Distinct mechanisms of replication stress induced by oncogenic RAS and cyclin E1 converge on R-loop-dependent fork reversal

Domain: biology
Published: 2026-04-03
Editorial outcome: major_revision

Abstract
Activated oncogenes elicit genomic instability by inducing DNA replication stress. Here we show that replication fork reversal and chromosome mis-segregation induced by oncogenic RAS (HRASV12) or cyclin E1 overexpression are largely caused by co-transcriptional RNA:DNA hybrids (R-loops) formed during S-phase. Furthermore, we demonstrate that replication stress induced by HRASV12, but not cyclin E1, is driven by reactive oxygen species (ROS) in a manner dependent on the replisome-associated ROS sensor peroxiredoxin 2 (PRDX2) and is linked to PRDX2-mediated release of the fork acceleration factor TIMELESS from the replisome. Inhibition of fork reversal in cells overexpressing HRASV12 or cyclin E1 induces unrestrained DNA synthesis mediated by the MUS81 endonuclease and the primase-polymerase PRIMPOL, thereby promoting proper chromosome segregation in mitosis. These results establish PRIMPOL repriming as part of the MUS81-dependent replication restart mechanism that operates at sites of R-loop-mediated transcription-replication conflicts to maintain genomic stability. Furthermore, our data indicate that, despite their protective role during S-phase, persistent reversed forks impair chromosome segregation in mitosis, potentially leading to DNA breaks and chromosomal rearrangements.

Introduction
Cancer is caused by genetic alterations that activate oncogenes and inactivate tumor suppressor genes. Oncogenes are known to induce stalling and collapse of DNA replication forks in precancerous lesions, leading to persistent activation of the DNA damage response machinery 1–4. This phenomenon, known as oncogene-induced replication stress creates a selective pressure for the outgrowth of malignant clones carrying mutations in the tumor suppressor proteins such as p53, which inactivate the DNA damage checkpoints, allowing the cancer to develop 5. One of the prototypical replication stress-inducing oncogenes is oncogenic RAS 3,6. The RAS family consists of three proto-oncogenes - KRAS, NRAS and HRAS - which function as membrane-associated GTPase signal transducers that, upon GTP binding, activate multiple mitogenic pathways including RAF/MEK/ERK and PI3K/AKT 7,8. RAS is turned into an oncogene by a mutation in the conserved codons 12, 13, or 61, which impairs intrinsic GTP hydrolysis or binding to GTPase-activating proteins, making RAS constitutively active. Such KRAS mutations are most commonly found in colorectal cancer (33%), pancreatic cancer (57%), and non-small-cell lung cancer, while HRAS mutations are associated with skin and head and neck cancers, and NRAS mutations are common in hematopoietic malignancies 7,8. Evidence suggests that replication stress induced by oncogenic RAS arises from increased global transcription driven by elevated expression of the TATA-box binding protein (TBP), leading to the accumulation of R-loops 9. These three-stranded nucleic acid structures, composed of an RNA:DNA hybrid and a single-stranded DNA (ssDNA) loop, can form co-transcriptionally by invasion of the nascent transcript into the underwound DNA duplex behind the transcription complex, and act as a strong barrier to replication fork progression 10. However, a recent study suggested that, rather than inducing R-loop formation, the transcriptional burst triggered by oncogenic RAS leads to H3K27me3- and HP1-associated chromatin compaction, thereby resulting in replication fork stalling and cell death 11. Moreover, it should be noted that sustained mitogenic stimulation by oncogenic RAS increases the intracellular concentration of reactive oxygen species (ROS) 12,13, which are known to induce R-loop-mediated fork stalling by promoting R-loop formation at sites of transcription-replication conflicts (TRCs) in a manner dependent on the replisome-associated ROS sensor peroxiredoxin 2 (PRDX2) 14. Thus, the molecular mechanism underlying DNA replication stress induced by this oncogene remains elusive.

Another prototypical oncogene that induces DNA replication stress is cyclin E1, the product of the CCNE1 gene 1,2. Cyclin E1, in complex with CDK2, normally drives the progression of the cell cycle from the G1 phase to the S phase by phosphorylating multiple proteins, including RB, leading to the expression of various genes required for DNA replication 15,16. However, cyclin E1 is frequently overexpressed in premalignant lesions and cancers such as ovarian tumors or uterine carcinomas due to gene amplification 15,17, which induces DNA replication stress 2. Accumulating evidence suggests that cyclin E1 overexpression induces DNA replication stress by enhancing replication origin firing 18,19. Cyclin E1 overexpression dramatically shortens the length of G1 phase, resulting in firing of intragenic origins that are normally inactivated by transcription in G1 19. The firing of these oncogene-induced origins gives rise to transcription-dependent chromosome breakage at sites that coincide with chromosomal rearrangement breakpoints in a large cohort of cancers 19. Thus, cyclin E1-induced replication stress is caused by aberrant firing of intragenic origins and subsequent head-on TRCs, which can lead to fork collapse and genomic instability. Although head-on TRCs are known to promote R-loop formation 20–23, particularly when replication fork progression is slowed due to impairment of the replication machinery 14,24, it remains to be determined whether R-loops are a major cause of DNA replication stress induced by cyclin E1 overexpression.

R-loops arrest replication fork progression by inducing replication fork reversal 14,25. This fork remodeling process, mediated by the RAD51 recombinase and the DNA translocases ZRANB3, HLTF, and SMARCAL1, results in the pairing of nascent DNA strands to form a four-way DNA structure, where the newly formed DNA arm is covered by BRCA2-stabilized RAD51 filament to prevent nucleolytic resection 25. Although overexpression of oncogenes such as cyclin E1 or CDC25A has been shown to induce fork reversal 26, the role of R-loops in these processes has not yet been investigated.

R-loop-stalled forks can be reactivated in a multistep process triggered by the action of two DNA helicases, namely RECQ1 and RECQ5 27. RECQ1 helicase mediates reverse branch migration to eliminate reversed forks while RECQ5 helicase removes RAD51 from the stalled fork to prevent further fork reversal and to promote fork cleavage by the MUS81-EME1 endonuclease 27–29. This initiates the replication restart process that additionally requires the strand-annealing factor RAD52, the DNA ligase IV (LIG4)/XRCC4 complex, the RNA polymerase II (RNAPII) elongation factor ELL, and the non-catalytic subunit of DNA polymerase delta POLD3 27. Defects in this DNA repair pathway lead to genomic instability due to impaired chromosome segregation in mitosis caused by the presence of regions of incompletely replicated DNA 27. However, its importance for DNA replication in oncogene-expressing cells remains to be determined.

Here, we show that the overexpression of HRASV12 or cyclin E1 induces R-loop formation specifically in S-phase of the cell cycle and that R-loops are the primary cause of replication fork reversal and chromosome mis-segregation induced by these oncogenes. Furthermore, we demonstrate that R-loop-mediated replication stress induced by HRASV12, but not by cyclin E1, is largely driven by ROS in a manner dependent on PRDX2 and is linked to PRDX2-mediated release of the fork acceleration factor TIMELESS from the replication fork. Suppression of fork reversal in cells overexpressing HRASV12 or cyclin E1 restores normal replication fork progression in a manner dependent on the primase-polymerase PRIMPOL and proteins that mediate the reactivation of R-loop-stalled forks. Our data suggest that persistent reversed forks can give rise to genomic instability if not restarted before the onset of mitosis. In addition, these findings identify the MUS81-PRIMPOL axis as a potential therapeutic target for cancers expressing oncogenes that induce transcription-dependent replication stress.

Results
HRASV12- and cyclin E1-induced replication stress is mainly caused by R-loops
To study the molecular mechanisms underlying oncogene-induced replication stress, we used pBABEneo-based retroviral vectors for overexpression of HRASV12 and cyclin E1, respectively, in human cells (Supplementary Fig. 1a). Target cells infected with the respective retroviruses were subjected to phenotypic analyses at various times after selection in G-418-containing medium (Day 0), thereby eliminating cells in which a DNA copy of the retroviral genome was not incorporated into a chromosome of the host cell. Using this retroviral transduction method, we were able to overproduce both HRASV12 and cyclin E1 in human osteosarcoma U2OS cells, as revealed by western blot analysis of total cell extracts (Supplementary Fig. 1b). To assess replication fork dynamics in these cells, we used the well-established DNA fiber assay based on pulse labeling of cells with the halogenated thymidine analogues 5-chloro-2’-deoxyuridine (CldU) and 5-iodo-2’-deoxyuridine (IdU), followed by visualization of labeled replication tracts on DNA fiber spreads by indirect immunofluorescence. As expected, by measuring the length of the labeled nascent DNA tracts, we found that the rate of replication fork progression in cells overexpressing either of these oncogenes was significantly lower than in cells harboring the empty vector at both 2 and 4 days after selection (Supplementary Fig. 1c). We also observed that in cells overexpressing HRASV12 or cyclin E1, labeled nascent DNA tracts of sister replication forks showed a marked asymmetry in their length (Supplementary Fig. 1d), a phenotype known as sister fork asymmetry, which indicates the occurrence of replication fork stalling events 30. Consistent with the replication fork stalling phenotype, cells overexpressing HRASV12 or cyclin E1 also exhibited an increased frequency of anaphase bridges and micronuclei (Supplementary Fig. 1e, f), which results from segregation of incompletely replicated DNA 31. Furthermore, using high content microscopy followed by unbiased quantitative image-based cytometry (QIBC), we found that overexpression of either oncogene induced the formation of γH2AX foci in S-phase cells identified by PCNA staining (Supplementary Fig. 2a-c). Notably, only a mild increase in 53BP1 foci - marker of DNA double-strand breaks (DSBs) - was detected in S/G2 cells upon overexpression of HRASV12 or cyclin E1 (Supplementary Fig. 2d, e), suggesting that replication stalling induced by these oncogenes does not lead to extensive DNA breakage during S-phase. Additionally, a modest increase in 53BP1 foci was detected in G1 cells upon overexpression of either oncogene (Supplementary Fig. 2d, e), potentially reflecting unresolved under-replicated DNA from the previous cell cycle that led to DNA breakage during chromosome segregation 31.

To elucidate the role of R-loops in DNA replication stress induced by overexpression of HRASV12 or cyclin E1, we introduced the retroviruses encoding these oncogenes into U2OS T-REx cells harboring an RNASEH1 transgene under the control of a doxycycline-regulated CMV promoter (U2OS T-REx [RNH1-GFP]) 32. RNase H1 is an endonuclease that specifically cleaves the RNA strand in RNA:DNA hybrids, thereby eliminating R-loops and preventing R-loop-mediated replication stress 27,30,33. By DNA fiber assay, we found that RNase H1 overexpression almost completely prevented replication fork slowing and sister fork asymmetry induced by HRASV12 or cyclin E1 overexpression in U2OS T-REx [RNH1-GFP] cells both 2 and 4 days after selection (Fig. 1a-c; Supplementary Fig. 3a, b). The same effects were observed when U2OS cells transduced with HRASV12 or cyclin E1 retroviruses were treated with the RNAPII transcription initiation inhibitor triptolide (TRP) for one hour prior to and during pulse labeling with CldU and IdU (Fig. 1d, e; Supplementary Fig. 3c, d). In addition, by analysis of DNA replication intermediates using electron microscopy, we found that RNase H1 overexpression completely suppressed replication fork reversal induced by HRASV12 or cyclin E1 overexpression in U2OS T-REx [RNH1-GFP] (Fig. 1f). Taken together, these results suggest that replication fork stalling induced by HRASV12 or cyclin E1 overexpression is mainly caused by co-transcriptional R-loops. Consistently, anaphase bridges and micronucleation induced by HRASV12 or cyclin E1 overexpression were suppressed by RNase H1 overexpression in U2OS T-REx [RNH1-GFP] cells (Fig. 1g, h; Supplementary Fig. 3e).

Cells overexpressing HRASV12 or cyclin E1 accumulate R-loops during S and G2 phases of the cell cycle
To investigate the molecular mechanisms underlying R-loop formation induced by overexpression of HRASV12 or cyclin E1, we used a U2OS T-REx cell line conditionally expressing catalytically inactive RNase H1 fused to green fluorescent protein [RNH1(D210N)-GFP] 32. This RNase H1 mutant, which binds but does not cleave RNA:DNA hybrids, accumulates at R-loops formed in cells and thus serves as an in vivo R-loop reporter 27,32–34. The oncogenes were introduced into these cells by retroviral transduction, and RNH1(D210N)-GFP expression was induced by the addition of doxycycline for the last 24 hours of post-selection growth (Fig. 2a). Prior to fixation, cells were pre-extracted with a Triton X-100-containing buffer to remove unbound RNH1(D210N)-GFP protein from nuclei. Fixed cells were subjected to immunofluorescence staining for PCNA to determine the DNA replication status and DAPI staining (DNA content) to determine the cell cycle stage of the individual cells (Fig. 2a). Using QIBC, we found that overexpression of either oncogene induced the formation of nuclear foci of RNH1(D210N)-GFP in S-phase cells (PCNA-positive), but not in G1 cells (PCNA-negative/low DAPI) (Fig. 2a-c), suggesting S-phase-specific R-loop formation, likely as a result of head-on TRCs. Consistently, proximity ligation assay (PLA) followed by QIBC revealed increased colocalization between PCNA and the elongating form of RNAPII in these cells (Supplementary Fig. 4a-c), indicating the occurrence of head-on TRCs 22. However, we cannot exclude the possibility that some of these PLA foci represent co-directional RNAPII collisions behind the replication fork 35, potentially arising from oncogene-induced fork slowing. Interestingly, we also observed a significant increase in the number of RNH1(D210N)-GFP foci in G2 cells (PCNA-negative/high DAPI) upon overexpression of either oncogene (Fig. 2c). These foci likely reflect R-loops persisting from S-phase because neither oncogene induced TRCs in G2 cells, as revealed by QIBC analysis of PCNA/RNAPII PLA foci (Supplementary Fig. 4c).

To confirm that the overexpression of HRASV12 and cyclin E1 via retroviral transduction induced R-loop formation, we isolated genomic DNA from U2OS cells transduced with the respective retroviral vector and performed slot blot analysis using the S9.6 antibody, which recognizes RNA:DNA hybrids. As expected, we observed a significantly higher R-loop signal in samples from oncogene-expressing cells compared to that of cells harboring the empty vector (Supplementary Fig. 5). Treating the genomic DNA samples with RNase H before slot blot analysis diminished the S9.6 signals, confirming the specificity of the R-loop signals detected with the S9.6 antibody (Supplementary Fig. 5).

It has been proposed that R-loops induced by oncogenic RAS result from increased transcriptional activity due to upregulation of the transcription factor TBP 9. Consistently, we found by 5-ethynyl uridine (5-EU) pulse labeling that overexpression of HRASV12, but not cyclin E1, increased the transcriptional activity in U2OS cells (Supplementary Fig. 6a). However, QIBC analysis showed that HRASV12 overexpression increased transcription not only in S-phase cells (PCNA-positive) but also in G1 cells (PCNA-negative/low DAPI) (Supplementary Fig. 6b-d), suggesting that the observed S-phase-specific accumulation of R-loops in HRASV12-expressing cells is not solely caused by increased transcription in these cells, but may be specifically associated with TRCs.

Suppression of replication fork reversal induces unrestrained DNA synthesis and prevents chromosome mis-segregation in cells overexpressing HRASV12 or cyclin E1
Replication fork stalling at co-transcriptional R-loops can be rescued by PARP inhibition or ZRANB3 depletion 14,27, which suppress replication fork reversal, leading to unrestrained DNA synthesis 14,27,28,36. Since we found that fork reversal induced by HRASV12 or cyclin E1 overexpression is caused by R-loops, we sought to test whether PARP inhibition with olaparib or siRNA-mediated depletion of ZRANB3 induces unrestrained replication fork progression in U2OS cells transduced with HRASV12 or cyclin E1 retroviral vectors. By DNA fiber assay, we found that the replication fork slowing and sister fork asymmetry phenotypes induced by these oncogenes were largely suppressed when cells were pretreated with olaparib or depleted of ZRANB3 (Fig. 3a-d; Supplementary Fig. 7a, b). Overexpression of HRASV12 or cyclin E1 mediated by the retroviral vectors also did not induce replication fork slowing in ZRANB3 knockout U2OS cells (Fig. 3e). Moreover, the fork slowing and sister fork asymmetry phenotypes of U2OS cells overexpressing HRASV12 or cyclin E1 were rescued by depletion of the SMARCAL1 translocase (Supplementary Fig. 7c, d), which acts in conjunction with ZRANB3 to promote fork reversal 25. To confirm these findings, we depleted ZRANB3 in hTert-immortalized human BJ fibroblasts expressing tamoxifen-inducible HRASV12 (BJ-hTert HRASV12ER-TAM) and in hTert-immortalized human retinal pigment epithelial-1 (RPE1) cells engineered to overexpress cyclin E1 from a doxycycline-regulated promoter 9,37. As expected, we observed that overexpression of HRASV12 and cyclin E1 dramatically slowed replication fork progression in the respective cells transfected with control siRNA (Supplementary Fig. 7e, f). On the contrary, cells depleted of ZRANB3 exhibited unrestrained replication fork progression upon oncogene overexpression as observed in U2OS cells (Supplementary Fig. 7e, f), excluding a cell line specific phenomenon. In addition, we found that PARP inhibition rescued fork slowing induced by HRASV12 or cyclin E1 overexpression in primary BJ fibroblasts (Supplementary Fig. 7g, h), excluding the possibility that this phenomenon is only seen in immortalized cells. These results prove that replication fork progression in cells overexpressing HRASV12 or cyclin E1 is restrained by fork reversal.

Next, we investigated whether persistent reversed forks are responsible for chromosome mis-segregation in cells overexpressing HRASV12 or cyclin E1. Notably, we found that lack of ZRANB3 completely prevented HRASV12- and cyclin E1-induced formation of anaphase bridges and micronuclei in U2OS cells (Fig. 3f, g; Supplementary Fig. 7i). The micronucleation phenotype of these cells was also rescued by SMARCAL1 depletion (Supplementary Fig. 7j). Moreover, induction of HRASV12 or cyclin E1 expression did not increase micronucleation in ZRANB3-depleted BJ fibroblasts and RPE1 cells, respectively (Supplementary Fig. 7k, l). These findings suggest that the defective chromosome segregation in cells overexpressing HRASV12 or cyclin E1 is caused by persistent reversed forks. Consistent with this hypothesis, we found that suppression of fork reversal by ZRANB3 depletion prevented micronucleation induced in U2OS cells by low doses of hydroxyurea (HU) or aphidicolin (Supplementary Fig. 8), both of which also induce R-loop-mediated TRCs 14,24.

Unrestrained replication fork progression in HRASV12- or cyclin E1-overexpressing cells depends on proteins involved in restarting R-loop-stalled forks
The unrestrained replication fork progression induced by PARP inhibition in cells treated with R-loop-inducing drugs is dependent on the MUS81-EME1 endonuclease, the DNA helicase RECQ1, the DNA ligase IV (LIG4), and the transcription elongation factor ELL, among others 14,27. We therefore tested whether these factors are also required for the unrestrained DNA synthesis observed in oncogene-expressing cells upon PARP inhibition. For this, we used siRNA technology to deplete RECQ1, MUS81, LIG4 or ELL in U2OS cells transduced with the HRASV12 or cyclin E1 retroviral vectors. By DNA fiber assay, we found that depletion of any of these proteins abolished the rescue of replication fork slowing in HRASV12- or cyclin E1-expressing cells by PARP inhibition with olaparib (Fig. 4a). In addition, we found that depletion of MUS81 attenuated olaparib-stimulated DNA synthesis in HRASV12-overexpressing immortalized or primary BJ fibroblasts as well as in cyclin E1-overexpressing RPE1 cells and primary BJ fibroblasts (Supplementary Fig. 9a-d). RECQ1 promotes the restart of R-loop-stalled forks by converting reversed forks to a three-way junction, which allows for fork cleavage by MUS81 endonuclease, initiating the replication restart process 27. Therefore, RECQ1 is not required for replication restart if fork reversal is prevented by ZRANB3 or HLTF depletion 14,27. Consistently, we found that unrestrained DNA synthesis induced in HRASV12- or cyclin E1-overexpressing U2OS cells by ZRANB3 depletion was dependent on the presence of MUS81 but not RECQ1 (Fig. 4b). Together, these results suggest that R-loop-mediated fork stalling in oncogene-expressing cells is counteracted by replication restart via the MUS81-LIG4-ELL axis.

Unrestrained replication fork progression in oncogene-expressing cells defective in fork reversal is mediated by PRIMPOL
Recent work has shown that the restart of R-loop-stalled forks via the MUS81-LIG4-ELL axis also requires the DNA primase-polymerase PRIMPOL 14, which is known to mediate repriming of DNA synthesis downstream of DNA lesions that block replication fork progression 38–40. We therefore investigated whether PRIMPOL also plays a role in the observed unrestrained replication fork progression in oncogene-expressing cells defective in fork reversal. For this, wild-type and ZRANB3 knockout U2OS cells transduced with HRASV12 or cyclin E1 retroviral vectors were transfected with control or PRIMPOL siRNA and replication fork progression was monitored by DNA fiber assay 72 hours post-transfection. Cells were also depleted of MUS81 as positive control. We found that, as with the proteins of the MUS81-LIG4-ELL axis, PRIMPOL depletion impaired unrestrained replication fork progression in ZRANB3 knockout cells overexpressing HRASV12 or cyclin E1 (Fig. 5a). PRIMPOL-mediated repriming causes discontinuous DNA replication, leaving a ssDNA gap behind the fork 39,40. To confirm that this replication restart mechanism is involved in the unrestrained replication fork progression observed in HRASV12- and cyclin E-overexpressing cells upon inactivation of fork reversal, we carried out a modified DNA fiber assay where cells were permeabilized after CldU/IdU labeling and then treated with S1 nuclease, which specifically cleaves single-stranded nucleic acids including single-stranded regions of duplex DNA 41,42. We found that DNA fibers in ZRANB3 knockout cells expressing HRASV12 or cyclin E1, which showed unrestrained fork progression, were sensitive to S1 nuclease digestion compared to cells carrying the empty vector (Fig. 5b). Taken together, these results indicate that unrestrained fork progression induced by fork reversal deficiency in HRASV12/cyclin E1-overexpressing cells is dependent on PRIMPOL and further support a role for PRIMPOL in restarting R-loop-stalled forks via the MUS81-LIG4-ELL axis. In agreement with this conclusion, we also found that PRIMPOL depletion did not further exacerbate replication fork stalling in MUS81 knockout HeLa cells (Supplementary Fig. 10).

HRASV12-induced replication stress is caused by reactive oxygen species generated by NADPH oxidases
Our finding that R-loop formation in HRASV12-overexpressing cells is caused by TRCs rather than increased transcription (Fig. 2; Supplementary Fig. 6) prompted us to further investigate the molecular mechanism underlying HRASV12-induced replication stress. It is well established that RAS oncogenes elevate intracellular ROS levels by upregulating the NADPH oxidases NOX1 and NOX4 43–46. In light of our recent findings that excessive ROS levels drive R-loop-mediated replication stress by promoting R-loop formation following head-on TRCs 14, we sought to determine whether HRASV12 induces replication fork stalling through a ROS-dependent mechanism. For this, we also generated stable U2OS cell lines expressing either HRASV12 or cyclin E1 under the control of a doxycycline-inducible TRE3G promoter (Fig. 6a, b). In this system, we observed a significant reduction in replication fork speed for both oncogenes two days after the addition of doxycycline (Fig. 6a, b). To eliminate ROS, we treated cells with the ROS scavenger N-acetyl-L-cysteine (NAC), which was present 2 hours prior to and during nascent DNA labeling for DNA fiber assay. Alternatively, cells were treated for 24 hours with setanaxib, a dual inhibitor of NOX1 and NOX4 46,47. We found that both NAC and setanaxib rescued the fork slowing phenotype of HRASV12-overexpressing cells but not cyclin E1-overexpressing cells (Fig. 6a, b). Similar results were obtained with U2OS cells overexpressing HRASV12 or cyclin E1 following transduction of the respective retroviral vectors (Supplementary Fig. 11a). Moreover, NAC and setanaxib rescued HRASV12-induced fork slowing in primary BJ fibroblasts (Supplementary Fig. 11b). In contrast, cyclin E1-induced fork slowing in primary BJ fibroblasts was not affected by NAC or setanaxib, although it was still rescued by the inhibition of transcription with triptolide (Supplementary Fig. 11b, c). In addition, we found that HRASV12-, but not cyclin E1-induced replication fork slowing was alleviated upon induction of antioxidant response with tert-butylhydroquinone (tBHQ) or sulforaphane (SFN) (Supplementary Fig. 11d, e), which activate NRF2 signaling 49,50. Together, these data suggest that HRASV12-induced replication stress is caused by ROS generated by NADPH oxidase enzymes. In support of this notion, we found that micronucleation induced by HRASV12 was suppressed when cells were treated with setanaxib, tBHQ or SFN, whereas cyclin E1-induced micronucleation was not affected by these treatments (Fig. 6c, d; Supplementary Fig. 11f, g).

HRASV12-induced replication stress depends on the replisome-associated ROS sensor PRDX2 and is linked to PRDX2-dependent dissociation of TIMELESS from the replisome
R-loop-mediated replication fork stalling caused by excessive ROS levels depends on the replisome-associated ROS sensor PRDX2 14. ROS disrupt the oligomeric state of PRDX2 and thereby enforce dissociation of the TIMELESS-TIPIN complex from the replisome, leading to a reduction in replication fork velocity 51. This ultimately increases the incidence of R-loop-mediated fork stalling events following head-on TRCs 14. To further explore the role of ROS in HRASV12-induced replication stress, we tested whether this phenomenon is suppressed by PRDX2 depletion. By DNA fiber assay, we found that siRNA-mediated depletion of PRDX2 almost completely rescued HRASV12-induced fork slowing in U2OS cells or primary BJ fibroblasts (Fig. 7a; Supplementary Fig. 12a). PRDX2 depletion also suppressed accumulation of R-loops and micronucleus formation in HRAS-overexpressing cells (Fig. 7b-e; Supplementary Fig. 12b, c), as did ZRANB3 depletion (Fig. 3g; Supplementary Fig. 12d-f), which promotes replication restart 14. Importantly, PRDX2 depletion had no effect on the replication stress phenotypes of cyclin E1-overexpressing cells (Supplementary Fig. 13a-d). These data suggest that oncogenic RAS induces R-loop-mediated fork stalling in a PRDX2-dependent manner.

To test whether HRASV12 overexpression triggers PRDX2-mediated dissociation of the TIMELESS-TIPIN complex from the replisome, we used SIRF, a PLA-based assay that enables quantitative analysis of protein interactions at active or stalled replication forks 52. In this assay, 5-ethynyl-2’-deoxyuridine (EdU) is first incorporated into nascent DNA at replication forks, followed by formaldehyde crosslinking and EdU biotinylation via click chemistry. PLA is then performed between biotin-labeled nascent DNA and the protein of interest. We performed SIRF against TIMELESS prior to and after induction of HRASV12 expression in U2OS-HRASV12-TetON cells transfected with control or PRDX2 siRNA. Concomitantly, we co-clicked EdU with Alexa Fluor 647-azide to mark S-phase cells. QIBC analysis revealed that the TIMELESS-SIRF signal in S-phase nuclei decreased significantly upon HRASV12 overexpression in control siRNA-transfected cells. This reduction was not observed in PRDX2-depleted cells, suggesting that HRASV12 induces PRDX2-dependent dissociation of TIMELESS from the replisome (Fig. 8a, b). In contrast to HRASV12, overexpression of cyclin E1 only mildly reduced the association of TIMELESS with the sites of DNA replication in PRDX2-proficient cells, with a further decrease observed upon PRDX2 depletion (Supplementary Fig. 13e, f). These findings support the conclusion that R-loop-mediated fork stalling induced by oncogenic RAS is a consequence of ROS-induced dissociation of the TIMELESS-TIPIN complex from the replisome, which slows down replication fork progression.

Methods
Plasmid constructions and protein purification
The plasmids pBABEneo-HRASV12 was kindly provided by Dr. Fabrizio d’Adda di Fagagna. For generation of empty vector (pBABEneo), pBABEneo-HRASV12 was digested with BamHI and EcoRI to excise the HRASV12 insert. The cleaved vector was religated by T4 DNA ligase (NEB, M0202S) after filling in DNA ends with Klenow Fragment (NEB, M0210S). pBABEneo-CCNE1 was constructed by inserting the Bsp1407I/HindIII fragment of pLXSNneo-CCNE1 (kindly provided by Dr. Stefano Ferrari) including the CCNE1 gene between the Bsp1407I and HindIII sites in pBABEneo. The RNH1(D210)-GFP sequence was amplified by PCR from the plasmid pAIOhM27RNaseH1-EGFP D210N 32 using the following primers: 5'-GGATCACATATGTTCTATGCCGTGAGGAGG-3' (forward); 5'-CATGAGAATTCCCATCTTGTACAGCTCGTCC ATGCC-3’ (reverse). The resulting PCR product was digested with NdeI and EcoRI and cloned along with the EcoRI/BspEI fragment of pQE60-yRad52-Tat11-His6 59 in the plasmid pTXB1 (New England Biolabs) to create a bacterial expression vector for RNH1(D210N)-GFP-Tat11-His6 fusion protein. Lentiviral vectors for inducible expression of HRASV12 and cyclin E1, respectively, were constructed by PCR amplification of the respective ORF [primers for HRASV12: 5’-GCAGCAGAATTCGCCACCATGACTGAATACAAG-3’ and 5’-GCAGCAGGATCCTCAGGACAGCACACATTTGC-3’; primers for CCNE1: 5’-GCAGCAGAATTCGCCACCATGCCGAGGGAGCGCAGGG-3’ and 5’-GCAGCAGGATCCTCACGCCATTTCCGGCCCGC-3’] and inserting the resulting PCR product into pLVX-TetOne-Puro (Clontech Laboratories, 631849) between EcoRI and BamHI sites.

RNH1(D210N)-GFP-Tat11-His6 protein was produced in E. coli BL21-CodonPlus (DE3)-RIL (Agilent Technologies, 230245) transformed with the above pTXB1-base vector. Protein expression was induced by 0.1 mM β-D-1-thiogalactopyranoside (IPTG) at 16°C for 16 h. Cells were lysed by sonication in buffer A [50 mM Tris-HCl (pH 7.5), 300 mM NaCl, 10% (v/v) glycerol, and 2 mM 2-Mercaptoethanol] supplemented with 10 mM imidazole, 0.1 mM phenylmethylsulfonyl fluoride (PMSF), and protease inhibitor cocktail (Roche cOmplete, EDTA free). Cell extract was clarified by ultracentrifugation and loaded onto 5-ml HisTrap FF column (Cytiva, 17531901). Bound proteins were eluded with a linear gradient of imidazole (40-300 mM) in buffer A. Peak fractions with RNH1(D210N)-GFP-Tat11-His6, determined by SDS-PAGE, were pooled and diluted with buffer B [50 mM Tris-HCl (pH 7.5), 0.1 mM EDTA, 10% (v/v) glycerol, and 1 mM dithiothreitol (DTT)] to a final concentration of NaCl of 150 mM. The sample was loaded onto a 5-ml HiTrap SP HP column (Cytiva, 17115101) equilibrated with buffer B/150 mM NaCl. Bound proteins were eluted with a 50-ml linear gradient of NaCl (150-500 mM) in buffer B. Peak fractions were pooled, dialyzed against 25 mM Tris-HCl (pH 7.5), 20% (v/v) glycerol, 0.1 mM EDTA 110 mM NaCl, and 0.1 mM PMSF, and stored at -80°C.

Cell culture
The following human cells lines we used in this study: U2OS (ATCC; HTB-96), HEK293T (ATCC; CRL-3216), BJ-hTert HRASV12ER-TAM 9, RPE1 CE-TetON (RPE1-pRetroX-Tet-On Advanced cells stably transfected with pRetroX- Tight-Pur-CCNE1) 60, U2OS ZRANB3-WT, U2OS ZRANB3-KO 36, Phoenix-AMPHO (ATCC; CRL-3213), HeLa Kyoto (Cancer Research UK, CVCL_1922), HeLa Kyoto MUS81 knockout 27, and U2OS T-REx (Invitrogen) cell lines carrying pAIO-based vectors encoding for RNaseH1-GFP or RNaseH1(D210N)-GFP 32. Cells were cultivated in Dulbecco's modified Eagle's medium (DMEM; Gibco, 41966-029) supplemented with 10% (v/v) fetal bovine serum (FBS; Tet-free approved, Gibco, 10270), 100 U/ml penicillin and 100 μg/ml streptomycin (Thermo Fisher Scientific, 15140122). Cells stably transfected with pAIO-based plasmids were selected in the presence of hygromycin B (50 μg/ml; Sigma-Aldrich, H3274) and puromycin (1 μg/ml, Sigma-Aldrich, P8833). Doxycycline (Takara Bio, 631311) was added for 24 h to induce overexpression of the RNaseH1 variants in U2OS T-REx cells (1 ng/ml) and for 72 h to induce overexpression of cyclin E1 in RPE1 CE-TetON (1 µg/ml). 4-Hydroxytamoxifen (Sigma-Aldrich, SML1666) at a concentration of 333 nM was added for 72 h to induce overexpression of HRASV12 in BJ-hTert HRASV12ER-TAM cells. Primary BJ HRASV12-TetON and BJ CE-TetON fibroblasts were obtained from Prof. Jiri Bartek 6, and grown in DMEM, low glucose (Gibco, 31885-023) supplemented with 10% (v/v) FBS, 100 U/ml penicillin, 100 μg/ml streptomycin and non-essential amino acids (Gibco, 11140050). Oncogene expression was induced by addition of doxycycline (2 µg/ml) for 48 h.

Retroviral transduction
For production of retroviruses, 5.5 x 106 of Phoenix-AMPHO cells were seeded to a 10-cm Petri dish 24 h prior to transfection of retroviral vectors. 1 h before transfection, cells were treated with 25 μM chloroquine (Sigma-Aldrich, C6628). For transfection of retroviral vectors, 500 μl of 2x HBS (50 mM HEPES, 10 mM KCl, 12 mM Dextrose, 280 mM NaCl, 1.5 mM Na HPO x 7H O; pH 7.04) were mixed with 500 μl CaCl plasmid DNA mix [61 μl of 2 M CaCl , 10 μg of retroviral vector], incubated for 5 min at room temperature (RT) and added drop-wise to the plate with Phoenix-AMPHO cells containing 5.5 ml of medium. After 8 h, the medium was replaced with fresh medium and this was repeated once again 24 h after transfection. 48 h post-transfection, the medium containing retroviral particles produced by Phoenix cells was collected, sterile-filtered (0.22 μm, Millipore) and polybrene (Sigma-Aldrich, H9268-5G) was added to a concentration of 8 μg/ml. This virus-containing medium was then added to target cells, which were seeded 24 h before the transduction at a density of 1.2 x 106 cells per 10-cm Petri dish. After a 3-h incubation, the medium was changed to freshly filtered virus-containing medium, and the plate was again incubated for 3 h. After this second transduction step, target cells were cultured in non-viral medium overnight. The next day, third retroviral transduction was done. After 3 h, cells were incubated for 18 h in a fresh non-viral medium. The following day, transduced and non-transduced cells were re-plated to 50% confluency in growth medium containing the selection antibiotic G-418 (500 μg/ml; Thermo Fisher Scientific, 10131027). The selection took typically 96 h, when the non-transduced control died (Day 0). The transduced cells were then used for experiments. Phenotypic analyses were conducted 2 to 4 days after the end of selection. Where specified, cells were transfected with siRNAs on the day 0 and harvested 72 h later.

Generation of stable cell lines via lentiviral transduction
For lentivirus production, HEK293T cells (Invitrogen) were transfected with the packaging plasmids pVSV, pMDL and pREV (kindly provided by Prof. Beat Schäfer), and the expression plasmid pLVX-TetOne-Puro-HRASV12 or pLVX-TetOne-Puro-CCNE1 using TransIT-X2 Dynamic Delivery System (Mirus, MIR 6004) according to the manufacturer’s instructions. 16 h after the transfection, medium was replaced with fresh medium. 48 h post-transfection medium containing lentiviral particles produced by HEK293T cells was collected, filtered through a 0.45 µm filter syringe and supplemented with polybrene (10 µg/ml). This virus-containing medium was then added to target U2OS cells which were seeded 24 h before transduction. After 24 h incubation, the medium was replaced with fresh virus-free medium. Transduced cells were selected in the presence of puromycin (1 μg/ml; Sigma-Aldrich, P8833). Clones were isolated after 10 to 14 days of growth in the selection medium. The expression of HRASV12 and cyclin E1 after induction with doxycycline (1 µg/ml, 48 h) was tested by western blotting.

Small-interfering RNA transfection
Transfections of siRNAs (a final concentration of 40 nM, in the case of double depletion 20 nM each) were done using Lipofectamine RNAiMAX (Thermo Fisher Scientific, 13778150) according to the instructions of manufacturer (reverse transfection protocol). Cells were typically seeded 6 h before siRNA transfection to 50% confluency. 24 h after siRNA transfection, the medium was replaced with fresh medium, and cells were replated if needed. Experiments were carried out 72 h after siRNA transfection. The majority of siRNA oligonucleotides used in this study were purchased from Microsynth AG. The sequences of the sense strand of siRNA duplexes are: siLUC (CGUACGCGGAAUACUUCGA); siMUS81 (CAGCCCUGGUGGAUCGAUA); siLIG4 (GCUAGAUGGUGAACGUAUG ); siPRIMPOL (GAGGAAAGCUGGACAUCGA); siRECQ1 (GCAAGAGAUUUACUCGAA); siSMARCAL1 (UUGCUAAGAAGGUCAAAGC); siPRDX2 (AGAUCAUCGCGUUCAGCAA). ZRANB3 siRNA was purchased from Dharmacon (84083, D-010025-03-0005), ELL siRNA was purchased from Santa Cruz Biotechnology (sc-38041).

DNA fiber assay
Cells were pulse-labeled with 30 μM thymidine analog 5-chloro-2′-deoxyuridine (CldU) (Sigma-Aldrich, C6891) for 30 min, then washed three times with PBS (pre-warmed to 37°C), followed by pulse-labeling with 250 μM 5-iodo-2′-deoxyuridine (IdU) (Sigma-Aldrich, I7125) for 30 min. Where required, cells were pre-treated with olaparib (10 μM; Selleckchem, S1060), triptolide (1 μM; Sigma-Aldrich, T3652), setanaxib (1 or 10 μM; Merck, S7171), sulforaphane (10 μM; Merck, 574215), or tert-butylhydroquinone (10 μM; Sigma-Aldrich, 112941) as specified in Figure legends. Labeled cells were washed with PBS, harvested by trypsinization, and resuspended in PBS to a concentration of 2.5x105 cells/ml. 2.5 μl of this cell suspension was mixed with 7.5 μl of lysis buffer [200 mM Tris–HCl (pH 7.5), 50 mM EDTA, 0.5% (w/v) SDS] directly on a slide and incubated for 5 min followed by tilting the slides to 30° causing the drops to run down by gravity. The spreads were then air-dried and fixed with methanol/acetic acid (3:1) at RT for 20 min. DNA was denatured in 2.5 M HCl for 1 h at RT and later washed three times with PBS. Slides were blocked in blocking solution [2% (w/v) BSA, 0.1% (v/v) Tween 20 in PBS] for 20 min and incubated with primary antibodies: rat monoclonal anti-BrdU (Abcam, ab6326; 1:500; detection of CldU) and mouse monoclonal anti-BrdU (BD Biosciences, 347580; 1:100; detection of IdU), in blocking solution for 2.5 h at RT in a humid chamber. Afterward, the samples were washed 3 times in PBS-T [0.2% (v/v) Tween 20 in PBS] and incubated with secondary antibodies: Cy3-conjugated donkey anti-rat secondary antibody (ImmunoResearch Europe, 712-166-153; 1:150) and goat anti-mouse Alexa Fluor 488 secondary antibody (Thermo Fisher Scientific, A11001; 1:300), in blocking solution for 1.5 h at RT in a humid chamber. The slides were washed in PBS-T and mounted using Fluoromount-G (Thermo Fisher Scientific, 00-4958-02) mounting media. Images were acquired with a Leica DM6000 upright fluorescent microscope (63×/1.40 oil immersion). CldU and IdU tract lengths were measured using ImageJ (version 1.54F).

S1 nuclease assay
Cells were pulse-labeled with CldU and IdU as described above. Subsequently, cells were washed with PBS and pre-extracted in CSK buffer [25 mM HEPES (pH 7.7), 50 mM NaCl, 1 mM EDTA, 3 mM MgCl , 300 mM sucrose, 0.5% (v/v) Triton X-100] for 10 min at RT. Cells were prewashed with S1 buffer [30 mM Sodium Acetate (pH 4.6), 10 mM Zinc Acetate, 5% (v/v) glycerol, 50 mM NaCl] and incubated in S1 buffer with and without S1 nuclease (20 U/ml; Thermo Fisher Scientific, EN0321) for 30 min at 37 °C. After incubation, cells were washed with 0.1% (w/v) BSA in PBS and scraped into an Eppendorf tube. Cells were pelleted by centrifugation at 4600 g for 10 min. The pellets were resuspended in PBS to a concentration of ~1500 cells per μl. 2.5 μl of this cell suspension was mixed with 7.5 μl of lysis buffer [200 mM Tris-HCl (pH 7.5), 50 mM EDTA, 0.5% (w/v) SDS] directly on a slide and incubated for 4 min followed by tilting the slides to 30° causing the drops to run down by gravity. The rest of the procedure was identical to the DNA fiber assay described above.

Immunofluorescence staining and analysis
For visualization of RNH1(D210N)-GFP foci and γH2AX foci, cells grown on autoclaved coverslips were washed with ice-cold PBS and pre-extracted for 5 min with CSK [25 mM HEPES–NaOH (pH 7.7), 50 mM NaCl, 1 mM EDTA, 3 mM MgCl , 0.3 M sucrose, 0.5% (v/v) Triton X-100] buffer on ice. For staining with recombinant RNH1(D210N)-GFP-Tat11 protein, cells were pre-extracted with 0.2% (v/v) Triton X-100 in PBS for 5 min on ice. Subsequently, cells were fixed with 4% (v/v) formaldehyde (Sigma-Aldrich, F8775) in PBS for 15 min in the dark at RT. For PCNA staining, formaldehyde fixation was followed by fixation with ice-cold methanol for 20 min at -20 °C. For 53BP1 staining, coverslips were washed with ice-cold PBS, fixed with 4% (v/v) formaldehyde in PBS for 15 min in the dark at RT, and then permeabilized with 0.1% (v/v) Triton X-100 for 10 min at RT. After fixations, coverslips were washed with PBS and blocked with 1% (w/v) BSA (Sigma-Aldrich, A7030) in PBS for 10 min at RT. Slides were incubated with primary antibodies at RT or with recombinant RNH1(D210N)-GFP-Tat11 protein at 37 °C for 90 min, subsequently washed with PBS, and incubated with appropriate Alexa Fluor-conjugated secondary antibodies for 30 min. Finally, the coverslips were incubated with DAPI (1 µg/ml; Merck, D9542) and mounted using Fluoromount-G mounting medium (Thermo Fisher Scientific, 00-4958-02). Cells stained with recombinant RNH1(D210N)-GFP-Tat11 protein were mounted with ProLong™ Gold Antifade Mountant (Thermo Fisher Scientific, P36930). The primary antibodies used for immunofluorescence staining: rabbit polyclonal anti-PCNA antibody (Abcam, ab18197; 1:200), rat polyclonal anti-PCNA (Abcam, ab252848; 1:500) mouse monoclonal Cyclin A (Santa Cruz Biotechnology, sc-271682; 1:200), rabbit polyclonal anti-53BP1 (H-300) (Santa Cruz Biotechnology, sc22760; 1:200) and mouse monoclonal anti-γH2AX (Ser139) (Millipore, 05-636; 1:200). The secondary antibodies used for immunofluorescence staining: goat anti-mouse Alexa Fluor 488 IgG (Thermo Fisher Scientific, A11001; 1:400), goat anti-rabbit Alexa Fluor 647 IgG (Thermo Fisher Scientific, A-21245; 1:400), goat anti-rat Alexa Fluor 555 (Thermo Fisher Scientific, A21434; 1:500). Cell images were acquired on a Leica DM6000 upright fluorescent microscope (63×/1.40 oil immersion), an IX83 microscope (Olympus) equipped with the ScanR imaging platform using a 60x/1.42 OIL objective with oil immersion, or an IXplore SpinSR10 (Olympus) with a UPLAN S Apo 40x/0.95 NA objective. Images were analyzed with ScanR Analysis software (version 3.5.0) to measure intensity of PCNA, Cyclin A, and with the edge detector plugin to detect RNH1(D210N)-GFP foci, RNH1(D210N)-GFP-Tat11 foci, γH2AX foci, and 53BP1 foci. A minimum of 600 cells per condition were analyzed in most experiments. The DAPI signal was used for segmentation of the images to identify individual nuclei. To prepare scatter plots, the values were exported to the TIBCO Spotfire software (version 10.10.1). An equal number of cells per condition was randomly selected using the Excel RAND() function.

Proximity ligation assay
For PLA, we used Duolink reagents and followed the Duolink® PLA Fluorescence Protocol (Sigma-Aldrich). Cells grown on coverslips were incubated with 25 μM EdU (Invitrogen, A10044) for 30 min. Cells were pre-extracted and fixed as for immunofluorescence staining. Then cells were blocked with 3% (w/v) BSA in PBS for 10 min. EdU click reaction was performed in buffer containing 100 mM Tris-HCl (pH 8.5), 2 mM CuSO , 100 mM ascorbate, and 5 µM Alexa Fluor™ 488 Azide (Thermo Fisher Scientific, A10266) for 30 min in the dark followed by two washes with 1% (w/v) BSA in PBS. Coverslips were then incubated with primary antibodies: rabbit polyclonal anti-PCNA antibody (ab18197, Abcam ; 1:1000) and mouse monoclonal anti-RNA polymerase II RPB1 (Biolegend, 920204, 1:1000) overnight at 4°C. Next day, coverslips were washed and incubated with PLA-probes for 1 h at 37°C: Duolink In Situ PLA Probe Anti Mouse MINUS (Sigma-Aldrich, DUO92004) and Duolink In Situ PLA Probe Anti Rabbit PLUS (Sigma-Aldrich, DUO92002), followed by ligation for 30 min at 37°C. For amplification, Duolink In Situ Detection Reagents Green (Sigma-Aldrich, DUO92014) was used. Coverslips were then washed and stained with DAPI (1 µg/ml; Merck, D9542) and mounted using Fluoromount-G mounting medium (Thermo Fisher Scientific, 00-4958-02). Cell images were acquired on an IX83 microscope (Olympus) equipped with the ScanR imaging platform using a 60x/1.42 OIL objective with oil immersion. A minimum of 750 cells per condition were analyzed in most experiments. The DAPI signal was used for segmentation of the images to identify individual nuclei. To prepare scatter plots, the values were exported to the TIBCO Spotfire software (version 10.10.1). An equal number of cells per condition was randomly selected using the Excel RAND() function.

SIRF
U2OS cells growing on autoclaved coverslips were incubated with 25 μM EdU for 10 min, washed twice with ice-cold PBS, pre-extracted with cold 0,2% (v/v) Triton X-100 in PBS for 5 min on ice and fixed with 4% (v/v) formaldehyde solution for 15 min. Subsequently, cells were fixed with methanol for 20 min at -20 °C. EdU click reaction was performed in buffer containing 100 mM Tris-HCl (pH 8.5), 2 mM CuSO , 100 mM ascorbate, and 100 µM biotin azide (Click Chemistry Tools, 1265-5) for 30 min. PLA was performed using Duolink PLA technology (Sigma-Aldrich) according to the manufacturer’s instructions. Cells were incubated with primary antibodies: mouse monoclonal anti-biotin antibody (Jackson ImmunoResearch, 200-002-211, 1:200) and rabbit monoclonal anti-Timeless antibody (Abcam, ab109512, 1:1000) at 4 °C overnight. Next day, coverslips with cells were incubated with PLA probes followed by ligation and amplification according to the manufacturer’s protocol. Click reaction to detect EdU was performed in buffer containing 100 mM Tris-HCl (pH 8.5), 2 mM CuSO , 100 mM ascorbate, and 2 µM Alexa fluor 647 azide (Thermo Fisher Scientific, A10277). Coverslips were stained with DAPI (1 µg/ml; Merck, D9542) and mounted with ProLong™ Gold Antifade Mountant (Thermo Fisher Scientific, P36930). Cells images were acquired on a IXplore SpinSR10 (Olympus) using UPLAN S Apo 40x/0.95 NA objective. Images were analyzed with ScanR Analysis software (version 3.5.0). TIMELESS SIRF foci were detected using spot detector. A minimum of 500 cells per condition were analyzed. The DAPI signal was used for segmentation of the images to identify individual nuclei. Mean intensity of EdU was measured to determine S-phase cells.

5-EU incorporation assay
Cells were incubated with 1 mM 5-ethynyl uridine (5-EU; Invitrogen, E10345) for 30 min, fixed with 4% (v/v) formaldehyde in PBS for 15 min in the dark at RT and then permeabilized with 0.1% (v/v) Triton X-100 for 10 min at RT. For PCNA staining, formaldehyde fixation was followed by PBS washes and fixation with ice-cold methanol for 20 min at -20 °C. The coverslips were washed with PBS and blocked with 1% (w/v) BSA (Sigma-Aldrich, A7030) in PBS for 10 min at RT. 5-EU click reaction was performed in buffer containing 100 mM Tris-HCl (pH 8.5), 2 mM CuSO , 100 mM ascorbate, and 5 µM Alexa Fluor™ 488 Azide (Thermo Fisher Scientific, A10266) for 30 min in the dark followed by two washes with 1% (w/v) BSA in PBS. Immunostaining was done with primary rabbit polyclonal anti-PCNA antibody (Abcam, ab18197; 1:200) for 90 min and later with Alexa Fluor 647 goat anti-rabbit (Life Technologies, A21245; 1:400) for 30 min. Finally, the coverslips were incubated with DAPI (1 µg/ml; Merck, D9542) and mounted using Fluoromount-G mounting medium (Thermo Fisher Scientific, 00-4958-02). Cell images were acquired on an IX83 microscope (Olympus) equipped with the ScanR imaging platform using a 60x/1.42 OIL objective with oil immersion. Images were analyzed with ScanR Analysis software (version 3.5.0) to measure intensity of 5-EU and PCNA. Minimum of 700 cells per condition were analyzed. The DAPI signal was used for segmentation of the images to identify individual nuclei. To prepare scatter plots, the values obtained were exported to the TIBCO Spotfire software (version 10.10.1), displaying the same number of cells per condition. An equal number of cells per condition was randomly selected using the Excel RAND() function.

Slot blot
To detect RNA:DNA hybrids on a slot blot, cells were first harvested by trypsinization, washed with PBS, and centrifuged (1200 × g, 4°C, 3 min). Pellets were resuspended in lysis buffer [40 mM Tris–HCl (pH 7.5), 1.28 M sucrose, 20 mM MgCl₂, 4% (v/v) Triton X-100], incubated for 10 min on ice, and centrifuged (1200 × g, 4°C, 15 min). The supernatant was removed, and pellets were again resuspended in lysis buffer and centrifuged (1200 × g, 4°C, 15 min). Isolated nuclei were incubated for 2 h at 50°C with Proteinase K (0.74 mg/ml; Roche, 03115852001) in digestion buffer [30 mM Tris-HCl (pH 8.0), 800 mM guanidine, 30 mM EDTA, 5% (v/v) Tween 20, 0.5% (v/v) Triton X-100]. After incubation, the supernatant was mixed with an equal volume of chloroform:isoamyl alcohol (24:1), thoroughly mixed, and centrifuged (8000 × g, 4°C, 25 min). The aqueous phase was mixed with an equal volume of isopropanol, centrifuged (8000 × g, 4°C, 10 min) and the precipitated DNA was washed with ethanol and dissolved in TE buffer [10 mM Tris–HCl (pH 8.0), 1 mM EDTA]. Genomic DNA concentration was measured by a NanoDrop. 1.5 µg of DNA was digested or not with 2.4 U of RNase H (NEB, M0297) at 37°C for 3 h. The DNA samples (750 ng) were spotted in duplicates onto nylon membranes (Hybond-N+; GE Healthcare, RPN203B) using a slot blot manifold (Cleaver Scientific, CSL-S48) and washed with 2xSSC solution [0.3 M sodium chloride, 30 mM trisodium citrate, pH 7.0]. For S9.6 antibody detection, the membrane was dried, UV-crosslinked (0.12 J/m²), and after blocking in 5% (w/v) Skimmed Milk Powder (Coop, Switzerland) in TBS-T, incubated with mouse monoclonal S9.6 antibody (Kerafast, ENH001; 1:1000). As a loading control, we used the ssDNA signal. Membrane was dried, denatured in denaturing solution (0.5 M NaOH, 1.5 M NaCl) for 10 min and subsequently neutralized in 0.5 M Tris–HCl (pH 7.2) containing 1.5 M NaCl for 10 min. The membranes were UV-crosslinked (0.12 J/m²) and immunostained with mouse monoclonal anti-ssDNA antibody (Millipore, MAB3868; 1:10,000). Both membranes were later incubated with goat anti-mouse IgG-HRP (Sigma-Aldrich, A4416) to detect RNA:DNA hybrids and total DNA, respectively, with chemiluminescence reagents (Pierce ECL Western Blotting Substrate, Thermo Fisher, 32209 and SuperSignal™ West Dura Extended Duration Substrate; Thermo Fisher Scientific, 34075). Signal quantification was performed using ImageJ (version 1.54F).

Preparation of cell extracts and western blot analysis
Cells were trypsinized, washed with PBS and resuspended in lysis buffer [50 mM Tris–HCl (pH 7.5), 120 mM NaCl, 0.5% (v/v) Nonidet P-40] supplemented with protease inhibitor cocktail (Roche, 11873580001) and incubated for 5 min on ice. The cell suspension was sonicated with a Diagenode sonicator. Cell lysates were clarified by centrifugation at 16 100 × g for 10 min at 4°C and protein concentration in cell extracts was determined using the Bradford assay (Bio-Rad). Samples (15-50 μg of total protein) were mixed with Laemmli SDS sample buffer, boiled at 95 °C for 10 min and loaded onto 8-12% SDS-PAGE gel. After electrophoresis, separated proteins were transferred onto a nitrocellulose membrane (VWR international, 10600003) in a wet-transfer apparatus (Bio-Rad, 350 mA, 90 min, 4°C) with buffer containing 2.5 mM Tris–HCl, 10% (v/v) methanol and 19.2 mM glycine. The membrane was blocked with 3% (w/v) milk in TBS-T [20 mM Tris–HCl (pH 7.5), 150 mM NaCl, 0.1% (v/v) Tween-20] for 30 min at RT and incubated with primary antibodies (diluted in 3% milk/TBS-T) overnight at 4°C. Afterward, membranes were washed 3 times in TBS-T and incubated with appropriate horseradish peroxidase-coupled (HRP) secondary antibody in TBS-T for 30 min at RT. Membranes were washed in TBS-T and protein bands were detected with a chemiluminescence reagent [Pierce ECL Western Blotting substrate (Thermo Fisher Scientific, 32209), SuperSignal™ West Dura Extended Duration Substrate (Thermo Fisher Scientific, 34075) or SuperSignal™ West Femto Maximum Sensitivity Substrate (Thermo Fisher Scientific, 34094)]. The primary antibodies used for western blotting: HRAS (259) rat monoclonal (Santa Cruz Biotechnology, sc-35; 1:2000); HRAS (C-20) rabbit polyclonal (Santa Cruz Biotechnology, sc-520; 1:500); HRAS rabbit polyclonal (GeneTex, GTX 116041; 1:500), Cyclin E (HE12) mouse monoclonal (Santa Cruz Biotechnology, sc-247; 1:700); TFIIH p89 (S-19) rabbit polyclonal (Santa Cruz Biotechnology, sc-293; 1:1000); RNASE H1 (A-9) mouse monoclonal (Santa Cruz Biotechnology, sc-365783; 1:500); ZRANB3 rabbit polyclonal (Proteintech, 23111-1-AP; 1:1000); GAPDH (D16H11) rabbit monoclonal (Cell Signaling Technology, 5174; 1:2000); GAPDH (0411) mouse monoclonal (Santa Cruz, sc-47724, 1:1000); SMARCAL1 (D3P5I) rabbit monoclonal (Cell Signaling Technology, 44717S; 1:2000); β-Tubulin (TUB 2.1) mouse monoclonal (Sigma-Aldrich, T4026; 1:1000); MUS81 (B-12) mouse monoclonal (Santa Cruz Biotechnology, sc-376661; 1:500); MUS81 (MTA30 2G10/3) mouse monoclonal (Santa Cruz Biotechnology, sc-53382; 1:500); PRIMPOL rat polyclonal (a gift from Juan Mendez); RECQ1 rabbit polyclonal (Novus Biological, NB100-618; 1:1000); LIG4 (D-8) mouse monoclonal (Santa Cruz Biotechnology, sc-271299; 1:500); ELL (B-4) mouse monoclonal (Santa Cruz Biotechnology, sc-398959; 1:500); PRDX2 (A-2) mouse monoclonal (Santa Cruz Biotechnology, sc-515428; 1:100). The secondary antibodies used for western blotting: goat anti-rabbit IgG-HRP (Sigma-Aldrich, A0545; 1:5000); goat anti-mouse IgG-HRP (Sigma-Aldrich, A4416; 1:5000) and goat anti-rat IgG-HRP (Sigma-Aldrich, A9037; 1:5000).

Electron microscopy
U2OS T-REx [RNH1-GFP] cells were transduced with HRASV12, cyclin E1 or empty retroviral vectors and either treated with doxycycline (1 ng/ml) for the last 24 h to induce RNaseH1-GFP expression or left untreated. For isolation of genomic DNA, 15-27×106 cells were used per sample. All subsequent steps were performed on ice. Cells were pelleted (600 × g, 5 min, 4°C) and washed with ice-cold PBS. Genomic DNA was then cross-linked by incubating cells with 200 µg/ml trimethylpsoralen for 5 min followed by irradiation with 365-nm UV light for 7 min on a precooled metal plate. This procedure was repeated three times. Cells were then lysed in lysis buffer [1.28 M sucrose, 40 mM Tris-HCl (pH 7.5), 20 mM MgCl , 4% (v/v) Triton X-100]. Nuclei were pelleted (1300 × g, 15 min, 4°C) and incubated for 3 h at 50°C in 5 ml of digestion buffer [800 mM guanidine-HCl, 30 mM Tris-HCl (pH 8.0), 30 mM EDTA, 5% (v/v) Tween-20, 0.5% (v/v) Triton X-100, 1200 U proteinase K]. Genomic DNA was isolated via chloroform/isoamylalcohol method, precipitated with isopropanol, and resuspended in TE buffer [10 mM Tris-HCl (pH 8.0), 1 mM EDTA]. Genomic DNA was then digested with PvuII HF (33 U per 10 µg of DNA; NEB, R3151L) plus 33.3 µg of RNase A (Thermo Fisher Scientific, EN0531) and 0.013 U ShortCut RNase III (NEB, M0245S) in CutSmart restriction buffer (3 h, 37°C) in a total volume of 250 µl. Digested DNA was concentrated and recovered using Microcon DNA Fast Flow Centrifugal Filters (Millipore, MRCF0R100), and DNA concentration was measured. DNA was then spread on a water surface using the benzyldimethylalkylammonium chloride (BAC) method, transferred onto the carbon-coated 400 mesh copper grids and coated with platinum using the Leica EM ACE900 sample preparation system. Grids were examined using the Jeol JEM-1400 Flash transmission electron microscope operated at ≤120 kV (0.2 nm resolution). Images were acquired with a Jeol Flash 2000 × 2000 pixels CMOS camera using the serial EM capturing mode for automated acquisition. Replication forks were analyzed with ImageJ (version 1.54F). For each sample, ~200 replication forks from three independent biological replicates were analyzed.

Analysis of bulky anaphase bridges
Cells grown on autoclaved coverslips were synchronized at G2/M phase transition by culturing in the presence of 9 µM RO-3306 (Merck, SML0569) for 16 h. This was followed by three brief washes with PBS for 5 min at RT and subsequent incubation in DMEM medium for a total time of 1.5 h at 37°C. Cells were fixed with 4% (v/v) formaldehyde in PBS for 15 min at RT, followed by staining with DAPI (1 µg/ml). Cell images were acquired on an IX83 microscope (Olympus) equipped with ScanR imaging platform using a 40×/0.95 dry objective. The percentage of anaphase cells with bulky bridges was determined using ImageJ (version 1.54F). At least 25 anaphase cells were scored per condition in each experiment.

Analysis of micronuclei
Cells were grown on autoclaved coverslips in the presence of the cytokinesis inhibitor cytochalasin B (2 μg/ml, Sigma-Aldrich, C6762) for 16 h, followed by a wash with PBS. Where required, cells were treated with 1 μM setanaxib, 10 μM sulforaphane or 10 μM tert-butylhydroquinone as specified in Figure legends. The cells were washed with PBS and fixed with 4% (v/v) formaldehyde in PBS for 15 min at RT and stained with DAPI (1 μg/ml, Sigma-Aldrich, D9542) for 2 min at RT in the dark. Images were acquired using a Leica DMI 6000 inverted microscope at 20x or 40x magnification or IX81 microscope (Olympus) equipped with a ScanR imaging platform using a 40x/1.3 NA objective with oil immersion. The percentage of binucleated cells with micronuclei was determined using ImageJ (version 1.54F). At least 100 binucleated cells were scored per condition in each experiment.

Statistical analysis
Statistical analysis was performed using GraphPad Prism 8 software (version 8.4.3). Details of how data are presented, including the definition of center (mean or median) and error bars, as well as the details of statistical tests for each experiment, including the type of statistical tests used and the number of repeats, can be found in the figure legends. Statistical test results, presented as p-values, are shown in the figures. Statistical differences in scatter plots of data from DNA fiber assays, QIBC analysis of 5-EU intensity and total number of PCNA/RNAPII PLA foci, γH2AX foci, 53BP1 foci, TIMELESS SIRF foci, RNH1(D210N)-GFP foci, and RNH1(D210N)-GFP-Tat11 foci were determined by a two-tailed non-parametric Kruskal-Wallis test followed by a Dunn’s multiple comparisons test. Statistical differences in the plot of average number of 53BP1 foci were determined by two-tailed paired t-test. Statistical differences for other grouped analyses, i.e., micronuclei, anaphase bridges, average number of γH2AX foci, or frequency of fork reversal were assessed by a repeated-measures one-way ANOVA followed by a Tukey’s multiple comparisons test.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "Manuscript: MUS81/PRIMPOL-dependent DNA synthesis counteracts R-loop-induced replication fork reversal to prevent chromosome damage in mitosis upon oncogene activation\nThe manuscript by Oravetzova et al. presents insights into the mechanisms underlying DNA replication stress induced by overexpression of oncogenic HRASV12 or Cyclin E1. The authors demonstrate that replication stress and DNA damage associated with HRASV12 or Cyclin E1 activation are driven by the accumulation of S-phase-specific R-loops. These R-loops promote replication fork reversal, leading to mitotic DNA damage and micronuclei formation. Supporting this model, the authors show that inhibition of PARP1 or depletion of ZRANB3—both of these conditions, which suppress fork reversal—results in unrestrained fork progression and rescues mitotic DNA damage in cells overexpressing HRASV12 or Cyclin E1. Based on these findings, the authors propose that persistent reversed replication forks are a significant source of mitotic DNA damage during oncogene-induced replication stress.\nFinally, they show that this unrestrained fork progression in HRASV12- and Cyclin E1-overexpressing cells depends on MUS81 and is accompanied by PRIMPOL-mediated single-stranded DNA gap formation.\nOverall, the experiments are generally well-conducted and supported by clear results. However, a key limitation of the manuscript is that the novelty of the insights is somewhat limited. The link between oncogene activation—such as HRASV12 or Cyclin E1—and replication stress, including replication fork slowing and the interplay between global transcription and R-loop formation, is well-established (e.g., DOI: 10.1038/onc.2012.387, DOI: 10.1038/ncomms13087). Similarly, the role of the MUS81–LIG4–PRIMPOL axis in mediating replication restart following R-loop-induced replication stress—though previously studied under different stress conditions—has already been demonstrated by the authors themselves (DOI: 10.1038/ncomms13087) and is here extended to the context of HRASV12- and Cyclin E1-overexpressing cells.\nThat said, the authors do note an interesting distinction between HRASV12 and Cyclin E1 in their ability to induce global transcription (Supplementary Figure 3b), even though both oncogenes ultimately lead to comparable levels of S/G2-specific R-loops (Figure 2a–c) and a similar rescue of mitotic DNA damage upon R-loop disruption (Figure 1a–h). A more systematic characterization of these differences—particularly in the context of the well-established role of Cyclin E1 in promoting new origin firing—could have provided valuable insights into the distinct upstream mechanisms leading to R-loop formation and replication stress. Such analysis might have clarified how divergent oncogenic triggers can converge on a common outcome—R-loop accumulation—as a key driver of replication stress and genome instability.\nBelow are some comments and suggestions that authors could improve and strengthen before publication.\nFigures 1 and 2:\n- In general, the experiments are well-performed. However, the manuscript lacks a comprehensive overview of cell cycle distribution and the extent of DNA damage or replication stress specifically within S-phase.\n- Since the authors utilize QIBC in Figure 2, a similar approach could be applied to assess S-phase-specific replication stress markers.\n- Along the same lines, it would be informative to evaluate R-loop levels (Figure 2c) upon HRASV12 or Cyclin E1 expression in the presence of CDC7 inhibition. This would help clarify how deregulated origin firing under these conditions influences R-loop formation and could be further extended to include replication fork speed measurements.\nFigure 3:\n- The authors use ZRANB3 depletion to disrupt fork reversal. It would strengthen the conclusions to test whether similar phenotypes are observed upon depletion of other fork reversal enzymes such as HLTF or SMARCAL1?\n- To further elucidate causal relationships, it would be valuable to assess R-loop levels (as shown in Figure 2c) following ZRANB3 or other fork reversal enzyme depletions. This could help determine if R-loops persist or resolve under conditions of oncogene-induced replication stress and unrestrained fork progression.\nFigures 4 and 5:\n- While the MUS81–LIG4 axis appears to play a critical role in promoting fork progression under oncogene-induced stress conditions, the lack of a comprehensive overview of S-phase DNA damage, global DNA synthesis, and the extent of R-loop accumulation makes it difficult to conceptualize the precise nature and extent of fork stalling that necessitates nucleases like MUS81.\n- It is also puzzling that ZRANB3 knockout cells, which replicate with persistent ssDNA gaps, nevertheless show a rescue of mitotic defects typically induced by oncogene-driven replication stress. This raises the question of whether these cells rely on daughter strand gap repair mechanisms, such as RAD51-mediated homologous recombination. This possibility could be addressed by assessing spontaneous RAD51 foci formation during S-phase (e.g., as described in DOI: 10.1016/j.devcel.2021.01.011).\nOther points:\n- The authors implicate the potential incidence of head-on transcription–replication conflicts as a driver of R-loop formation and replication stress. However, this has not been directly tested. It would be informative to apply genomics-based approaches, for R-loop mapping in relation to replication origin orientation, to explore TRC landscapes in HRASV12 versus Cyclin E1-overexpressing cells. Alternatively, it should be phrased cautiously.\n- It is unclear why 5-FU was used to assess global transcriptional activity instead of more conventional and direct methods such as EU (5-ethynyl uridine) incorporation, which is widely used for nascent RNA labelling.", "reviewer_2": "Oravetzova A. et al. show how an increase of transcriptional levels or replication origin firing mediated by oncogene overexpression are linked to genomic instability associated to R-loop formation in S phase and fork reversal. They also showed that inhibition of fork reversal promotes fork progression re-start and suppression of mitotic DNA damage mediated by MUS81 and PRIMPOL repriming. Reinforcing in the idea that fork reversal have both a protective but also a damaging role if not resolved properly. The manuscript presents a huge amount of work. While the results presented are solid and it is an interesting observation, I do have some concerns regarding the novelty of the presented data.\nMayor comments\nThe authors propose that oncogene overexpression (OE) leads to R-loop formation specifically in S phase, possibly due to TRCs, and inducing persistent replication fork reversal. The RNH1 foci seen in G2 phase in Fig2C can represent persistent R-loops formed in S phase as the author suggested but also, R-loop formed in G2 if late replication and TRCs occurs in G2/M phases. Late replication has been shown to occur at TSS regions that is where more R-loops are formed (Wang J. et al Cell Report 2021). This possibility could be tested by for example checking RNH1 foci when inhibiting transcription during S phase and reactivating it during G2, or inhibiting fork reversal resolution and checking whether the frequency increases From S to G2.\nAlso, to confirm that it is produced as a result of TRC, authors could check directly measuring the frequency of EdU-RNAPII or PCNA-RNAPII interactions by PLA assays after oncogene OE.\nThe authors also propose that the formation of fork reversal in HRASV12 OE could be due to a slow fork progression mediated by ROS. Their data of fork velocity in the presence of RNH1 OE (Fig1B) support the idea that something else apart from R-loops is delaying the movement of the fork. They could check whether TIMELESS/TIPIN is indeed detached from the replisome and whether this is also the case of Cyclin E OE.\nIs ATR active in both oncogene OE conditions?", "reviewer_3": "In this manuscript, the authors show that HRAS or cyclin E1 overexpression results in enhanced replication fork reversal and increased mitotic DNA damage, which is primarily caused by R-loop mediated transcription-replication conflicts (TRåCs) during S-phase. Inhibition of replication fork reversal creates a condition of unrestrained replication fork progression which prevents mitotic DNA damage in oncogene overexpressing cells. This process engages the MUS81/LIG4/EEL axis and PRIMPOL-mediated repriming to support unrestrained fork progression, where the presence of persistent reversed forks contributes to enhanced fork slowdown/stalling and an increase in mitotic DNA damage. Overall, experimental evidence is solid and well-controlled. Understanding the nature of DNA replication stress caused by oncogenes is considered significant. Nevertheless, results of this study seem incremental; it is uncertain how much new molecular insight can be deduced from the findings presented here. It is already well-established in the field that (1) oncogene activation, including Cyclin E, causes TRCs, (2) R-loop formation is associated with frequent fork reversal, (3) unrestrained fork progression upon PARP inhibition is dependent the MUS81 activity and the RECCQ1 helicase, and (4) restart of stalled forks, including R-loops, via MUS81 requires repriming mediated by PRIMPOL, all of which are the key takeaways of this study. In this case, overexpression of oncogenes is used to trigger DNA replication stress - unless new mechanistic insight of how HRASG12V and Cyclin E uniquely compromises replication fork stability and causes R-loop is presented, the current study lacks sufficient molecular details and is only limited to the phenotypic analyses using loss-of-function approaches (e.g., siRNA or knockout).\n1. One major concern is that the phenotypic analyses are limited to measuring DNA fiber tracks and there is little molecular insight to explain mitotic DNA damage. This is a major weakness of the manuscript in its current form.\n2. Figure 1 to establishes R-loop as a source of oncogene-induced replication stress is performed only in a cancer cell line (U2OS), which has already undergone a transformation process. On the other hand, Fig. S4 employs immortalized yet non-transformed fibroblast or epithelial cells. The authors should at least confirm the key results of Figure 1 using non-transformed cells.\n3. In Figure 1, it should be tested whether RNaseH1 expression can suppress DNA damage markers (e.g.,  H2AX). In relation to this, the nature of DNA lesions (e.g., the status of DSBs) has not been scrutinized. What would be the underlying basis of DNA damage that contributes to defective chromosome segregation and mitotic DNA damage after R-loop accumulation?\n4. The authors uses RNaseH1-dependent rescues of fork progression as sole evidence to support that R-loops are the cause of oncogene-induced replication stress. However, there is no direct evidence of R-loop formation in these oncogene overexpressing cells. This could be validated by using the S9.6 antibody previously used by the lab in Ref#25 either through immunofluorescence or DRIP-seq. A positive control such as PDS or even HU or APH can be included as mentioned by the authors.\n5. 5-FU staining suggests that there is an increase in transcriptional activity in G1, S, and G2/M cells that are overexpressing HRASG12V but not in cyclin E1 overexpressing cells, while the authors claim that R-loops seen in S-phase are carried over into mitosis. Can the authors exclude the possibility of R-loop formation within G2/M itself independently of DNA replication?\n6. The authors show that ZRANB3 completely prevents oncogene-induced anaphase bridges and micronuclei (Fig. 3). It is an interesting result since other translocases such as SMARCAL1 or HLTF are also known to be involved in fork reversal. These factors should be tested to determine how much individual translocase contributes to the fork reversal process. If solely dependent on ZRANB3, some mechanistic explanation should be presented considering the way ZRANB3 acts to promote fork reversal.\n7. It would be important to test whether PRIMPOL’s primase activity is actually required for unrestrained replication fork progression.\n8. Figure 6 model – there is no experimental evidence supporting that ZRANB3 deficiency rescues the mitotic defect. In relation to this, the physiological relevance of inhibiting fork reversal is not clear. The phenotype of unrestrained DNA synthesis is revealed when cells are forced to lose fork reversal process, and whether unrestrained DNA synthesis plays a meaningful role in normal DNA replication and mitosis is not addressed. Additionally, there is no mechanism of fork remodeling and processing that is uniquely engaged by oncogenes."}, "author_response": "Point-by-point response to the reviewers' comments (NCOMMS-25-21283A)\nWe would like to thank the reviewers for taking the 4me to carefully assess our manuscript\nand for providing helpful and construc4ve feedback that helped us significantly improve the\nmanuscript.\nAnswers to specific ques4ons of reviewers:\nReviewer #1 (Remarks to the Author):\nManuscript: MUS81/PRIMPOL-dependent DNA synthesis counteracts R-loop-induced\nreplica4on fork reversal to prevent chromosome damage in mitosis upon oncogene ac4va4on\nThe manuscript by Oravetzova et al. presents insights into the mechanisms underlying DNA\nreplica4on stress induced by overexpression of oncogenic HRASV12 or Cyclin E1. The authors\ndemonstrate that replica4on stress and DNA damage associated with HRASV12 or Cyclin E1\nac4va4on are driven by the accumula4on of S-phase-specific R-loops. These R-loops promote\nreplica4on fork reversal, leading to mito4c DNA damage and micronuclei forma4on.\nSuppor4ng this model, the authors show that inhibi4on of PARP1 or deple4on of ZRANB3—\nboth of these condi4ons, which suppress fork reversal—results in unrestrained fork\nprogression and rescues mito4c DNA damage in cells overexpressing HRASV12 or Cyclin E1.\nBased on these findings, the authors propose that persistent reversed replica4on forks are a\nsignificant source of mito4c DNA damage during oncogene-induced replica4on stress.\nFinally, they show that this unrestrained fork progression in HRASV12- and Cyclin E1-\noverexpressing cells depends on MUS81 and is accompanied by PRIMPOL-mediated single-\nstranded DNA gap forma4on.\nOverall, the experiments are generally well-conducted and supported by clear results.\nHowever, a key limita4on of the manuscript is that the novelty of the insights is somewhat\nlimited. The link between oncogene ac4va4on—such as HRASV12 or Cyclin E1—and\nreplica4on stress, including replica4on fork slowing and the interplay between global\ntranscrip4on and R-loop forma4on, is well-established (e.g., DOI: 10.1038/onc.2012.387, DOI:\n10.1038/ncomms13087). Similarly, the role of the MUS81–LIG4–PRIMPOL axis in media4ng\nreplica4on restart following R-loop-induced replica4on stress—though previously studied\nunder different stress condi4ons—has already been demonstrated by the authors themselves\n(DOI: 10.1038/ncomms13087) and is here extended to the context of HRASV12- and Cyclin\nE1-overexpressing cells.\nThat said, the authors do note an interes4ng dis4nc4on between HRASV12 and Cyclin E1 in\ntheir ability to induce global transcrip4on (Supplementary Figure 3b), even though both\noncogenes ul4mately lead to comparable levels of S/G2-specific R-loops (Figure 2a–c) and a\nsimilar rescue of mito4c DNA damage upon R-loop disrup4on (Figure 1a–h). A more\nsystema4c characteriza4on of these differences—par4cularly in the context of the well-\nestablished role of Cyclin E1 in promo4ng new origin firing—could have provided valuable\ninsights into the dis4nct upstream mechanisms leading to R-loop forma4on and replica4on\nstress. Such analysis might have clarified how divergent oncogenic triggers can converge on a\n1\ncommon outcome—R-loop accumula4on—as a key driver of replica4on stress and genome\ninstability.\nResponse: We would like to thank the reviewer for valuable sugges4ons on how to improve\nour manuscript. To address the reviewer’ issues regarding limited novelty, we have included,\nin the revised manuscript, the results of our follow-up project aiming to clarify the molecular\nmechanism leading to R-mediated fork stalling in HRASV12-expressing cells. As men4oned in\nour manuscript, apart from increasing global transcrip4on, oncogenic RAS is known to\nincrease the cellular levels of reac4ve oxygen species (ROS) by upregula4ng NADPH oxidases\n1 and 4 (Kodama et al., 2013; Mitsushita et al., 2004; Ogrunc et al., 2014; Weyemi et al., 2012).\nWork in Jiri Lukas lab has shown that excessive ROS disrupt the oligomeric state of the\nreplisome-associated ROS sensor peroxiredoxin 2 (PRDX2) and thereby enforce dissocia4on\nof the TIMELESS-TIPIN complex from the replisome, leading to a global reduc4on in replica4on\nfork velocity (Somyajit et al., 2017). Moreover, our recent study revealed that this ROS-\ninduced fork slowdown ul4mately causes R-loop-mediated fork stalling at sites of head-on\nTRCs (Andrs et al., 2023). We therefore asked whether ROS play a role in R-loop-mediated\nreplica4on stress induced by oncogenic RAS. Our experiments revealed that HRASV12-\ninduced replica4on stress is largely driven by ROS in a manner dependent on PRDX2 and is\nlinked to PRDX2-mediated release of the fork accelera4on factor TIMELESS from the\nreplica4on fork. On the other hand, replica4on fork stalling in cyclin E1-overexpressing cells\nwas found to be ROS- and PRDX2-independent, and is presumably a consequence of increased\nintragenic origin firing as reported previously (Macheret and Halazone4s, 2018). These new\nfindings, presented in Fig. 6-8, and Supplementary Fig. 11-13, and described on pages 11-13,\nprovide explana4on for our observa4on that HRASV12 induces R-loop forma4on in S phase\nbut not in G1, although it increases transcrip4on at all stages of the cells cycle. Moreover,\nthese findings argue against the model wherein HRASV12-induced replica4on stress stems\nfrom increased global transcrip4on due to upregula4on of TATA-box binding protein (TBP),\nwhich increases R-loop forma4on (Kotsan4s et al., 2016). Instead, we propose that oncogenic\nRAS induces R-loop-mediated fork staling by increasing the cellular levels of ROS, which in\nturn induce TIMELESS-TIPIN dissocia4on from the replisome, leading a reduc4on in replica4on\nfork velocity. Conflicts of such slowly moving forks with head-on transcrip4on complexes lead\nto R-loop forma4on which induces fork reversal and thereby halts fork progression. In support\nof this model, we also show that HRASV12-induced replica4on stress was rescued by NOX1/4\ninhibi4on (Fig. 6a, and Supplementary Fig. 11b). The rescue of HRASV12-induced replica4on\nstress by TBP deple4on reported by Eva Petermann lab could be explained by downregula4on\nof NOX1/4 enzymes, which generate ROS in HRASV12-expressing cells. This would be expected\nin case of a deficiency in a general transcrip4on factor such as TBP. Nevertheless, we cannot\nexclude the possibility that increased transcrip4on ac4vity enhances R-loop forma4on at sites\nof head-on TRCs in HRASV12-expressing cells. This is now men4oned in Discussion (page\n13/14). In any case, we think that our new findings provide important insight into the\nmolecular mechanism underlying DNA replica4on stress induced by oncogenic RAS.\nPlease note that, due to the addi4on of these new results, we have changed the 4tle of the\nmanuscript to “Mechanis)c insights into oncogene-induced replica)on stress”.\nBelow are some comments and sugges4ons that authors could improve and strengthen before\npublica4on.\n- In general, the experiments are well-performed. However, the manuscript lacks a\ncomprehensive overview of cell cycle distribu4on and the extent of DNA damage or\nreplica4on stress specifically within S-phase.\n- Since the authors u4lize QIBC in Figure 2, a similar approach could be applied to assess S-\nphase-specific replica4on stress markers.\nResponse: We tested the effect of HRASV12 and cyclin E1 overexpression on the forma4on of\nγH2AX and 53BP1 foci in different stages of the cell cycle. By QIBC analysis, we found that\noverexpression of either oncogene increased γH2AX foci in S-phase cells iden4fied by PCNA\nstaining. Notably, only a mild increase in 53BP1 foci - marker of DNA double-strand breaks -\nwas detected in S/G2 cells (cells posi4ve for cyclin A staining) upon overexpression of\nHRASV12 or cyclin E1, sugges4ng that replica4on fork stalling induced by these oncogenes\ndoes not lead to extensive DNA breakage during S-phase. Addi4onally, a modest increase\nin 53BP1 foci was detected in G1 cells upon overexpression of either oncogene, poten4ally\nreflec4ng unresolved under-replicated DNA from the previous cell cycle that led to DNA\nbreakage during chromosome segrega4on (Ying et al., 2013). This is consistent with the\nfinding that overexpression of these oncogenes increased the frequency of anaphase bridges\nand micronuclei (Fig. 1g, h; Supplementary Fig. 1e, f). Please note that the data from our\nanalysis of γH2AX and 53BP1 foci are now presented in Supplemetary Fig. 2 and described on\npage 6.\nAs suggested by the reviewer, we also tested the effect of HRASV12 or cyclin E1\noverexpression on cell cycle distribu4on. We did not observe any major differences in cell\ncycle distribu4on compared to control cells except for a small reduc4on in the propor4on of\nG1 cell in cyclin E1-overexpressing cell popula4ons, and a small reduc4on in the propor4on of\nS-phase cells in HRASV12-overexpressing cell popula4on. The former finding is consistent with\nthe finding of Halazone4s lab that cyclin E1 overexpression shortens the length of G1 phase.\nWe think that it is not necessary to include these data in the manuscript and therefore we\nshow them only in this response leqer (please see Fig. R1 below).\nFigure R1. Cell cycle profile of U2OS cells transduced with HRASV12 (RAS) or cyclin E1 (CE) retroviral vector.\nAsynchronously growing cells were treated with EdU for 30 min prior to harvest for FACS analysis to determine\nacHvely replicaHng cells. DAPI counterstaining was used to follow DNA content. EV, empty vector. Data are mean\n+/- SD, n = 4\n- Along the same lines, it would be informa4ve to evaluate R-loop levels (Figure 2c) upon\nHRASV12 or Cyclin E1 expression in the presence of CDC7 inhibi4on. This would help clarify\nhow deregulated origin firing under these condi4ons influences R-loop forma4on and could\nbe further extended to include replica4on fork speed measurements.\noverexpressing cells by trea4ng cells with 2 μM XL413 for 2 h followed by fixa4on and staining\ncells with recombinant RNH1(D210N)-GFP protein. QIBC analysis showed that HRASV12\noverexpression increased R-loops (RNH1 foci), but this was not significantly affected by CDC7\ninhibi4on (please see Fig. R2 below). We also tested the effect of CDC7 inhibi4on on\nreplica4on fork progression in cells overexpressing HRASV12 or cyclin E1. In this experiment,\nXL413 was present during DNA fiber labeling with CldU and IdU. We found that CDC7\ninhibi4on could rescue both HRASV12- and cyclin E1-induced replica4on fork slowing. These\nresults are now shown in Supplementary Fig. 14 and discussed in the manuscript on page 14.\nFigure R2. Top: Experimental workflow. Doxycycline (DOX; 1 μg/ml) was\nadded for the last 48 h to induce HRASV12 expression. Cells were treated\nwith 2 μM CDC7i (XL413) for 2 h. Bo&om: QuanHficaHon of the number of\nRNH1(D210N)-GFP-Tat11 foci per PCNA-posiHve nucleus for the indicated\ncondiHons (n≥401). Black horizontal lines indicate the mean; p-values were\ncalculated by the Kruskal-Wallis test followed by Dunn’s mulHple\ncomparisons test.\nFigure 3:\n- The authors use ZRANB3 deple4on to disrupt fork reversal. It would strengthen the\nconclusions to test whether similar phenotypes are observed upon deple4on of other fork\nreversal enzymes such as HLTF or SMARCAL1?\nResponse: We tested the effect of SMARCAL1 deple4on on replica4on fork slowing and\nmicronuclea4on induced by HRASV12 or cyclin E1 overexpression. We observed similar\nphenotypes as in case of ZRANB3 deple4on. These data are now shown in Supplementary Fig.\n7c, d, j.\n- To further elucidate causal rela4onships, it would be valuable to assess R-loop levels (as\nshown in Figure 2c) following ZRANB3 or other fork reversal enzyme deple4ons. This could\nhelp determine if R-loops persist or resolve under condi4ons of oncogene-induced replica4on\nstress and unrestrained fork progression.\nResponse: We tested the effect of ZRANB3 deple4on on R-loop accumula4on in HRASV12-\noverexpressing U2OS cells. R-loops were detected by staining cells with recombinant\nRNH1(D210N)-GFP protein followed by QIBC analysis. We found that upon ZRANB3 deple4on,\nHRASV12 overexpression did not induce accumula4on of RNH1(D210N)-GFP foci in S-phase\ncells. This is consistent with the model wherein R-loop resolu4on is a prerequisite for\nreplica4on restart via MUS81-LIG4 axis (Rao et al., 2024). The data showing that ZRANB3\n4\ndeple4on suppresses R-loop accumula4on in HRASV12-overexpressing cells are now shown\nin Supplementary Fig. 12d-f.\nFigures 4 and 5:\n- While the MUS81–LIG4 axis appears to play a cri4cal role in promo4ng fork progression\nunder oncogene-induced stress condi4ons, the lack of a comprehensive overview of S-phase\nDNA damage, global DNA synthesis, and the extent of R-loop accumula4on makes it difficult\nto conceptualize the precise nature and extent of fork stalling that necessitates nucleases like\nMUS81.\nResponse: Our aim was to determine whether the MUS81-LIG4-PRIMPOL axis is required for\nunrestrained DNA synthesis induced by inhibi4on of fork reversal in oncogene expressing cells.\nWe believe that the DNA fiber data shown in Fig. 4 and 5 strongly support this claim.\nMeasurement of global DNA synthesis for example by quan4fying EdU incorpora4on cannot\nprovide accurate informa4on about replica4on fork stalling events because it also scores for\ndormant origin firing. We believe that single molecule approaches such as DNA fiber assay are\nmore appropriate to study the processes occurring at stalled replica4on forks. To support our\nclaim that MUS81 is essen4al for restar4ng stalled replica4on forks in HRASV12/cyclin E1-\noverexpressing cells, we reanalyzed DNA fibers in Fig. 4b to determine sister fork ra4o, a\nmeasure of fork stalling frequency, for each condi4on. The obtained data, which are shown\nbelow in Fig. R3, demonstrate that the rescue of sister fork asymmetry (fork stalling) in\nHRASV12/cyclinE1-overexpressing cells by ZRANB3 deple4on depends on MUS81 but not on\nRECQ1.\nFigure R3. MUS81 is required for the rescue of oncogene-induced replicaHon fork stalling by ZRANB3 depleHon.\nLe+: Workflow for DNA fiber labeling in transduced U2OS cells 3 days ajer selecHon. Cells were transfected with\nappropriate siRNAs on day 0. Right: Plot of the values of IdU tract length raHo of sister forks (sister fork raHo) (n\n≥ 100). Black horizontal lines indicate the median; p-values were calculated by the Kruskal-Wallis test followed\nby Dunn’s mulHple comparisons test.\n- It is also puzzling that ZRANB3 knockout cells, which replicate with persistent ssDNA gaps,\nnevertheless show a rescue of mito4c defects typically induced by oncogene-driven\nreplica4on stress. This raises the ques4on of whether these cells rely on daughter strand gap\nrepair mechanisms, such as RAD51-mediated homologous recombina4on. This possibility\ncould be addressed by assessing spontaneous RAD51 foci forma4on during S-phase (e.g., as\ndescribed in DOI: 10.1016/j.devcel.2021.01.011).\noncogene-expressing cells results from incompletely replicated DNA regions caused by\npersistent reversed forks. It is likely that ssDNA gaps generated during PRIMPOL-mediated\nrestart of R-loops-stalled forks in oncogene-expressing cells lacking ZRANB3 are filled by\nhomologous recombina4on before the onset of mitosis. In any case, it is not expected that\npersistence of such ssDNA gaps would impair chromosome segrega4on in mitosis giving rise\nto anaphase bridges and micronuclei. As suggested, we tried to monitor these puta4ve HR\nevents by measuring RAD51 foci during S-phase. However, we found that overexpression of\nHRASV12 or cyclin E1 did not significantly increase the frequency of RAD51 foci in S-phase\ncells even upon deple4on of ZRANB3 (please see Fig. R4 below). Obviously, this does not\nexclude the possibility that the ssDNA gaps are eliminated by HR. It is also possible that the\nssDNA gaps are simply filled in by DNA polymerases as the DNA template does not contain\nobstacles that would block DNA synthesis as in case PRIMPOL-mediate repriming at bulky DNA\nadducts (e.g. cispla4n crosslinks).\nFigure R4. a Top: Experimental\nworkflow. U2OS HRASV12 TetON cells\nwere transfected with siLUC or\nsiZRANB3 three days before fixaHon.\nDoxycycline (DOX; 1 μg/ml) was\nadded for the last 48 h to induce\nHRASV12 expression. Bo&om:\nQuanHficaHon of the number of\nRAD51 foci per PCNA-posiHve\nnucleus for the indicated condiHons\n(n = 760). b Top: Experimental\nworkflow. U2OS CE TetON cells were\ntransfected with siLUC or siZRANB3\nthree days before fixaHon.\nDoxycycline (DOX; 1 μg/ml) was\nadded for the last 48 h to induce\ncyclin E1 (CE) expression. Bo&om:\nQuanHficaHon of the number of RAD51 foci per PCNA posiHve nucleus for the indicated condiHons (n = 640). a,\nb A representaHve plot from two independent experiments yielding similar results is shown. Black horizontal\nlines indicate the median; p-values were calculated by the Kruskal-Wallis test followed by Dunn’s mulHple\ncomparisons test.\nOther points:\n- The authors implicate the poten4al incidence of head-on transcrip4on–replica4on conflicts\nas a driver of R-loop forma4on and replica4on stress. However, this has not been directly\ntested. It would be informa4ve to apply genomics-based approaches, for R-loop mapping in\nrela4on to replica4on origin orienta4on, to explore TRC landscapes in HRASV12 versus Cyclin\nE1-overexpressing cells. Alterna4vely, it should be phrased cau4ously.\nResponse: To obtain evidence that overexpression of HRASV12 or cyclin E1 increases the\nfrequency of head-on TRCs, we measured colocaliza4on between RNAPII and PCNA by\nproximity liga4on assay (PLA). Please note that work in Karlene Cimprich lab has shown that\nhead-on but not co-direc4onal TRCs increase PLA foci between RNAPII and PCNA (Hamperl et\nal., 2017). We found that overexpression of both HRASV12 and cyclin E1 increases the\nfrequency of RNAPII/PCNA PLA foci in U2OS cells, indica4ng head-on TRCs. These data are\n6\nnow shown in Supplementary Fig. 4. Nevertheless, as suggested by the reviewer, we have\ntuned down our statements in the manuscript because the above PLA data do not provide\ndirect evidence for the link between TRCs and R-loop forma4on.\nWe would like to men4on here that we have recently ini4ated experiments to map the\nsites of R-loop forma4on in HRASV12-overexpressing cells by Cut&Tag technology. In the first\nexperiment, we iden4fied 34,607 R-loop peaks for the empty vector (EV) sample and 51,634\nR-loop peaks for the HRASV12 sample. As expected, the most significant peaks of R-loops were\nobserved at promoters/transcrip4on start sites for both condi4ons (47% for EV, 43% for\nHRASV12). However, we also found a large number of intragenic peaks that were enriched in\nthe HRASV12 sample (12043 for EV vs. 19827 for HRASV12). These peaks were predominantly\nunique and did not overlap between the two condi4ons. We also analyzed the loca4ons of R-\nloops rela4ve to the origins of DNA replica4on. To do this, we used publicly available data from\nSNS-seq performed by Guilbaut et al. 2022 (DOI: 10.1093/nar/gkac555). We selected\nreplica4on origins within the genes on the (+) strand. We detected the presence of R-loops\nnot only at the sites of the origins but also approximately 3 to 4 kb upstream of the origins.\nThis phenomenon was par4cularly evident in the regions of common fragile sites (Figure R5a),\nwhich are known to be highly prone to breakage upon HRASV12-induced replica4on stress.\nImportantly, no such peaks were observed downstream of the intragenic origins or at the\nintergenic origins, suppor4ng the no4on that HRASV12-induced R-loops form at sites of head-\non TRCs (Figure R4b). Obviously, these are very preliminary data that need to be confirmed by\nat least one more Cut&Tag experiment. Therefore, we cannot include these data in the current\nmanuscript. These data will be part of a follow-up manuscript, where we also intend to\nperform Cut&Tag experiments to map the loca4on of stalled forks (MCM7, pol epsilon) and\ntranscrip4on complexes (elonga4ng form of RNAPII) not only for HRASV12 but also for cyclin\nE1 overexpression to obtain a complete genome-wide profile of the sites of R-loop-mediated\nfork stalling induced by these oncogenes.\nFigure R5. R-loops form at sites of head-on transcrip>on-replica>on conflicts in HRASV12-overexpressing cells.\n(A) S9.6-CUT&Tag-Seq signal across intragenic origins for forward oriented genes (+ strand) within common\nfragile sites (CFSs). (B) S9.6-CUT&Tag-Seq signal across intergenic origins. (A and B) Yellow line: U2OS cells\noverexpressing HRASV12; Blue line, U2OS cells harboring empty vector; Dark purple line: U2OS cells harboring\nempty vector and treated with RNaseA prior to CUT&Tag assay. The data show that R-loops accumulate on the\nhead-on side of origins in gene bodies\n- It is unclear why 5-FU was used to assess global transcrip4onal ac4vity instead of more\nconven4onal and direct methods such as EU (5-ethynyl uridine) incorpora4on, which is widely\nused for nascent RNA labelling.\noverexpressing cells and obtained essen4ally the same results as with 5-fluorouridine. The\nnew data are shown in Supplementary Fig. 6.\nReviewer #2 (Remarks to the Author)\nOravetzova A. et al. show how an increase of transcrip4onal levels or replica4on origin firing\nmediated by oncogene overexpression are linked to genomic instability associated to R-loop\nforma4on in S phase and fork reversal. They also showed that inhibi4on of fork reversal\npromotes fork progression re-start and suppression of mito4c DNA damage mediated by\nMUS81 and PRIMPOL repriming. Reinforcing in the idea that fork reversal have both a\nprotec4ve but also a damaging role if not resolved properly. The manuscript presents a huge\namount of work. While the results presented are solid and it is an interes4ng observa4on, I\ndo have some concerns regarding the novelty of the presented data.\nResponse: We are pleased that the reviewer found our study interes4ng. We believe that the\nnew findings presented in the revised manuscript, par4cularly those providing the\nmechanis4c insights into the role of ROS in HRASV12-induced replica4on stress, will allay the\nreviewer's concerns about the novelty of the data (for further details, please refer to our\nresponse to Reviewer 1 on page 2).\nMajor comments\nThe authors propose that oncogene overexpression (OE) leads to R-loop forma4on specifically\nin S phase, possibly due to TRCs, and inducing persistent replica4on fork reversal. The RNH1\nfoci seen in G2 phase in Fig2C can represent persistent R-loops formed in S phase as the author\nsuggested but also, R-loop formed in G2 if late replica4on and TRCs occurs in G2/M phases.\nLate replica4on has been shown to occur at TSS regions that is where more R-loops are formed\n(Wang J. et al Cell Report 2021). This possibility could be tested by for example checking RNH1\nfoci when inhibi4ng transcrip4on during S phase and reac4va4ng it during G2, or inhibi4ng\nfork reversal resolu4on and checking whether the frequency increases From S to G2.\nResponse: Wang et al. (2021) show that a subset of transcrip4on start sites (TSSs) are\nreplicated in G2/M due to high levels of an4sense transcrip4on, making them difficult to\nduplicate during S phase. It is possible that some R-loops may result from TRCs at sites of late\nreplica4on in G2. However, it is likely that these head-on TRCs occur already in S-phase and\nprevent replica4on due to R-loop forma4on. To monitor head on TRCs in HRASV12/cyclin E1-\noverexpressing cells, we performed PLA between elonga4ng form of RNAPII and PCNA\n(Hamperl et al., 2017). By QIBC analysis, we found that overexpression of either oncogene\nincreased RNAPII/PCNA PLA foci in S-phase cells but not in G2 cells, sugges4ng that RNH1 foci\nseen in G2 cells rather reflect R-loops generated in S-phase. These data are now shown in\nSupplementary Fig. 4 and described on page 7.\nWe are not sure whether the experiments suggested by the reviewer would yield\nconclusive results. It is challenging to establish experimental condi4ons to eliminate\ntranscrip4on or fork reversal selec4vely during the en4re S phase. It is also likely that\nprolonged inhibi4on of transcrip4on would have a nega4ve impact on cell cycle progression.\nthe frequency of EdU-RNAPII or PCNA-RNAPII interac4ons by PLA assays azer oncogene OE.\nResponse: As men4oned above, we found that overexpressed oncogenes increase\nRNAPII/PCNA PLA foci in S-phase, but not G2 cells (Supplementary Fig. 4). This suggests that\nTRCs only occur in S, and not in G2.\nThe authors also propose that the forma4on of fork reversal in HRASV12 OE could be due to\na slow fork progression mediated by ROS. Their data of fork velocity in the presence of RNH1\nOE (Fig1B) support the idea that something else apart from R-loops is delaying the movement\nof the fork. They could check whether TIMELESS/TIPIN is indeed detached from the replisome\nand whether this is also the case of Cyclin E OE.\nResponse: To determine whether HRASV12 overexpression induces dissocia4on of the\nTIMLESS/TIPIN complex from the replisome, we used a PLA-based assay monitoring\ncolocaliza4on between TIMELESS and replica4on sites labeled by EdU incorpora4on into\nnascent DNA. QIBC analysis revealed that HRASV12 overexpression significantly reduced the\nnumber of TIMELESS/EdU PLA foci in nuclei of U2OS cells in a manner dependent on the\nreplisome-associated ROS sensor PRDX2. On the contrary, upon cyclin E1 overexpression, only\na marginal reduc4on in the number of TIMELESS/EdU PLA foci was observed in PRDX2-\nproficient cells, with a further decrease upon PRDX2 deple4on. These data, which are now\nshown in Fig. 8 and Supplementary Fig. 13e, f, support the conclusion that R-loop-mediated\nfork stalling induced by oncogenic RAS is a consequence of ROS-induced dissocia4on of the\nTIMELESS/TIPIN complex from the replisome, which results in reduced fork velocity.\nIs ATR ac4ve in both oncogene OE condi4ons?\nResponse: We observed that overexpression of both oncogenes induced γH2AX forma4on in\nS-phase cells, without causing extensive DNA breakage, which suggests ATR ac4va4on by\nstalled forks (please see Supplementary Fig. 2). Nevertheless, we did not observe a\npronounced increase in the phosphoryla4on of ATR targets such Chk1 Ser317 and Chk1 Ser345\nupon overexpression of HRASV12 or cyclin E1 in U2OS cells (data not shown).\nReviewer #3 (Remarks to the Author)\nIn this manuscript, the authors show that HRAS or cyclin E1 overexpression results in\nenhanced replica4on fork reversal and increased mito4c DNA damage, which is primarily\ncaused by R-loop mediated transcrip4on-replica4on conflicts (TRåCs) during S-phase.\nInhibi4on of replica4on fork reversal creates a condi4on of unrestrained replica4on fork\nprogression which prevents mito4c DNA damage in oncogene overexpressing cells. This\nprocess engages the MUS81/LIG4/EEL axis and PRIMPOL-mediated repriming to support\nunrestrained fork progression, where the presence of persistent reversed forks contributes to\nenhanced fork slowdown/stalling and an increase in mito4c DNA damage. Overall,\nexperimental evidence is solid and well-controlled. Understanding the nature of DNA\nreplica4on stress caused by oncogenes is considered significant. Nevertheless, results of this\nstudy seem incremental; it is uncertain how much new molecular insight can be deduced from\nthe findings presented here. It is already well-established in the field that (1) oncogene\nac4va4on, including Cyclin E, causes TRCs, (2) R-loop forma4on is associated with frequent\n9\nfork reversal, (3) unrestrained fork progression upon PARP inhibi4on is dependent the MUS81\nac4vity and the RECCQ1 helicase, and (4) restart of stalled forks, including R-loops, via MUS81\nrequires repriming mediated by PRIMPOL, all of which are the key takeaways of this study. In\nthis case, overexpression of oncogenes is used to trigger DNA replica4on stress - unless new\nmechanis4c insight of how HRASG12V and Cyclin E uniquely compromises replica4on fork\nstability and causes R-loop is presented, the current study lacks sufficient molecular details\nand is only limited to the phenotypic analyses using loss-of-func4on approaches (e.g., siRNA\nor knockout).\nResponse: We are pleased that the reviewer found our results to be solid and well-controlled.\nWe agree that the impact of our study would be further enhanced by providing insights into\nhow HRASV12 and cyclin E1 induce R-loop-mediated fork slowing. In response, we have\nincorporated data from our follow-up project that specifically addressed this ques4on. We\nkindly ask the reviewer to refer to our response to Reviewer 1, where we summarized our\nfindings (Page 2).\n1. One major concern is that the phenotypic analyses are limited to measuring DNA fiber\ntracks and there is liqle molecular insight to explain mito4c DNA damage. This is a major\nweakness of the manuscript in its current form.\nResponse: We have realized that the term “mito)c DNA damage” has been inappropriately\nused in our manuscript. We show that both oncogenes increase the frequency of anaphase\nbridges and micronuclei in mitosis. It has been demonstrated that these phenotypes, within\nthe context of DNA replica4on stress, are the result of underreplicated DNA regions, which\nimpair chromosome segrega4on (Ying et al., 2013). Our data show that anaphase bridges and\nmicronuclei induced by overexpression of HRASV12 or cyclin E1 are caused by R-loops (rescue\nby RNH1 overexpression) and persistent reversed forks (rescue by ZRANB3/SMARCAL1\ndeple4on). Thus, we believe that our findings provide molecular insight into how the studied\noncogenes impair chromosome segrega4on and thereby induce chromosomal instability. We\nhave modified the manuscript text accordingly.\n2. Figure 1 to establishes R-loop as a source of oncogene-induced replica4on stress is\nperformed only in a cancer cell line (U2OS), which has already undergone a transforma4on\nprocess. On the other hand, Fig. S4 employs immortalized yet non-transformed fibroblast or\nepithelial cells. The authors should at least confirm the key results of Figure 1 using non-\ntransformed cells.\nResponse: We agree with the reviewer that non-transformed cells are more appropriate for\ninves4ga4ng the mechanisms underlying oncogene-induced replica4on stress. To address this\npoint, we u4lized primary BJ fibroblasts with inducible expression of HRASV12 or cyclin E1\n(Maya-Mendoza et al., 2015). Similar to U2OS cells, we observed that replica4on fork stalling\ninduced by HRASV12 or cyclin E1 in these cells depends on RNAPII transcrip4on (rescue by\nRNAPII inhibi4on with triptolide; Supplementary Fig. 11b, c) and is alleviated by PARP1\ninhibi4on in a MUS81-dependent manner (Supplementary Fig. 7g, h, and Supplementary Fig.\n9b, d). Furthermore, in these cells, we could reproduce our new findings that HRASV12,\nbut\nnot cyclin E1, overexpression induces replica4on fork stalling through a mechanism\ndependent on ROS and the replisome-associated ROS sensor PRDX2 (Supplementary Fig.\n11b, c; Supplementary Fig. 12b; Supplemetary Fig. 13c). Consistently, PRDX2 deple4on\naqenuated R-loop forma4on in HRASV12-expressing BJ fibroblasts (Supplementary Fig. 12c).\n10\n3. In Figure 1, it should be tested whether RNaseH1 expression can suppress DNA damage\nmarkers (e.g., gH2AX). In rela4on to this, the nature of DNA lesions (e.g., the status of DSBs)\nhas not been scru4nized. What would be the underlying basis of DNA damage that contributes\nto defec4ve chromosome segrega4on and mito4c DNA damage azer R-loop accumula4on?\nResponse: We tested the effect of HRASV12 or cyclin E1 overexpression on the forma4on of\nγH2AX and 53BP1 foci in different stages of the cells cycle. As men4oned in our response to\nReviewer 1 (page 3), we found that none of the oncogenes tested induce extensive DNA\ndamage in S-phase cells (Supplementary Fig. 2). Therefore, we did not perform the suggested\nRNaseH1 overexpression experiment. Our data suggest that defec4ve chromosome\nsegrega4on (anaphase bridges and micronuclei) in these cells results from persistent reversed\nforks as evidenced by rescue of these phenotypes by ZRANB3/SMARCAL1 deple4on.\n4. The authors uses RNaseH1-dependent rescues of fork progression as sole evidence to\nsupport that R-loops are the cause of oncogene-induced replica4on stress. However, there is\nno direct evidence of R-loop forma4on in these oncogene overexpressing cells. This could be\nvalidated by using the S9.6 an4body previously used by the lab in Ref#25 either through\nimmunofluorescence or DRIP-seq. A posi4ve control such as PDS or even HU or APH can be\nincluded as men4oned by the authors.\nResponse: To confirm that overexpression HRASV12 or cyclin E1 induces accumula4on of R-\nloops in U2OS cells, we isolated genomic DNA from these cells and performed slot blot using\nS9.6 an4body, which specifically binds to RNA:DNA hybrids. We observed a significantly higher\nR-loop signal in samples from oncogene-expressing cells compared to that of cells harboring\nempty vector. Trea4ng the genomic DNA samples with RNase H before slot blot analysis\ndiminished the S9.6 signals, confirming the specificity of the R-loop signals detected with the\nS9.6 an4body. These data are now shown in Supplementary Fig. 5 and described on page 7.\n5. 5-FU staining suggests that there is an increase in transcrip4onal ac4vity in G1, S, and G2/M\ncells that are overexpressing HRASG12V but not in cyclin E1 overexpressing cells, while the\nauthors claim that R-loops seen in S-phase are carried over into mitosis. Can the authors\nexclude the possibility of R-loop forma4on within G2/M itself independently of DNA\nreplica4on?\nResponse: We consider it unlikely that R-loops detected in G2/M cells were formed\nindependently of DNA replica4on. If these R-loops resulted from increased transcrip4on, we\nwould expect to see R-loop accumula4on in G1 cells as well. However, this is not the case in\nHRASV12-expressing cells (Fig. 2c). As men4oned in our response to Reviewer 2 (point #1,\npage 8), in addi4on to R-loops, we monitored head-on TRCs by measuring colocaliza4on\nbetween elonga4ng form of RNAPII and PCNA using PLA. QIBC analysis revealed that\noverexpression of HRASV12 or cyclin E1 increased RNAPII/PCNA PLA foci in S-phase cells but\nnot in G2 cells (Supplementary Fig. 4; page 7 of the manuscript). These data provide further\nsupport for our hypothesis that R-loop forma4on in HRASV12-/cyclin E1-overexpressing cells\nis driven by head-on TRCs during S-phase and that a frac4on of these R-loops can persist into\nG2/M, thereby blocking comple4on of DNA replica4on.\n6. The authors show that ZRANB3 completely prevents oncogene-induced anaphase bridges\nand micronuclei (Fig. 3). It is an interes4ng result since other translocases such as SMARCAL1\nor HLTF are also known to be involved in fork reversal. These factors should be tested to\ndetermine how much individual translocase contributes to the fork reversal process. If\nsolely\n11\ndependent on ZRANB3, some mechanis4c explana4on should be presented considering the\nway ZRANB3 acts to promote fork reversal.\nResponse: We tested the effect of SMARCAL1 deple4on on replica4on fork stalling and\nmicronuclea4on induced by HRASV12 or cyclin E1 overexpression in U2OS cells. We found\nthat SMARCAL1 deple4on restored normal fork progression and prevented micronuclea4on\nin these cells, as did ZRANB3 deple4on. These data are now shown in Supplementary Fig. 7c,\nd, j.\n7. It would be important to test whether PRIMPOL’s primase ac4vity is actually required for\nunrestrained replica4on fork progression.\nResponse: We agree that this is an interes4ng ques4on, but it should be noted that it was not\naddressed in previous studies on PRIMPOL-mediated repriming (Bai et al., 2020; Quinet et al.,\n2020; Tirman et al., 2021). All of these studies only used PRIMPOL deple4on to assess its role\nin repriming. Evidence that PRIMPOL-mediated repriming is involved in unrestrained fork\nprogression is provided by S1 nuclease assay demonstra4ng the presence of ssDNA gaps on\nDNA fibers. This assay was also used in our study (Fig. 5b). PRIMPOL mediates repriming\nthrough its primase and polymerase ac4vi4es. Although PRIMPOL uses the same ac4ve site\nfor both ac4vi4es, the cataly4c core alone is sufficient for polymerase ac4vity but not primase\nac4vity. Primase ac4vity requires a zinc finger module located C-terminal to the cataly4c core\n(residues ~372 to 487), which has been shown to be capable of binding to ssDNA (Rechkoblit et\nal., 2016). Therefore, one could mutate this zinc finger module to generate a PRIMPOL\nmutant that is defec4ve in primase ac4vity, yet s4ll proficient in polymerase ac4vity. However,\nto our knowledge, such mutants have not yet been characterized in vitro. Therefore,\ninves4ga4ng the role of this region of PRIMPOL in repriming would require extensive research,\nincluding biochemical characteriza4on of the mutants, genera4on of stable cell lines\nexpressing PRIMPOL mutants, and phenotypic analyses. This is beyond the scope of our study.\nNevertheless, we firmly believe that the data from our S1 nuclease/DNA fiber assays strongly\nsupport our conclusion that unrestrained DNA synthesis induced by fork reversal deficiency in\nHRASV12-/cyclin E1-overexpressig cell involves PRIMPOL-mediated repriming (demonstra4on\nof ssDNA gaps).\n8. Figure 6 model – there is no experimental evidence suppor4ng that ZRANB3 deficiency\nrescues the mito4c defect. In rela4on to this, the physiological relevance of inhibiting fork\nreversal is not clear. The phenotype of unrestrained DNA synthesis is revealed when cells are\nforced to lose fork reversal process, and whether unrestrained DNA synthesis plays a\nmeaningful role in normal DNA replica4on and mitosis is not addressed. Addi4onally, there is\nno mechanism of fork remodeling and processing that is uniquely engaged by oncogenes.\nResponse: Please note that we demonstrate that ZRANB3 deple4on prevents the forma4on\nof anaphase bridges and micronuclei in HRASV12- or cyclin E1-overexpressing U2OS cell\n(mito4c defects). As men4oned above these phenotypes are caused by incompletely\nreplicated DNA regions, which impair chromosome segrega4on in mitosis. If cells do not have\nZRANB3, forks stalled at R-loops are restarted during S-phase, which allows for normal\nchromosome segrega4on in mitosis and hence prevents chromosomal instability. We have\npreviously shown that in fork reversal-proficient cells, there is an equilibrium between fork\nreversal and fork restart, which is regulated by RECQ1 and RECQ5 helicases, and that the\nmajority of forks stalled by an R-loop are restarted prior to onset of mitosis (Chappidi et al.,\n2020). Chromosome segrega4on errors (anaphase bridges and micronuclei) likely result from\n12\nreplica4on fork stalling events in late S-phase, when there is not enough 4me to restart\nreplica4on before mitosis. 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Cell Biol. 15, 1001–1007.\n14"}, {"round": 2, "reviewer_comments": {"reviewer_1": "The revised manuscript by Oravetzova, Dvorakova et al. includes substantial additional work and improved clarity compared to the initial submission. As noted previously, the study is technically well executed, supported by complementary experimental approaches and appropriate statistical analyses. In the original version, however, the novel mechanistic insights were somewhat limited.\nIn the revised manuscript, the authors now more effectively develop the concept of mechanistic convergence in oncogene-induced replication stress, focusing on HRASV12 and Cyclin E1. In particular, the clarification that the ROS–PRDX2 axis acts as an upstream driver of replication fork slowing, and that it contributes to the formation of transcription–replication conflicts (TRCs) in HRASV12 but not in Cyclin E1, strengthens the overall contribution of the study. The additional data, including the use of NOX inhibitors, ROS quenching, and PRDX2 depletion experiments, provide useful insight into how distinct oncogenic programs can lead to overlapping replication stress outcomes.\nThe authors have adequately addressed the specific comments raised during revision, and the revised manuscript presents a large amount of data more systematically and coherently, improving its overall impact for understanding how oncogenes shape replication stress and genome stability. One minor comment from my side is that the authors should consider using a more specific title that better reflects the key data and conclusions presented in the study, rather than relying on a broad and generic title.", "reviewer_2": "I appreciate the authors’ substantial effort to improve the quality and novelty of the manuscript. The revised version is clearly stronger than the original one, and the inclusion of additional mechanistic data regarding the role of ROS and PRDX2 in the replication stress induced by HRASV12 has significantly improved the novelty of the study. Nevertheless, I still have some concerns that should be addressed.\nThe authors conducted new PLA experiments to obtain evidence that overexpression of HRASV12 or cyclin E1 increases the frequency of head-on TRCs. Although they acknowledge that PLA data do not provide direct evidence for the link between TRCs and R-loop, and they have turn down the statements, they claimed that the increase of PLA foci when overexpressing HRASV12 is linked to HO TRCs. They based their conclusions in the observation made by Hamperl et al., 2017 where head-on but not co-directional TRCs increase PLA foci between RNAPII and PCNA. However, these results have been obtained from artificial episomal systems. A recent study has demonstrated that co-directional TRCs, where the RNAPII is which collides with the replication fork traveling ahead, also exists (Bruno et al Mol Cell 2024; doi:10.1016/j.molcel.2023.11.036). These new type of TRCs that should not block replication fork progression can also be detected by these PLA experiments. Consequently, this possibility should be discussed, and it should be acknowledged that at least a fraction of the increase in PLA foci may not correspond to head-on (HO) TRCs. This is especially relevant in the context of HRASV12 overexpression that leads to an increase in transcriptional activity.\nMoreover, to clearly determine whether HO TRCs are responsible for the observed genomic and mitotic instability, the manuscript would greatly benefit from the inclusion of the HRASV12 overexpression CUT&Tag genomic data the authors are generating and the estimation of replication fork directionality that can be done using public U2OS Repli-seq data. Alternative, they should be more cautious when assigning directionality to TRCs.", "reviewer_3": "The authors addressed many concerns raised in the previous version and strengthened the manuscript. The revised title does not summarize their original findings, though. It is recommended to come up with a more specific title."}, "author_response": "Point-by-point response to the reviewers' comments (NCOMMS-25-21283A)\nWe would like to thank the reviewers for taking the 4me to evaluate the revised version of\nour manuscript and for their posi4ve feedback. We are delighted that all reviewers found\nthe manuscript to be substan4ally improved.\nAnswers to specific ques4ons of reviewers:\nReviewer #1 (Remarks to the Author):\nThe revised manuscript by Oravetzova, Dvorakova et al. includes substan4al addi4onal work\nand improved clarity compared to the ini4al submission. As noted previously, the study is\ntechnically well executed, supported by complementary experimental approaches and\nappropriate sta4s4cal analyses. In the original version, however, the novel mechanis4c\ninsights were somewhat limited.\nIn the revised manuscript, the authors now more effec4vely develop the concept of\nmechanis4c convergence in oncogene-induced replica4on stress, focusing on HRASV12 and\nCyclin E1. In par4cular, the clarifica4on that the ROS–PRDX2 axis acts as an upstream driver\nof replica4on fork slowing, and that it contributes to the forma4on of transcrip4on–\nreplica4on conflicts (TRCs) in HRASV12 but not in Cyclin E1, strengthens the overall\ncontribu4on of the study. The addi4onal data, including the use of NOX inhibitors, ROS\nquenching, and PRDX2 deple4on experiments, provide useful insight into how dis4nct\noncogenic programs can lead to overlapping replica4on stress outcomes.\nThe authors have adequately addressed the specific comments raised during revision, and\nthe revised manuscript presents a large amount of data more systema4cally and coherently,\nimproving its overall impact for understanding how oncogenes shape replica4on stress and\ngenome stability. One minor comment from my side is that the authors should consider\nusing a more specific 4tle that beXer reflects the key data and conclusions presented in the\nstudy, rather than relying on a broad and generic 4tle.\nResponse: We thank Reviewer 1 for the thougthful assessment of our revised manuscript.\nWe agree that the previous 4tle was somewhat broad, and we have revised it to beXer\nreflect the key findings of our study. The new 4tle “Dis$nct mechanisms of replica$on stress\ninduced by oncogenic RAS and cyclin E1 converge on R-loop-dependent fork reversal” more\nspecifically captures the main conclusions, highligh4ng the dis4nct oncogene-induced\nreplica4on stress mechanisms and the common downstream outcome involving R-loop-\nmediated fork reversal.\nReviewer #2 (Remarks to the Author):\nI appreciate the authors’ substan4al effort to improve the quality and novelty of the\nmanuscript. The revised version is clearly stronger than the original one, and the inclusion of\naddi4onal mechanis4c data regarding the role of ROS and PRDX2 in the replica4on stress\n1\ninduced by HRASV12 has significantly improved the novelty of the study. Nevertheless, I s4ll\nhave some concerns that should be addressed.\nResponse: We thank Reviewer 2 for the careful evalua4on of our revised manuscript and for\nacknowledging the substan4al improvements in quality and novelty compared to the\noriginal submission. We are pleased that the reviewer finds the addi4onal data on the role\nof ROS and PRDX2 in HRASV12-induced replica4on stress to significantly strengthen the\nstudy. We also appreciate the reviewer’s con4nued engagement and construc4ve feedback.\nThe remaining concerns are addressed below.\nThe authors conducted new PLA experiments to obtain evidence that overexpression of\nHRASV12 or cyclin E1 increases the frequency of head-on TRCs. Although they acknowledge\nthat PLA data do not provide direct evidence for the link between TRCs and R-loop, and they\nhave turn down the statements, they claimed that the increase of PLA foci when\noverexpressing HRASV12 is linked to HO TRCs. They based their conclusions in the\nobserva4on made by Hamperl et al., 2017 where head-on but not co-direc4onal TRCs\nincrease PLA foci between RNAPII and PCNA. However, these results have been obtained\nfrom ar4ficial episomal systems. A recent study has demonstrated that co-direc4onal TRCs,\nwhere the RNAPII is which collides with the replica4on fork traveling ahead, also exists\n(Bruno et al Mol Cell 2024; doi:10.1016/j.molcel.2023.11.036). These new type of TRCs that\nshould not block replica4on fork progression can also be detected by these PLA\nexperiments. Consequently, this possibility should be discussed, and it should be\nacknowledged that at least a frac4on of the increase in PLA foci may not correspond to\nhead-on (HO) TRCs. This is especially relevant in the context of HRASV12 overexpression that\nleads to an increase in transcrip4onal ac4vity.\nResponse: As recommended by the reviewer, we now discuss this possibility in the\nmanuscript. The following sentence has been added to page 7: “However, we cannot exclude\nthe possibility that some of these PLA foci represent co-directional RNAPII collisions behind\nthe replication fork, potentially arising from oncogene-induced fork slowing”, ci4ng the\nar4cle by Bruno et al. (2024). Indeed, co-direc4onal transcrip4on resuming on the leading\narm of a slowed replica4on fork (e.g., following ROS-induced TIMLESS-TIPIN dissocia4on)\ncould bring the RNAPII complex into close proximity with the PCNA clamp and thereby\ngenerate a posi4ve signal in RNAPII/PCNA PLA assay.\nMoreover, to clearly determine whether HO TRCs are responsible for the observed genomic\nand mito4c instability, the manuscript would greatly benefit from the inclusion of the\nHRASV12 overexpression CUT&Tag genomic data the authors are genera4ng and the\nes4ma4on of replica4on fork direc4onality that can be done using public U2OS Repli-seq\ndata. Alterna4ve, they should be more cau4ous when assigning direc4onality to TRCs.\nResponse: We prefer not to include CUT&Tag data in the current manuscript. To date, we\nhave performed only a single CUT&Tag experiment to map the sites of R-loop forma4on in\ncells expressing HRASV12. To draw any conclusions, at least one addi4onal biological\nreplicate would be required, and the analysis would need to be extended to cyclin E1-\noverexpessing cells. We hope the reviever will agree that comple4ng this work would\nrequire several addi4onal months. While we do plan to perform these experiments, we\nprefer to include them in a future study, in which we also aim to perform CUT&Tag\n2\nexperiments to map the loca4on of stalled forks (MCM7, pol epsilon) and transcrip4on\ncomplexes (elonga4ng form of RNAPII) in both HRASV12- and cyclin E1-overexpressing cells.\nWe believe that analysis of these datasets will ul4mately allow to draw more robust\nconclusions regarding TRC direc4onally, R-loop forma4on and fork stalling upon oncogene\nac4va4on.\nWe would like to emphasize that the current manuscript provides experimental\nevidence demonstra4ng that oncogene-induced fork stalling and chromosomal instability\n(anaphase bridges and micronuclei) are caused by R-loops (rescue by RNH1 overexpression).\nOur hypothesis that these R-loops may result from head-on TRCs is based on previously\npublished findings (e.g., doi: 10.1016/j.cell.2017.07.043) and is not presented as a main\nconclusion of the manuscript. To avoid any such impression, we have removed the\nstatement “…, likely as a consequence of head-on transcrip4on-replica4on conflicts (TRCs)”\nfrom the abstract.\nReviewer #3 (Remarks to the Author):\nThe revised 4tle does not summarize their original findings, though. It is\nrecommended to come up with a more specific 4tle.\nResponse: We appreciate the reviewer’s feedback and are grateful for his/her recogni4on of\nthe improvements made to the manuscript. We understand the concern regarding the\nrevised 4tle and agree that it should more accurately summarize our key findings. To address\nthis, we have modified the 4tle to: “Dis$nct mechanisms of replica$on stress induced by\noncogenic RAS and cyclin E1 converge on R-loop-dependent fork reversal”. We believe that\nthis revised 4tle more precisely reflects the core aspects of our study.\n3"}]

Decision letter