAAGGTTCGGTGGGTTCCGCGGT ACGCGTTCCCCTCGTCCCCAAGCGCGATCCCGTGGTGCGACGACCTCTCAACCCCCCCCCCCACCTCTAGCCTCCTAAAGATGTGCTGGACTAGGCGCCAAACGCCCCGTCCCCCTTCCAAGCCACCCAAGGCGCCA [20] [9] GGAGCTCTGGAGATCGGAGGATTTCTACACGACCTGATCCGCGGTTTGCGGGGAAGGGGGAAGGTTCGGTGGGTTCCGCGGAACTGCGCGAGCCAGGGTGCTGGCGTGTGCGCTGCCCCCCCCCCCCCCCCCCCGTTC CCTCGAGACCTCTAGCCTCCTAAAGATGTGCTGGACTAGGCGCCAAACGCCCCTTCCCCCTTCCAAGCCACCCAAGGCGCCTTGACGCGCTCGGTCCCACGACCGCACACGCGACGGGGGGGGGGGGGGGGGGGCAAG [7] [21] 810 [63] 828 AGAGGATTTTAGCACAACTCCAATTGTGCTGCGATTCTGGTTGTGCCGAGTTGCGAGAGAGGCTAAGGGTGAAAAGCTGCACAAGGAGGGGATTCGGAGGGTTTAGAGGGTTCCGCGGCACTGCGCCGCAGTGCTGCCCG TCTCCTAAAATCGTGTTGAGGTTAACACGACGCTAAGACCAACACGGCTCAACGCTCTCTCCGATTCCCACTTTTCGACGTGTTCCTCCCCTAAGCCTCCCAAATCTCCCAAGGCGCCGTGACGCGGCGTCACGACGGGC [34] [18] AGGGTTCGGAGCAATTCCGGTTGTGCCGTGATCCTTGGTTGTGCTGAGTTGCGGTGAGGGAAAGGGAAGGGAAAGCTCAGAGGGTTCCGAGCTATCCTGAGGGTGCGAAGCTGCACAAGGGCAGGGTTCCGAGGGTTCC TCCCAAGCCTCGTTAAGGCCAACACGGCACTAGGAACCAACACGACTCAACGCCACTCCCTTTCCCTTCCCTTTCGAGTCTCCCAAGGCTCGATAGGACTCCCACGCTTCGACGTGTTCCCGTCCCAAGGCTCCCAAGG [8] [3] [20] GCGGCACTGCGTGAGCCCCGGTGCCGCGCTGCCCAAAGGTTCGGAGGGTTTTAATGCGATTCCGGTTGGGCTGTGATTCTGGTTATGCCAAGTTGCGCGAGGGGAGGGGAAGAGACGGCTCGGAGGATTCCGAGGTATC CGCCGTGACGCACTCGGGGCCACGGCGCGACGGGTTTCCAAGCCTCCCAAAATTACGCTAAGGCCAACCCGACACTAAGACCAATACGGTTCAACGCGCTCCCCTCCCCTTCTCTGCCGAGCCTCCTAAGGCTCCATAG [15][14] CTGAGGGTGCAAACTGCACAAGGGGAGGATTCGAAGGGTTCTGTGATCAGGGCCAACGCTCAAAAGTGCCATGTTACAGGAGAGATGAACTCAAAAGTGCCACGTTGCAAGAGAGGCAAGCTCC - 3’ GACTCCCACGTTTGACGTGTTCCCCTCCTAAGCTTCCCAAGACACTAGTCCCGGTTGCGAGTTTTCACGGTACAATGTCCTCTCTACTTGAGTTTTCACGGTGCAACGTTCTCTCCGTTCGAGG - 5’

B - + RNase A C CD HO 123456789 123456789 Fraction Number

kbp

kbp EtBr 8.6 Plasmid + R-loop Plasmid + R-loop 10 2.3

3 8.6 Plasmid + R-loop 2.3 0.7 1 Free

EtBr Autoradiograph RNA

D nm 2.0 1 µm 1 µm 1 µm 1.5

1.0

0.5

0.0 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

Figure S2

A CD HO - + ClaI - + ClaI +- +- R-loop +- +- R-loop kb - + - + - + - + DDK kb - + - + - + - + DDK 8.5 Full-length 8.5 Full-length 6.4 Restart 6.4 4.8 4.8 Restart 3.7 3.7 2.3 Stall Stall 2.3 Left lead 1.4 1.4 Left lead Left lead, cut 0.7 Left lead, cut 0.7 denaturing denaturing

0.2 Okazaki Okazaki fragments 0.2 fragments

1 2 3 4 5 6 7 8 1 2 3 4 5 6 7 8

B CD HO - + R-loop - + R-loop 30 120 30 120 30 120 30 120 15 60 15 60 minutes 15 60 15 60 minutes

Stalled fork / Stalled fork / kbp RI kbp RI native 8.5 Linear native 8.5 Linear kb Uncoupled kb Uncoupled 8.5 Full-length 8.5 Full-length 6.4 6.4 4.8 Restart 4.8 Restart 3.7 3.7 2.3 Stall Stall 2.3 1.4 1.4 0.7 Left lead Left lead 0.7 denaturing denaturing Okazaki 0.2 Okazaki 0.2 fragments fragments

1 2 3 4 5 6 7 8 1 2 3 4 5 6 7 8 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

Figure S2

C - R-loop 100 75 CD, dNTP pulse-chase Linear 50

- + R-loop RI/Stalled fork native gel products

% total 25 3 12 48 3 12 48 minutes .25 6 24 .25 6 24 0 0 25 50 Time (min)

kbp Stalled fork / RI + R-loop native 8.5 Linear 100 Linear Uncoupled RI/Stalled fork kb 75 8.5 6.4 Full-length 50 4.8 3.7 % total 25

2.3 0 Stall 0 25 50 Time (min) 1.4 Left lead Full-length (-) R-loop

0.7 100 gel products denaturing Full-length (+) R-loop denaturing Stall (+) R-loop Okazaki 75 fragments 0.2 50 25 % right lead 0 1 2 3 4 5 6 7 8 9 10 11 12 0 25 50 Time (min)

- R-loop 100 HO, dNTP pulse-chase 75 - + R-loop Linear 50 RI/Stalled fork native gel products 3 12 48 3 12 48 minutes % total .25 6 24 .25 6 24 25 0 0 25 50 Stalled fork / Time (min) kbp RI

native 8.5 Linear + R-loop Uncoupled 100 Linear RI/Stalled fork kb 75 8.5 Full-length 6.4 50 4.8 % total 3.7 25 Stall 2.3 0 0 25 50 1.4 Time (min) Left lead 0.7 100 Full-length (-) R-loop gel products denaturing Full-length (+) R-loop denaturing 75 Okazaki Stalled (+) R-loop 0.2 fragments 50 25 % right lead 0 1 2 3 4 5 6 7 8 9 10 11 12 0 25 50 Time (min) bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

Figure S2

D Cla I BbvCI ARS305 mAirn 803 925 2728 4102 4795 8624

normal leading strand replication restart Cla I Cla I

0.8 kb 0.8 kb Nt.BbvCI Nt.BbvCI Nb.BbvCI Nb.BbvCI 7.8 kb 2.3 - 2.8 kb 5.0 - 5.5 kb

1.2 - 1.7 kb 0.8 kb 4.0 kb 3.8 kb 0.8 kb 2.3 - 2.8 kb 3.8 kb

CD HO - + - + R-loop - Nt Nb - Nt Nb - Nt Nb - Nt Nb BbvCI kb 8.5 Full-length 6.4 Restart 4.8 FL / Restart, cut 3.7 2.3 Stall

1.4 Left lead 0.7

denaturing Okazaki fragments 0.2

1 2 3 4 5 6 7 8 9 10 11 12

E CD HO +- +- R-loop - + - + - + - + CTM

Stalled fork / kbp RI

native 8.5 linear 6.4 Uncoupled kb 8.5 Full-length 6.4 4.8 Restart 3.7

2.3 Stall

1.4

denaturing 0.7 Left lead

Okazaki 0.2 fragments

1 2 3 4 5 6 7 8 bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

Figure S3

CD ARS305

(-) RNase H1 (+) RNase H1

Stalling

0.8 kb 2.3 kb Complete

0.8 kb or 7.8 kb Uncoupling

0.8 kb 2.3 kb

B CD ARS305

(-) RNase H1 (+) RNase H1

Restart Restart

0.8 kb 0.8 kb 2.3 kb 5.5 kb 2.3 kb 5.5 kb bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

Figure S3

C HO ARS305

G4 (-) RNase H1 (+) RNase H1

Stalling Stalling

0.8 kb 0.8 kb 2.8 kb 2.8 kb

D HO ARS305

G4 (-) RNase H1 (+) RNase H1

Restart Complete

0.8 kb 0.8 kb 2.8 kb 5.0 kb 7.8 kb bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

Figure S4

A HO - R-loop MscI BseRI - + - + - + Pif11 kb 8.5 Full-length 6.4 LagDS 4.8 FL+ LagDS,cut 3.7 Stall / LagUS 2.3

1.4 FL+Stall / LagUS,cut Left lead / lag 0.7 denaturing

0.2

1 2 3 4 5 6

B MscI BseRI

Cla I HO ARS305 mAirn

803 9251881 2728 4770 8624 4102

replicate

0.8 kb 7.8 kb

0.8 kb 1.1 kb 6.7 kb MscI 0.8 kb BseRI 4.0 kb 3.8 kb

C MscI BseRI

Cla I HO ARS305 mAirn

803 9251881 2728 4770 8624 4102

replicate G4810-828

0.8 kb 2.5 kb 5.3 kb

0.8 kb 7.8 kb

0.8 kb 1.1 kb 1.4 kb 5.3 kb MscI 0.8 kb BseRI 2.5 kb 1.5 kb 3.8 kb bioRxiv preprint doi: https://doi.org/10.1101/2021.07.16.452753; this version posted July 17, 2021. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.

Figure S5

A B CD 0.061 RNase H1 (nM) CD - + - + Pol δ - + -- + Pol ε Stalled fork / - + - + - Pol δ kbp RI native - + + -- Pol α 8.5 Linear kb Uncoupled 8.5 End labeled kb 6.4 8.5 4.8 6.4 Full-length 3.7 4.8 Restart 3.7 2.3 2.3 Stall 1.4 1.4 0.7 0.7 Left lead denaturing 0.2 denaturing Okazaki 0.2 fragments

1 2 3 4 5 1 2 3 4

C CD HO +- +- R-loop RNase H2 - + - + - + - + RNase H2

Stalled fork / kbp RI native 8.5 Linear Uncoupled kb 8.5 6.4 Full-length 4.8 Restart 3.7 RNH202 Stall RNH201 2.3 1.4 Left lead 0.7

denaturing Okazaki 0.2 fragments RNH203 1 2 3 4 5 6 7 8