RF-SIRF reveals a replication stress-specific epigenetic code by spatio-temporal mapping of reversed forks

Domain: biology
Published: 2026-03-23
Editorial outcome: major_revision

Abstract
DNA replication stress responses are guardians of genomiEc stability critical during development, hematopoiesis, cancer therapy response, aging and disease suppression. Central to these responses are reversed forks (RF), which are distinct four-way DNA structures formed during DNA replication stalling to protect against toxic DNA lesions. Historically, RF detection relies on specialized electron microscopy, precluding studies within their native cellular context. By harnessing intrinsic bio-physical properties of RFs, we here present a quantitative method to map RFs with single-cell resolution (RF-SIRFI). RF-SIRF reveals that RFs accumulate at the nuclear periphery during early-mid S-phase of the cell cycle. Crucially, RFs possess a specialized chromatin landscape and utilize an epigenetic replication stress code distinct from transcription, explaining the selective recruitment of DNA stress response proteins to RFs. Collectively, RF-SIRF enables robust quantitative, temporal, spatial and proteomic analyses of reversed forks, empowering advIanced cellular and medical investigations of DNA replication stress responses.

Introduction
Growing evidence establishes fork reversal (RF) as a pivotal mechanism across diverse biological contexts, including aging, hematopoiesis, and cellular senescence, where RFs dictate stem cell maintenance and therapeutic vulnerability in cancer. RF are formed during stress conditions as a protective measure that promotes DNA damage tolerance in actively replicating cells to avoid exposure to DNA lesions and formation of toxic DNA double strand breaks 1-4. Thus, RF are of particular importance in highly proliferating cells, those exposed to DNA damage and those that are under metabolic stress. RFs accumulate in both aged and embryonic stem cells 5, 6, are needed to suppress myeloid and lymphoid dysregulation in both long-term hematopoietic stem cells as well as progenitor cells 6, and form at telomeres as implied in preventing replicative senescence 7, 8. On the flip side, RFs also can be detrimental within certain genetic backgrounds leading to telomere catastrophe as well as genome-wide instability such as in BRCA1/2 fork protection defective cancer cells, where formation of RFs controls cellular sensitivity and resistance to standard of care and PARP inhibitor cancer therapies 9-17.

Fork reversal involves the annealing of newly synthesized DNA strands, converting a three- way replication fork into a four-way junction and so creating a regressed double-stranded DNA arm. In vitro, positive super-coiling of DNA provides sufficient free energy to achieve fork reversal 17. In cells, fork reversal is achieved by designated translocases including SMARCAL1, HLTF and ZRAN3B, with added DNA remodeling proteins FBH1, FANCM helicases, RAD51 recombinase and RAD51 paralogs being also implied in the process 18-24. SMARCAL1 holds a biochemically distinct role in annealing RPA coated stalled leading strands and displays a somewhat dominant function under several cellular contexts including in BRCA2 defective cells and at telomeres 12-15, 20, 25-28. The recent discovery of SMARCAL1 status controlling inflammation and its use as a biomarker for immune checkpoint therapy response corroborates this notion and further emphasizes the importance of RF reactions in diverse cancer cell therapy settings 26.

Despite their fundamental role in health and disease, the study of reversed forks in their native cellular context has been hindered by a lack of robust, highS-resolution tools. Surrogate RF assays that indirectly measure RF include DNA fiber analySsis, 2D-gel electrophoreses and immunofluorescence approaches 1, 2, 22. Yet, these are limited in efficiency, sensitivity, resolution, specificity, or quantitative robustness, restricting their broad usage. To date, the prevailing accepted assay to measure RFs with sufficient resolution and robustness is by electron microscopy (EM) 1, 2, 22, 29. EM visualizes RFP structures, provides quantitative information regarding the relative abundance of RF and allows for single-molecule size measurements. However, this in vitro method is technically challenging requiring sensitive processing steps, biophysics expertise and sIpecialized instrumentation, making the assay a time-consuming and costly effort that is prohibitive for most laboratories. Moreover, it is a bulk- DNA technique that lacks single-cell resolution and spatial context about the cellular macro- environment that defines fork reversal.

Given the persistent challenIges to examining RFs, there is a critical need for alternative, sensitive, and reproducible methods to monitor these structures in their native cell environment. Here, we directly harnessR the unique geometry of the reversed DNA fork structure, which enforces the proximityA of nascently replicated leading and lagging strand to develop a robust reporter to examine fork reversal by RF-SIRF. RF-SIRF enables the quantitative, sensitive, and reproducible analysis of reversed forks within single-cells.

Results
RF-SIRF signals are elevated under conditions permissive for fork reversal

We aspired to devise an efficient, robust and technically amicable assay system to detect reversed DNA replication forks by considering the bio-physical aspects of these DNA structures. In a canonical Y-form DNA replication fork, the newly synthesized strands on the leading and lagging strand templates are spatially separated (Fig. 1a, top panel, red dashed lines). In contrast, reversed forks form a four-way DNA junction containing a regressed arm, in which the nascently replicated leading and lagging DNA strands are annealed, thus are in direct proximity to each other (Fig. 1a, bottom panel, red bracket).

Proximity ligation assay (PLA) exploits the spatial distance of two antibody-directed epitopes to yield a fluorescent signal only if the two epitopes are within close proximity (nominally <40nm) (Fig. 1a) 30-32. Based on the unique structural elements contained within reversed fork, we reasoned that the distance between the newly incorporated Ethynyl-2'-deoxyuridine (EdU) residues is reduced in the regressed arm of the reversed fork (Fig. 1a, red-green bubble, bottom panel), compared to a canonical Y-shaped structure at non-stressed forks (Fig. 1a, red-green bubble, top panel). We therefore hypothesized that reversed replication forks will produce a greater EdU-PLA signal compared to Y-forks (Fig. 1a). This hypothesis builds upon and extends our previous reports on SIRF, which is an in situ assay to detect proteins at nascent replication forks using a modified PLA approach 33-36. For the development of SIRF, in which we biotinylate EdU via Click-it chemistry, we showed a direct correlation between the proximity of biotin-EdU moieties on nascently replicated DNA and the productivity of a PLA signals when using primary antibodies against biotin-EdU 36. Importantly, protein-SIRF signals are compared and normalized to the total amount of incorporated EdU to differentiate between signal changes caused by changes in EdU substrates (e.g. amounts of nascent DNA) from those caused by an actual loss of proximity (e.g. loss of protein-DNA interaction), distinguishing it from PLA. To achieve this, we perform simultaneous Click-it chemistry with biotin-azide and Alexa488-azide so that both biotin and Alexa488 directly correlate to the total amount of EdU moieties present. Given that the Click-it reaction is a covalent chemical reaction thaSt depends on substrate availability but not proximity, Alexa488 serves as a direct meaSsure and normalization control for the total amount of incorporated EdU. In contrast, EdU-PLA signals using anti-biotin antibodies depend on the proximity between the nascently incorporated EdU moieties, which we suggest will be greater at RFs compared to ongoing Y-forks (Fig. 1a).

To test our RF hypothesis, we performed a rev ersed fork (RF)-SIRF whereby PLA against biotinylated EdU is performed and compared to the total amount of EdU as measured by Alexa488-co-Click-it (Fig.1a). To devise a robIust and specific assay system we considered three distinct features. First, for RF-SIRF we used cells and replication stalling conditions extensively validated to promote fork reversal 12, 15, 22, 37, 38. Second, EdU is incorporated within a short 8’ pulse only prior to exposure to the reversal-inducing DNA replication stress. This ensures that a change in nascent DNA signal reflects fork regression of previously incorporated EdU molecules prior to reversalI. Third, EdU-Alexa488 is used to normalize PLA signals to total amounts of incorporated EdU with single-cell resolution. This enables to determine if a loss or gain of EdU-PLA signal is due to EdU proximity or a loss or gain in total amounts of EdU- substrate. The experimAental set up of RF-SIRF thus predicts the anti-biotin EdU-PLA signal to increase with fork reversal. In contrast, the EdU-Alexa488 signal is predicted to remain unchanged regardless of fork reversal (Fig. 1a).

Low dose exposure to camptothecin (CPT, 50nM) for one hour substantially induces fork reversal and concomitantly slows replication speed, without evoking double strand breaks (DSB) as extensively validated in U2OS cells by electron microscopy (EM) and single-molecule DNA fiber assay 1-3, 39-41. We performed RF-SIRF under these proven fork reversal conditions (Fig. 1a). Using anti-biotin antibodies for the EdU-PLA reaction, we observed a substantial increase in EdU-PLA signals in U2OS cells challenged with CPT compared to medium-treated cells (Fig. 1b,c; median EdU-PLA intensity of 0.584 with compared to 0.432 without CPT, Supplementary Fig. 1a). PLA-control reactions without EdU showed no appreciable PLA signal (Fig. 1b). In stark contrast to EdU-PLA, the EdU-Alexa488 signals decreased with CPT to some extent (Fig. 1b,d; median Alexa488 intensity of 0.21 with compared to 0.294 without CPT, Supplementary Fig. 1b). While in an ideal experimental set-up we expect EdU-Alexa488 signals to remain unchanged since EdU is incorporated prior to replication stalling, we considered that there could be residual EdU incorporation even after EdU wash-out as previously described42, and this may explain the reduced EdU-Alexa488 signals with CPT given the reported slowing of replication under these conditions39. Consistently, EdU-Alexa488 signals intensities increase over time despite EdU wash-out (Supplementary Fig.1d). To account for EdU-substrate changes with CPT treatment, we normalized the EdU-PLA signals to the EdU-Alexa488 signals on a single-cell level for an objective and robust SIRF measurement for reversed forks (RF- SIRF), which shows a significant increase in RF-SIRF signals under conditions that promote reversed forks (Fig. 1b,e; median RF-SIRF of 0.514 with CPT compared to 0.360 without, Supplementary Fig. 1c). Importantly, the data shows that the increase in RF-PLA signals seen with CPT is not caused by an inadvertent increase in EdU incorporation with CPT post-stalling. Instead, the greater EdU-PLA signals evidently are caused by a closer proximity of the previously incorporated EdU moieties. This increase is optimal with higher EdU concentrations that can maximize proximity between EdU moieties and so maximizes EdU-PLA signals under fork reversal conditions (Supplementary Figs. 1e-g) 36. Further validating this, increasingly delaying CPT-mediated fork reversal, which allows the fork to move away from the EdU-pulse label, decreases RF-EdU-PLA intensities, confirming that RF-signals are dependent on nascent EdU incorporated prior to CPT-stalling (Supplementary Figs 1h-j).

Fork reversal swiftly occurs upon exposure to diverse genotoxic agents

Aside from CPT, reversed replication forks are significantly indSuced with low doses of diverse genotoxic reagents 22. To test if the increase in RF-SIRF signal represents reversed forks under conditions other than CPT, we exposed U2OS cells to one hour hydroxyurea (HU, 500nM), aphidicolin (APH, 100 nM), or hydrogen peroxide (H2O2, 20 M) as previously reported permissive for fork reversal 22 (Supplementary Fig. 2a-l). When considering EdU-PLA only, H2O2 causes a significant increase in signals. Bot h APH and HU also show increased EdU-PLA signals, albeit without statistical significance (Supplementary Fig. 2a; median EdU-PLA intensities of 0.276 with HU, 0.292 with APH,I 0.379 with H2O2 and 0.255 without external replication stress). Similar to CPT conditions, there was a reduction in EdU-substrate in one- hour treated compared to untreated cells (Supplementary Fig. 2b; median EdU-Alexa488 intensities of 0.143 with HU, 0.187 with APH, 0.227 with H2O2 and 0.419 without external replication stress). However, when calculating RF-SIRF, which takes into account the total available EdU substrate per cellI, the RF-SIRF signal are significantly elevated under all RF- promoting conditions, corroborating the importance of normalization for EdU substrate availability (SupplementarRy Fig. 2c; median RF-SIRF of 0.342 with HU, 0.347 with APH, 0.413 with H2O2 and 0.214 Awithout external replication stress).

To better control for EdU substrate changes, we tested if shorter exposures to CPT are sufficient to observe fork reversal by RF-SIRF. In contrast to the longer one-hour incubations with CPT, there is only a small and non-significant change in the EdU-Alexa488 signals with or without 15 minutes of CPT treatments (Figure 2a; median EdU-Alexa488 intensities of 0.293 with CPT and 0.32 without). Yet, the EdU-PLA signal intensity significantly increases under these conditions, resulting in robust RF-SIRF signals that are significantly higher when cells are challenged with CPT, thus under conditions promoting fork reversal (Fig. 2b,c; median EdU-PLA intensities of 0.432 with CPT 0.173 without, Supplementary Fig. 2m-o). Similarly, there are smaller differences in EdU-Alexa488 signals with or without 15 minutes of HU, APH or H2O2 compared to the one-hour exposures (Fig. 2d; median EdU-488 intensities of 0.255 with HU, 0.211 with APH, 0.321 with H2O2 and 0.355 without external replication stress). In contrast to the Alexa488 intensities, the EdU-PLA signals substantially increase with all agents tested, resulting in robust RF-SIRF signals upon exposure to the various DNA replication stresses (Fig. 2 e,f; median EdU-PLA intensities of 0.265 with HU, 0.422 with APH, 0.477 with H2O2, 0.166 without external replication stress, and median RF-SIRF of 0.252 with HU, 0.4 with APH, 0.413 with H2O2 and 0.14 without external replication stress, Supplementary Fig. 2p-x).

Fork reversal is not restricted to low damage conditions and also occurs with high concentrations of HU13. Consistently, EdU-PLA but not EdU-Alexa488 and consequently RF- SIRF signals significantly increase after exposure to high concentrations of hydroxyurea (Fig. 2g-i; median Alexa488-PLA intensities of 0.105 with 4mM HU and 0.145 without, EdU-PLA intensities of 0.010 with HU and 0.006 without, RF-PLA intensities of 0.134 with HU and 0.089 without). Of note, the short damage exposure conditions limit loss of nascently DNA incorporated prior to stalling as seen by DNA fiber analysis (Fig. 2j, median IdU tract length of 9.03m without, 9.63 m with 0.5mM HU, 10.0 m with CPT and 9.18 m with 4mM HU, respectively). In contrast, residual nascent DNA replication post-damage exposure varies depending on the stalling agents and concentration used (Supplementary Fig. 2y). Under any circumstance, it is decreased with any damage compared to no treatments, further corroborating that inadvertent residual post-damage incorporation of EdU is unlikely to be a primary cause for the observed increase in RF-SIRF signals.

Taken together, the data show that RF-SIRF signals increase within short, fifteen-minute exposures to diverse genotoxic replication stalling agents known to reverse DNA replication forks, whereby oxidative damage apparently promotes the most robust RF-SIRF signals amongst the agents tested.

Single-stranded (ss)DNA-SIRF signals are elevated under conditions that promote fork reversal

In principle, increased EdU-PLA signals could arise from proximity of nascent EdU at template switched forks as an alternative to one-ended reversed forks 43. To further validate that RF-SIRF preferentially detects reversed replication forIks, we again took advantage of the biophysical features of reversed forks. Intrinsic and unique to reversed forks, the regressed arms of reversed forks contain short nascent ssDNA overhangs 22, 29, 40, 44 (Figure 3a, schematic to the right, dashed box), in contrast to nascently replicated DNA at an ongoing replication fork, which is paired with its template (Figure 3a, schematic to the left), or nascent template switched strands that anneal to complemIentary nascent strands without DNA ends (Supplementary Fig. 3a). We therefore hypothesized that SIRF signals using an antibody with specificity against ssDNA 45-47 will increase uRnder conditions favoring reversed forks. Consistent with this notion, ssDNA-SIRF signals sAignificantly increased with one hour CPT treatments (Fig. 3b, median ssDNA-SIRF of 0.476 with CPT and 0.245 without, and Supplementary Fig. 3b-f). Thus, nascent ssDNA, which is limited at ongoing and template-switched replication forks but known to form at reversed forks 22, 29, 40, 44, is elevated under conditions that promote RF-SIRF, further supporting RF-SIRF as a measure of reversed forks.

Exonuclease T (ExoT) is a 3’-5’ and RecJ is a 5’-3’ structure specific DNA exonuclease, respectively 48 49. Both act on ssDNA but cannot initiate from or extend resection into duplex DNA. Similar to CPT, H2O2 strongly increases ssDNA-SIRF signals (Fig. 3d,e median ssDNA-SIRF of 0.08 with H2O2 compared to with 0.01 without, and Supplementary Fig. 3g,h). Treatment of cells with either RecJ or ExoT subsequent to fork stalling by H2O2 significantly decreased ssDNA-SIRF signals (Fig. 3d,e median ssDNA-SIRF of 0.08 with H2O2 compared to 0.015 with ExoT+H2O2 and 0.03 with RecJ+H2O2 treatment, and Supplementary Fig. 3g,h). These data with structure-specific ssDNA exonucleases verify that the nascent ssDNA detected by ssDNA-SIRF is located at DNA ends, e.g. not at template-switched or otherwise annealed DNA structures. Taken together the results show that RF-SIRF conditions are amendable for reversed fork detection.

RF-signals form at one-ended DNA structures controlled by fork-reversal enzymes

SMARCAL1 is a major fork reversal DNA translocase, that dislodges nascent DNA strands at stalled DNA forks to drive the formation of reversed fork structures 14, 20, 25. We therefore tested the effect of SMARCAL1 ablation on RF-SIRF, which inhibits fork reversal. For this we induced reversed forks by treating U2OS cells with low-dose CPT, and compared RF-SIRF results with those in cells knocked-down for SMARCAL1 (Fig. 4a, b). In direct reversal to the results obtained under conditions that promote fork reversal (Fig. 1), knocking down SMARCAL1 dramatically reduced RF-SIRF signals caused by a significant reduction in EdU-PLA signal (Fig. 4c,d; median RF-SIRF of 0.4452 with CPT and 0.2411 with CPT plus SMARCAL1 knockdown, and median EdU-PLA intensities of 0.473 with CPT and 0.315 with CPT plus SMARCAL1 knockdown, Supplementary Figs. 4a-c). The reduction cannot be attributed to decreased EdU substrate incorporation as overall EdU- Alexa488 signals are increased, rather than decreased under SMARCAL1 knock-down conditions (Fig. 4e; median EdU-PLA intensities of 0.304 with CPT and 0.405 with CPT plus SMARCAL1 knockdown), which is consistent with accelerated DNA replication that is seen with resolution of reversed forks 39, 50. Similarly, HU-induced RF- SIRF signals are reduced upon SMARCAL1 knockdown, suggesSting RF-SIRF signals measure reversed forks under diverse conditions (Supplementary Fig. 4Sd,e). Corroborating this notion, ssDNA-SIRF signals characteristic of reversed forks too are reduced in cells treated with CPT or HU when SMARCAL1 expression is knocked down (Supplementary Fig. 4f-i). Verifying these results, CPT-induced RF-SIRF signals are starkly reduced in U2OS cells with SMARCAL1, ZRAN3B and HLTF knock-out (3KO 21) that lack all tPhree major fork reversal enzymes (Fig. 4f, median RF-SIRF 0.53 with CPT in U2OS and 0.21 with CPT in 3KO, Supplementary Fig. 4j,k).

DNAPKcs is an obligate DNA-end bindingI protein that binds both dsDNA and ssDNA and more recently has been implied at reversed forks 51,52. DNAPKcs-SIRF signals significantly increase with one hour CPT in U2OS but not 3KO cells, further supporting the formation of reversed forks containing a free end under our testing conditions (Fig. 4g,h, median DNAPKcs- SIRF 0.38 with CPT in U2OS and 0.26 with CPT in 3KO, Supplementary Fig. 4l,m). Taken together, the data shows that limIiting reversed forks by eliminating the fork reverser enzymes SMARCAL1, HLTF and ZRAN3B significantly reduces RF-SIRF signals formed at DNA ends, demonstrating that RF-SIRRF detects reversed DNA replication fork structures.

Spatial and temporal characteristics of RF-SIRF

An advantage using a single-cell fluorescence microscopy-based approach to assess reversed forks is that it provides temporal and spatial resolution of fork reversal reactions within cells allowing to investigate so far unexplored research questions. To further probe the RF-SIRF assay system, we performed cell-cycle analysis of RF-SIRF (Fig. 5a,b). By plotting the cell cycle profiles of cells treated with and without CPT, the data demonstrates that cells in early to mid-S- phase show the greatest increase in RF-signals in cells treated with CPT compared to untreated cells (Fig. 5a,b). Additionally, RF-signals are significantly reduced in late S-phase (Fig. 5c).

Moreover, early to mid S-phase RF-signals formed with CPT or HU are reduced with SMARCAL1 knock-down (Supplementary Fig. 5a-d). Of note, while non-treated cells overall have significantly less RF-SIRF signal compared to those that are stressed by exogenously provided replication stalling agents, there are more RF signals in early-mid compared to late S- phase also in untreated cells (Supplementary Fig. 5e), suggesting that also spontaneous reversed forks occur less frequently in late S/G2. Together, these analyses suggest that reversed forks predominantly form in early/mid S-phase of the cell cycle.

Stalled DNA replication forks can be tethered to nuclear pores at the nuclear periphery 53-55. In addition to temporal resolution, we sought to query the subcellular localization of RF-signals to obtain spatial knowledge of reversed forks. We noticed that RF-SIRF often form a prominent signal at the nuclear periphery in proximity to the nuclear lamina, which is in contrast to EdU- Alexa488 signals (Fig. 5d, Supplementary Fig. 5f). Moreover, the number of cells containing prominent peripheral nuclear RF-ring like structures increase significantly with treatment of CPT (Fig. 5e, 72.6% cells with nuclear ring-like RF-signal with CPT compared to 54.4% without), whereby CPT-treated cells with greater RF-SIRF signals show considerably more RF-SIRF ring- structures compared to those with less RF-SIRF signals (Fig. 5f, top 25% of RF-SIRF signal positive cells with 75.8% ring-structures, compared to bottom 25% of RF-SIRF signal positive cells, with 14.8% ring-structures).

To better dissect the spatial-temporal features of RF-SIRF signals, we performed cell cycle analysis using nuclear EdU-Alexa488 S-phase pattern formation to categorize cells into early-, mid-, and late S-phase cells (Fig. 5g,h and Supplementary Fig. 5g). Investigating the top 25% with RF-SIRF signal positive cells with CPT, the cell cycle pattern of nuclear EdU-Alexa488 revealed predominantly early S-phase cells compared to the bottom 25th percentile, which show a sizable amount of late S-phase EdU patterned cells (SupplemeSntary Fig. 5g). These results mirror the earlier results obtained by cell cycle analysis using SDAPI immunofluorescence (compare Fig. 5c and Supplementary Fig. 5g). No late S-phase patterned cells were detected in the top 25% RF-SIRF percentile.

Within the top 25th percentile of RF-SIRF-positive cells, 73% showed early S-phase diffuse EdU-Alexa488 pattern, with roughly two-thirds of th ese early S-phase cells showing a pronounced ring-shaped RF-SIRF signal (Fig. 5g,h). The remaining 27% cells showed condensing mid-S-phase EdU-Alexa488 pattIerns with ring-shaped RF-SIRF pattern (Fig. 5g,h). Of note, RF-SIRF signals localize to the nuclear periphery irrespective of significant EdU- Alexa488 signals being present within the nuclear lumen, suggesting that peripheral RF-SIRF rings may differentiate between reversed forks and ongoing replication forks in early- and mid- S-phase. In stark contrast, RF-SIRF signals overlap with EdU-Alexa488 signals during late S-phase, which are devoid of RF-rIing structures (Fig. 5g,h). Nevertheless, late S-phase RF-SIRF signals also significantly decreased with SMARCAL1 knockdown (Supplementary Fig. 5a-d, p<0.0001). This suggestsR that RF form in late-S-phase, however, at a nuclear location that is distinct from early S-phAase reversed forks. Collectively, these data suggest that reversed forks preferentially form at the nuclear periphery during early and mid S-phase.

Local chromatin changes at reversed replication forks

While canonically considered for transcriptional control, local chromatin changes during DNA stress are critical intracellular communication markers enabling the specific recruitment of DNA stability factors 17, 34, 35, 55-57. Whether fork reversal reprograms the epigenetic landscape is currently elusive. Having established a robust assay system to interrogate reversed fork reactions, we sought to test its usage for understanding potential epigenetic changes that may occur at reversed forks (Fig. 6a). H3K9me3 is a heterochromatin mark that is induced with replication stress 55, 58,. To test if heterochromatin forms at reversed forks, we performed H3K9me3-SIRF with and without CPT (Fig. 6b,c, and Supplementary Fig. 6a,b). Under conditions promoting fork reversal, H3K9me3-SIRF signals are significantly enriched (Fig. 6c, median H3K9me3-SIRF of 0.347 with CPT and 0.091 without CPT). Importantly, H3K9me3- SIRF signals are reduced to non-stressed levels when knocking down SMARCAL1 (Fig. 6c, median H3K9me3-SIRF of 0.347with CPT and 0.049 without CPT plus SMARCAL1 knockdown). These data demonstrate that H3K9me3 is enriched at reversed replication forks, and not at nascent DNA behind the fork, which is not expected to change with SMARCAL1 knock down.

H4K16ac is an epigenetic mark for transcriptionally active chromatin. In the context of DNA stress responses, it is a critical mark for suppressing TP53BP1 binding to damaged DNA sites 57, 58. TP53BP1 is also recruited to stalled replication forks and acts in replication fork protection 21. Under the same fork reversal promoting conditions that show an increase in H3K9me3, H4K16ac-SIRF signals are significantly decreased (Fig. 6d, median H4K16ac-SIRF of 0.070 with CPT and 0.251 without CPT, and Supplementary Fig. 6c,d). These data show that distinct epigenetic chromatin changes occur under conditions that promote fork reversal.

Like H4K16ac, H3K4m3 is an epigenetic mark for transcriptionally active chromatin. Yet in the context of DNA stress responses, it is required for MRE11 recruitment to stalled replication forks suppressing cancer therapy resistance in BRCA1/2 defective cancer cells 17, 35. Intriguingly, despite both H4K16ac and H3K4me3 marking active transcription, they show opposite behavior at reversed replication forks with CPT, whereby H3K4me3-SIRF signals are significantly enriched at CPT-stalled forks (Fig. 6e, median H3K4me3-SIRF of 0.23 with CPT and 0.143 without CPT, and Supplementary Fig. 6e,f). Taken togSether the data shows that the epigenetic code for replication stress differs from that at transScription sites and explains previously observed proteomic changes at reversed forks.

Seemingly paradoxically, the data suggests that both transcription activating and suppressing epigenetic marks are enriched under rePversed fork conditions. To test if both marks occur concomitantly at reversed forks, we performe d a PLA assay between H3K4me3 and H3K9me3. H3K4me3-H3K9me3 PLA signals increase significantly with CPT, suggesting simultaneous enrichment at reversed forks (FIig. 7a,b, median H3K4me3-H3K9me3 PLA of 0.42 with CPT and 0.29 without CPT, Supplementary Fig. 7a). Confirming that bivalent H3K4me3- H3K9me3 are induced with fork reversal, the PLA signals are significantly reduced in 3KO cells (Fig. 7a,b, median H3K4me3-H3K9me3 PLA of 0.43 with CPT compared to 0.25 with CPT in 3KO cells). Importantly, the observed chromatin-modification changes cannot be explained by total histone changes as evidenIt by H3-SIRF that in contrast to the bi-valent H3K9me3- H3K4me3 signals, slightly decrease with CPT (Fig. 7c,d, median H3-SIRF PLA of 0.34 without and 0.271 with CPT, and RSupplementary Fig. 7b,c). These data imply that active chromatin modification changes oAccur at reversed forks, resulting in an over-representation of otherwise rare transcriptionally bivalent H3K4me3-H3K9me3 marks.

Local chromatin changes principally can occur via re-localization or new deposition 55, 59, 60. Given that reversed forks preferentially form during early S-phase when heterochromatin is principally limited, and total H3-association with nascent DNA does not increase with fork reversal, we considered active chromatin-mark depositing as a means for epigenetic changes at reversed forks. In the context of stalled replication forks, PTIP, which is the functional binding partner of the MLL3/4 methyltransferases, is required for H3K4 tri-methylation promoting MRE11 recruitment 17, 61. Using HAP1 cells, H3K4me3-SIRF signals significantly increase with CPT confirming the previous results in multiple cell lines (Fig. 7e,f, median H3K4me3-SIRF of 0.196 with CPT and 0.036 without, Supplementary Fig. 7d,e). Contrasting wild-type HAP1 cells, H3K4me3-SIRF is severely compromised in PTIP knock-out HAP1 cells (Fig. 7e,f, median H3K4me3-SIRF of 0.083 with CPT and 0.091 without, Supplementary Fig. 7d,e). Collectively, the data suggest that PTIP mediates de novo H3 tri-methylation at reversed forks.

Methods
Reagents, cell lines and culture conditions: Hydroxyurea (HU, H8627), Aphidicolin (A0781), Hydrogen Peroxide (H1009), X-tremeGENE™ HP DNA Transfection Reagent (6366244001) and Duolink proximity ligation assay reagents (DUO92001, DUO92005, and DUO92008) were from Sigma-Aldrich; Ethynyl-2'-deoxyuridine (EdU, A10044), biotin azide (B10184), Alexa Fluor 488 azide (A10266), Alexa Fluor 647 azide (A10277) and ProLong Gold (P36934) were from Invitrogen; 4′,6-diamidino-2- phenylindole (DAPI, 62248) and Lipofectamine™ RNAiMAX Transfection Reagent (13778075) was obtained from ThermoFisher Scientific; 32% Paraformaldehyde (PFA, 15714) was from Electron Microscopy Science, protein block buffer was from Abcam (ab64226), Camptothecin (S1288) was from Selleck chemicals, T4 DNA ligase buffer from New England Biolabs (B0202S). Antibodies used for SIRF assays are as follows: mouse anti-biotin (BN-34, 1:100; Sigma-Aldrich), rabbit antiS-biotin (D5A7, 1:200; Cell Signaling Technology), mouse anti-dsDNA (MAB1293, 1:100; Millipore), mouse anti- dsDNA (CBL186, 1:100; Millipore), rabbit anti- GFP (2555S, 1:200; Cell signaling technology), rabbit anti-H3K9me3 (ab8898, 1:500; Abcam), mouse anti-ssDNA (clone 16-19, 1:100; Millipore), mouse anti- Lamin A/C (4C11, 1:50; Cell signaling technology) rabbit anti-H3K4me3 (ab8580, 1:500; Abcam), rabbit anti-H4K16ac (ab109463, 1:200; Abcam), mouse anti-BrdU (B-44, 1:100; BD34 7580), rabbit anti-BrdU (1:400; GTX128091), mouse anti-DNAPKcs (1:100; sc-390849), rabbit anti-histone H3 (1:200; ab1791), mouse anti-H3K4me3 (1:100; Sigma Aldrich 05-1339), mouse anti-MRE11 (1:200, ab214). Antibodies used for iEmmunoblotting are mouse anti-SMARCAL1 (SC- 376377, 1:500; Santa Cruz BiotecLhnology) and mouse anti-beta-actin (A5441, 1:5000, sigma). SMARCAL1 siRNA waCs obtained from Horizon (J-013058-06-0002). Reagents for ExoT/RecJ nuclease assays: MOPS (Sigma M1254), MgCl2 (Ambion AM9530G), Sucrose (Sigma S0389), RTecJ (NEB M0264L), ExoT (NEB Mo265L) U2OS cells (ATCC) and U2OS 3KOR (gift from the David Cortez lab) were grown in Dulbecco's Modified Eagle Medium (DMEM, Life Technologies) supplemented with 10% fetal bovine serum (Gemini Bio-Products). HAP1 PTIP (Horizon Discovery) were grown in IMDM (Life Technologies) supplemented with 10% fetal bovine serum (Gemini Bio- Products)

Western blot for siRNA: SMARCAL1 siRNA transfection was performed using RNAiMAX for 48 hrs following manufacturer’s instructions. Cell pellets were lysed in NETN buffer (50 mM Tris-HCl, pH 7.4, 1% NP-40, 200 mM NaCl, and 1 mM EDTA) on ice for 30mins. Lysates were centrifuged at 13,000 × g , 4°C for 15 mins and supernatant proteins were resolved by SDS-PAGE. Proteins were transferred to PVDF membranes, blocked with TBST and 5% milk for 1 hr and incubated with primary antibodies overnight. Corresponding secondary antibodies were incubated for 1 hr at RT. Signals were detected using enhanced chemiluminescence.

RF-SIRF protocol: Cells are grown in log-phase and plated the day before the experiment at 50–60% confluence onto microscope chamber slides (NuncTM Lab- TekTM). Cells are treated with 125 µM EdU for 8 minutes, washed twice with Phosphate buffered saline (PBS, pH7.4), followed by addition of media (no treatment), or genotoxic agent for 1 hour or 15 minutes, as indicated in the figures. Cells are fixed with 4% Paraformaldehyde (PFA, Electron Microscopy Science) in PBS (pH 7.4) for 15 minutes at room temperature. After fixation, slides are washed two times with PBS (pH 7.4) for 5 minutes each. Cells are permeabilized using 0.25% Triton X-100 in PBS (pH 7.4) for 15 minutes at room temperature and washed in PBS (pH 7.4) twice for 5 minutes each. Click-iT reaction cocktail is prepared fresh using 2mM copper sulfate (Sigma-Aldrich) 9 µM biotin-azide (Invitrogen), 1 M Alexa-488 azide (Invitrogen) and 100mM freshly prepared sodium ascorbate (Sigma-Aldrich) in PBS (pH 7.4) in that order and mixed. Click-It reaction cocktail is added to slides (30 µl/well) and incubated at room temperature for 30 minutes in a humid chamber. Slides are washed three times in PBS (pH 7.4) for 5 minutes each and blocked with protein block buffer (ab64226, Abcam) for 1 hour at room temperature in a humid chamber. Primary antibodies (anti-mouse-biotin and anti-rabbit-biotin for RF-SIRF) are diluted in blocking buffer, dispensed onto slides (30–40 µl/well), and incubated at 4°C overnight in a humid chamber. Slides are washed three times with wash buffer A (0.01 M Tris, 0.15 M NaCl, and 0.05% Tween 20, pH 7.4, (Sigma Duolink) for 5 minutes each. Duolink In Situ PLEA probes anti–mouse plus and anti–rabbit minus are diluted 1:5 in blocking solutioRn and incubated on slides for 1 hour at 37°C. Slides are again washed in wash buffer A solution three times for 5 minutes each. Pre-ligation mix is prepared by diluting D uolink ligation stock (1:5) in sterile water. Slides are incubated in humid chamber with pre-ligation mix at 37°C for 30 min and washed in wash buffer A three times for 5 minutes each. Slides are incubated with ligation mix [1x T4 ligase buffer (B0202 S, New England Biolabs), 1X Duolink ligase enzyme in sterile water] in the humid chamber at 37°C for 30 minutes, followed by three washes with wash buffer A three times for 5 minutes each. PLA amplification mix is prepared according to manufacturer’s instructions (Duolink, Sigma) and slides are incubated with it at 37°C for 100 minutes in a humid chamber. Slides are washed in wash buffer B solution (0.2M Tris and 0.1M NaCl, Sigma Duolink) three times for 10 minutes each and one time in 0.01x diluted wash buffer B solution for one minute. Slides are incubated with 4′,6-diamidino-2-phenylindole (DAPI, 1µg/ml; Life Technologies) at room temperature for 5 minutes, followed by one wash with PBS (pH 7.4) for 5 minutes before mounting with Prolong Gold antifade (Life Technologies). Slides are imaged using a Nikon Eclipse Ti-U inverted microscope and analyzed using Nikon NIS elements software.

Image acquisition and analyses: Images are obtained using Nikon Eclipse Ti-U inverted microscope with Andor Zyla camera and Plan Apochromat objective lens at 40X magnification. 3-6 image fields were acquired for each condition, yielding 100-300 nuclei for image analyses per condition and biological independent experiment. Mean fluorescent intensity for DAPI, GFP, Texas-red and Cy5 channels is determined using NIS elements software, except Fig. 4h, Supplementary Fig.1 e-g and 4l,m, which were determined using ImageJ software with macro provided in Supplementary Data. RF- SIRF and SIRF values are calculated using the ratio between PLA intensity and corresponding EdU-Alexa488 intensity per cell. Fluorescent intensity data is min-max range normalized for each independent biological experiment (x = (x-x ) / normalized minimum (x -x ). The Minima and maxima are determined based on the entirety of a maximum minimum given data set that is being compared. Normalized values are pooled from 2-4 biological repeats as indicated in the figure legends. The two-sided Mann-Whitney test is applied to determine statistical significance when two variables are being compared. The one-way ANOVA is applied to determine statistical significance when more than two variables are being compared. For cell cycle plots, the sum DAPI fluorescence intensity is plotted on the X-axis and the mean EdU-Alexa488 intensity is plotted on the Y-axis after min-max normalization. The data point color represents the RF-SIRF value. The RF-SIRF values plotted on the cell cycle plots are min-max normalized for each biological repeat between NT and CPT conditions for direct comparison between the two conditions.

DNA Fiber Assay: DNA fiber experiments were performed as described previously 10, 11, 24, 62, 65. Briefly, log-phase cells were pulse-labeled with 50 μM IdU for 30 min and 50 M CldU for 15min with or without replication stress agents as indicated in figures. Cells were harvested, lysed, and spread to obtain single DNA molecules on microscope slides before standard immunofluorescence with antibodies against IdU and CldU (BD Biosciences, GeneTeX).

ssDNA SIRF with nuclease treatment: For nucPlease assay, cells were washed with PBS after treatments in chamber slides and pr e-extracted with CSK100 buffer (100mM NaCl, 10mM MOPS pH7, 3mM MgCl , 300NmM sucrose, 0.5% TritonX-100) in RT for 10min. Cells were washed and treated foIr 30min at 37C with respective nucleases (20 units/ml in reaction buffer). After nuclea se treatment, solutions were aspirated, and cells were fixed with 4%PFA and proceeded for SIRF assay as described 33.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "The manuscript by Roy et al describes an application of their original SIRF assay to detection and quantification of reversed replication forks in vivo. Their approach is ingenious, innovative and can potentially simplify and expedite replication fork metabolism studies, while also adding a single-cell and intranuclear spatial resolution levels to the analysis. This is an important work that will offer immediate, practical benefits to the community of researchers working in the areas of replication stress and genomic instability in relation to cancer and aging. I hope the authors can address two questions that should make their claims more robust.\n1) The nature of the fork structure recognized by RF-SIRF. siSMARCAL1-dependent reduction of the RF-SIRF signal is used as proof that RFs are the substrate. Ideally, additional lines of evidence are needed. Can template-switched strands in replicated DNA away from the fork or replicated sister duplexes that are in close proximity due to sister cohesion be recognized as well? Both of these can in theory be affected by CPT or other damage. By definition, a RF occurs at a replication junction with extrusion of free ends of leading and lagging strands. It can be thus inferred that the fraction of EdU-labeled DNA involved in daughter strand pairing depends on: 1) how much DNA the fork was synthesized since the EdU pulse and 2) how far did reversal go. These two factors apply even if replication during CPT treatment is viewed as a persistence of a dynamic equilibrium between forks moving forward and reversing themselves. To verify that their conditions are permissive for pairing of the EdU-labeled daughter strands the authors need supporting data for the CPT and other treatments, showing what length of DNA is synthesized in their model system in 1hr after an EdU label pulse (by carrying out DNA fiber analyses). This could include conditions with and w/o reversal (siSMARCAL1) to estimate the length of reversed arms (or with and w/o degradation to estimate the same, assuming the entire reversed arm is degraded). Also, a testable prediction should be that the RF-SIRF signal will decrease as a function of the duration of the post-EdU period since the EdU segment will be less and less likely to be in the reversing segment. The data will help to boost the interpretation that only RFs but not the template switched segments internal to the replicated DNA or even the newly replicated sisters contribute to the EdU/EdU PLA signal. Does RF-SIRF signal show the same pattern if the drugs are used at concentrations that completely block fork movement (I.e. HU at 2-4mM)?\nIn addition, EdU concentration is quite high, 125uM, and it does not appear that the authors use a thymidine chase. It is likely that the internal pool of EdU will enable additional incorporation after the end of pulse (for example, this paper http://dx.doi.org/10.1016/j.cub.2016.07.020 showed that EdU accumulation in DNA continued for 9 min after a 30 sec pulse with 10uM EdU). Likely, this accounts for higher EdU level in NT controls compared to the CPT-treated samples 1hr after the EdU pulse that the authors see. If so, the EdU-substituted positions produced during the chase may be, for example, too widely separated (in accordance with the authors paper, ref 36) to produce PLA signal in NT but not in CPT-treated samples, due to a more rapid global EdU/dT re-equilibration in NT with its faster forks/more origin firing.\nParticularly because the authors showed that the intensity of EdU/EdU SIRF in-cis (on the same daughter strand) depends on the distance between EdU residues in a labeled DNA fiber (ref 36), why do they use the concentration that they showed to minimize this distance and thus maximize in-cis SIRF signal in regular replicated DNA? Would it not be more informative to use lower EdU concentrations, 10-20uM, to produce widely spaced EdU positions in DNA that may not be SIRF-able in-cis but will be SIRFable in-trans, i.e., in a reversed fork? Can the authors show data on EdU titration effects on RF-SIRF?\n2) The interpretation of enrichment of RF-SIRF signal at the nuclear periphery. Line 266: “RF-SIRF signals show a preference for the nuclear periphery and are more prevalent during early/mid S-phase of cell cycle, a pattern shared with heterochromatin” -- this is potentially an overstatement or a confusing statement. The references given in its support describe relocation of heterochromatic DSBs and collapsed forks to nuclear periphery/pores. First, this relates to heterochromatin that is not already at the nuclear periphery, i.e., nucleolar heterochromatin. Second, more importantly, the authors’ observation that peak RF SIRF signal is in early/mid S phase (rather than, say, mid/late) is inconsistent with the assertion that these RFs come from replicating heterochromatin. Do the authors suggest that RF-SIRF detects a particular subset of early S phase-replicating, non-peripherally localized heterochromatin? This is a speculation I am not sure the authors need to make. The authors’ detection of H3K9me3 gain and H4K16ac loss at stalled forks mirrors the findings of Gaggioli et al (ref 52) on de novo assembly of heterochromatin-like state at stalled forks that supports their protection form degradation, as the authors point out in the discussion. However, at present there is no strong evidence in this manuscript or elsewhere, as far as I know, to suggest that all RFs, euchromatic or not, go to the nuclear periphery in mammalian cells, or that heterochromatin is more prone to RF formation than euchromatin. In fact, from a recent paper 10.1093/nar/gkae721 it can be inferred that RFs are degraded more and thus are shorter, in effect, less abundant in heterochromatin, though the paper did not examine specifically the RF formation step. This paper was not referenced in the manuscript. The enhanced peripheral RF SIRF alone is insufficient to draw conclusions about RF-SIRF and heterochromatin. If the authors want to make this conclusion they need more data, for example, use the Gaggioli et al de novo fork heterochromatization mutants to see if any of these disrupt peripheral localization of RF-SIRF signal. Does peripheral ring of RF-SIRF appear in early S cells?\nMinor points:\nFigure 4C-E: please include siSMARCAL1 data for non-treated control.\nFigure 5b: RF-SIRF intensity color code was missing in the figure.\nFigure 5c: how were cells selected for early vs late S phase? Was it based on the incorporation pattern or EdU incorporation vs DAPI content?", "reviewer_2": "In this work Roy et al present an original method named RF-SIRF based on proximity ligation assay to quantify reversed forks (RFs), they also suggest that an epigenetic code for replication stress may exist. The paper is well written with clear a clear introduction, the discussion may be improved by discussing more some results (see below).\nIn the recent years RFs has been observed in various contexts, however the lack of simple methods renders their study difficult. The state of art method to quantify them is electronic microscopy, however this approach is really tough and not accessible to most researchers of the field. Clearly there is a need for an easier method. Roy et al developed the RF-SIRF method that relies on Proximity Ligation Assay, a method accessible to most researchers that requires a low number of cells. The results are clear and the required controls are present, although I have comments (see below). It seems that the RF-SIRF is a robust method that will be really useful for many researchers in the field.\nI am little bit less convinced by the part regarding the spatio-temporal characteristics of RFs and the possible existence of a specific epigenetic code. To improve this part, I recommend experiments (see below) and new analysis and also strongly suggest the reviewers to discuss more the data.\nAltogether, this a very interesting paper, really useful for the community in term of methodology, that deserves to be published. In contrast the new results leave one hungry and should be improved before publication.\nMajor comments\nLines 137-138: I do not understand why CPT would decrease EdU-Alexa488 signal since CPT is added after EdU and therefore should not impact its incorporation. Does it mean there is DNA resection? In fact they do write at line 168 that the loss of EdU signal is caused by loss of fork protection. Authors need to homogenize here.\nIn several figures (1C, 1D, 2A etc.) it is written norm. int., I supposed it stands for normalized intensities but it should be described in the legend. According to the methods section, they did a normalization procedure in order to pool together 2-4 experiments, it makes sense since PLA intensities varies between experiments. But it would be nice to show systematically the repetitions in Supdata like they did for Fig1 and Fig 3.\nIn Fig4 the authors show clear data demonstrating that, as expected, SMARCAL1 depletion decreases the formation of RFs using RF-SIRF. I think it would nice to see if this is also the case for the other translocases (HLTF and ZRANB3) known to be involved in fork reversal. It would strengthen the results.\nOne of the most important issue of the paper is that the authors does not really consider S-phase patterns in their analysis (Fig5), I think it could provide a more precise description of the results. It is well established that the EdU patterns are diffused in early-S (euchromatin), more condensed in mid-S (facultative heterochromatin) and very condensed in late S (constitutive heterochromatin replication). For instance, the ring in Fig5D could correspond to heterochromatin replication and not to a relocalization of the fork to the nuclear membrane, the patterns of EdU presented in this figure could actually correspond to mid-S in my opinion. Actually, they do write on line 266 that RF-SIRF signals have a preference for early/mid S, but on Fig5C the just compare early-S and late-S. At the end I am a little bit lost. To increase the clarity of the results, it would be determinant to clearly decipher if RF-SIRF is occurring more in early-S or in mid-S. To do so the authors need to re-analyze the RF-SIRF data using the replication patterns and present the RF-SIRF level in Mid-S vs Early-S cells, this would complement their study based on dapi level in a more precise way.\nThe data from Fig6 regarding the increase level of H3K9me3/H3K4me3 and decrease level of H4K16ac during fork reversal induced by CPT are interesting. However, I do not really understand the mechanism. Does it mean that somehow these modified histones are recruited or excluded during fork reversal? But how? Or does it mean that the enzymes required (such as SUV39H1 for H3K9me3 or MLL3/4 for H3K4me3) for these modifications are recruited to reversed fork? I think the authors should discuss more their data and provide a little bit of mechanistic otherwise it remains really descriptive.\nOn line 293 they write: “these data explain how MRE11 is recruited there”. I think the data provided in the current version of this manuscript are not sufficient to support completely this claim. As cited by the reviewer (Ray Chauduri 2016), MLL3/4 is required to recruit MRE11 to reversed forks. It would great if the authors could test by SIRF the impact of MLL3/4 inactivation on MRE11 recruitment and on H3K4me3 recruitment in the conditions used in Fig6.\nMinor comments\nI would put the SupFig1A instead of 1A because it simplifies the life of the reader to understand the principle of the method directly.\nOn line 187 the write that the strongest effect of RF is H2O2, in Zelwegger 2015 JCB they also found that H2O2 has one of the strongest effects (with CPT). I would be curious to know authors hypothesis to explain this.\nOn Fig3 and SupFig3 it is written that data are derived from3 independent repeats, why only 2 repeats are shown for SupFig3 c, d and e?\nIn Fig5 the gradient colors are missing.", "reviewer_4": "Replication fork reversal is a protective mechanism during DNA replication stress that occurs when replication forks encounter an obstacle and reverse their course, forming a four-way \"chicken foot\" structure through the annealing of newly synthesized DNA strands, while the parental strands re-anneal. This structure has gained significant interest over the past decade, as it stabilizes the replication fork, prevents DNA damage, and allows time for repair. As stated by the authors, direct analysis of these structures requires complex electron microscopy-based techniques, mastered by only a few specialized laboratories worldwide. In this article, Roy and colleagues propose an elegant new approach to monitor fork reversal using a modified proximity ligation assay (PLA) that specifically detects EdU on the regressed arm of the reversed fork. The assay is based on the bold assumption that EdU epitopes incorporated into the two sister chromatids of a normal Y-shaped fork do not generate a PLA signal, due to their physical separation—even though they are held together by cohesin. Nevertheless, the authors convincingly demonstrate a specific RF-SIRF signal at reversed forks, normalized to total EdU incorporation. They further show that this signal increases upon treatment with drugs known to induce fork reversal and is lost following depletion of SMARCAL1, a key factor involved in fork reversal. Additionally, they show that this assay provides insights into the subnuclear organization of reversed forks, the presence of single-stranded DNA (ssDNA) at reversed forks, and their relative frequency during S phase. Altogether, RF-SIRF emerges as a promising assay that complements indirect methods such as DNA fiber analysis and serves as an accessible alternative to electron microscopy. This approach fulfills a clear need in the field and will be of great utility to the growing number of laboratories investigating replication stress responses and fork reversal. However, the authors should address the following points to strengthen their conclusions:\n1. Simultaneous click chemistry using biotin-azide and Alexa488-azide may compete for the single alkyne group on EdU. This competition could significantly impact data interpretation and may explain the substantial variability observed in EdU-PLA signals across biological replicates, as shown in Extended Data Fig. 1.\n2. In the legend of Extended Data Fig. 1a, the authors mention anti-biotin primary antibodies. These should presumably be derived from two different species (e.g., mouse and rabbit). The authors should clarify this in the Methods section.\n3. In the ssDNA-SIRF assay, what is the minimum ssDNA length required for effective recognition by the anti-ssDNA antibody?\n4. Could the authors perform a time-course experiment to assess whether RF-SIRF signals increase during genotoxic treatment in the presence of nuclease inhibitors such as Mirin?\n5. The authors present convincing evidence that ssDNA can be detected at reversed forks upon resection. It would be valuable to demonstrate that the RF-SIRF signal decreases as ssDNA-SIRF increases. A time-course experiment in BRCA2-deficient cells could illustrate these signal dynamics.\n6. The EdU-H3K9me and H4K16ac-EdU PLA experiments do not presently add substantial value, as the authors do not directly demonstrate that the observed signal arises from reversed forks. Unless supported by further experimental validation, this section should be removed.\n7. The word \"telomere\" is misspelled multiple times in the manuscript and should be corrected throughout."}, "author_response": "We thank the reviewers for their expert, rigorous, and thoughtful reviews. We appreciate their enthusiastic comments and consequently put major resources into the added new data resulting in 18 new Figure panels in the main Figures and 25 new Figure panels in the Extended Data Figures. We also did re-analysis and editorial changes to address all reviewers’ concerns, which without doubt greatly improved the manuscript. Please see reviewer remarks in italic, and our responses marked in blue font.\nReviewer #1 (Remarks to the Author):\nThe manuscript by Roy et al describes an application of their original SIRF assay to detection and quantification of reversed replication forks in vivo. Their approach is ingenious, innovative and can potentially simplify and expedite replication fork metabolism studies, while also adding a single-cell and intranuclear spatial resolution levels to the analysis. This is an important work that will offer immediate, practical benefits to the community of researchers working in the areas of replication stress and genomic instability in relation to cancer and aging. I hope the authors can address two questions that should make their claims more robust.\nThank you for the encouraging comments and thoughtful assessment. We split up responses for question nr 1) into multiple sections.\n1) The nature of the fork structure recognized by RF-SIRF. siSMARCAL1-dependent reduction of the RF-SIRF signal is used as proof that RFs are the substrate. Ideally, additional lines of evidence are needed. Can template-switched strands in replicated DNA away from the fork or replicated sister duplexes that are in close proximity due to sister cohesion be recognized as well?\nBoth of these can in theory be affected by CPT or other damage. By definition, a RF occurs at a replication junction with extrusion of free ends of leading and lagging strands.\nThe ssDNA SIRF data in Fig. 3 argue against a template switch product of nascent strands, which in contrast to reversed forks does not contain nascent ssDNA.\nNonetheless, we now added multiple additional lines of evidence that the RF-SIRF products detect reversed replication forks and are unlikely to represent template switched products under the conditions tested.\n1) We employed structure specific ssDNA exonucleases as evidence that the RF-SIRF products contain DNA ends with overhangs, which do not occur at template switched/paired regions. Specifically, ExoT is a 3’-5’ssDNA exonuclease, and RecJ is a 5’-3’ ssDNA exonuclease that cannot act on dsDNA or gapped DNA.\nUsing ssDNA-SIRF, which increases under fork-reversal conditions, we show that the products are significantly decreased when cells are processed with either ExoT or RecJ nucleases. Since neither nuclease is active on gapped DNA or dsDNA, this provides strong evidence that ssDNA-SIRF signals are located at DNA ends (added Fig. 3d,e, Ext. Data Fig. 3g,h)\n2) We show increased binding of DNAPKcs, a canonical DNA end binding protein, to nascent forks under fork reversal conditions, which reduces in cells devoid of fork reversal enzymes (“3KO” cells with knock out of SMARCAL1, ZRAN3B and HTLF). (added Fig.4 g,h, Ext. Data Fig. 4l,m)\nIt can be thus inferred that the fraction of EdU-labeled DNA involved in daughter strand pairing depends on: 1) how much DNA the fork was synthesized since the EdU pulse and 2) how far did reversal go. These two factors apply even if replication during CPT treatment is viewed as a persistence of a dynamic equilibrium between forks moving forward and reversing themselves. To verify that their conditions are permissive for pairing of the EdU-labeled daughter strands the authors need supporting data for the CPT and other treatments, showing what length of DNA is synthesized in their model system in 1hr after an EdU label pulse (by carrying out DNA fiber analyses).\nThis could include conditions with and w/o reversal (siSMARCAL1) to estimate the length of reversed arms (or with and w/o degradation to estimate the same, assuming the entire reversed arm is degraded). Also, a testable prediction should be that the RF-SIRF signal will decrease as a function of the duration of the post-EdU period since the EdU segment will be less and less likely to be in the reversing segment. The data will help to boost the interpretation that only RFs but not the template switched segments internal to the replicated DNA or even the newly replicated sisters contribute to the EdU/EdU PLA signal. Does RF-SIRF signal show the same pattern if the drugs are used at concentrations that completely block fork movement (I.e. HU at 2-4mM)?\nTo our understanding, there is no data to believe that fork reversal is dependent on DNA synthesis post replication stalling. Rather, the previously synthesized strands prior to DNA damage make up the reversed strands. Following the reviewer’s thoughtful suggestions, we further provide added data here for the latter:\n1) Chasing the EdU label with media before stalling by CPT progressively reduces RF signals, showing that it is the nascent DNA synthesized prior – not after- CPT that makes up RF-SIRF signals and reversed fork (added Ext. Data Fig 1h-j).\n2) We also now show that RF-SIRF is significantly increased with high concentrations of HU when DNA synthesis after stalling is greatly inhibited as seen by DNA fibers analysis (added Fig. 2 g-j, Ext Data Fig. 2y).\n3) Moreover, while there can be residual EdU incorporation after EdU wash out (Ext. Data Fig. 1d), nucleoside incorporation post-DNA damage is slowed with CPT treatment compared to non-treated cells as seen by fiber analysis (Ext. Data Fig. 2y), thus cannot explain the increase in EdU-PLA with CPT.\nIn addition, EdU concentration is quite high, 125uM, and it does not appear that the authors use a thymidine chase. It is likely that the internal pool of EdU will enable additional incorporation after the end of pulse (for example, this paper http://dx.doi.org/10.1016/j.cub.2016.07.020 showed that EdU accumulation in DNA continued for 9 min after a 30 sec pulse with 10uM EdU). Likely, this accounts for higher EdU level in NT controls compared to the CPT-treated samples 1hr after the EdU pulse that the authors see. If so, the EdU-substituted positions produced during the chase may be, for example, too widely separated (in accordance with the authors paper, ref 36) to produce PLA signal in NT but not in CPT-treated samples, due to a more rapid global EdU/dT re-equilibration in NT with its faster forks/more origin firing.\nParticularly because the authors showed that the intensity of EdU/EdU SIRF in-cis (on the same daughter strand) depends on the distance between EdU residues in a labeled DNA fiber (ref 36), why do they use the concentration that they showed to minimize this distance and thus maximize in-cis SIRF signal in regular replicated DNA? Would it not be more informative to use lower EdU concentrations, 10-20uM, to produce widely spaced EdU positions in DNA that may not be SIRF-able in-cis but will be SIRFable in-trans, i.e., in a reversed fork? Can the authors show data on EdU titration effects on RF-SIRF?\nIn hindsight, the increased PLA signal in re 36 may have been a reflection of increased detetion of endogenous reversed forks.\nThis is a logical hypothesis. Alternatively, a productive PLA signal requires a minimum of 40nm distance or less between EdU moieties. Thus, even if within a reversed fork, if the overall concentration and incorporation rate results in EdU moieties that are too far apart, it will not result in a productive PLA signal irrespective if at reversed forks or in cis. The latter is supported by our new data: we added EdU titration data and show that 125uM EdU provides optimal detection of reversed forks. (Ext. Data Fig. 1e-g). Indeed, the combined data argues that both, the high EdU concentration and normalization to total EdU (e.g. Alexa488) are critical components in detecting reversed forks vs. simply nascent forks.\nWe conclude that these data explain an apparent discrepancy between our results that show H3K9me3 enriched at reversed forks in multiple independent cell settings, and those results reported by the Taneja group. The group found that while H3K9me3 is upregulated at nascent forks, it was unaffected by SMARCAL1 knock down under their experimental conditions, with these negative results prompting them to conclude it is not enriched at reversed forks (Gaggiolo et al Nature Cell Biol 2023). However, the group applied a straightforward PLA between H3K9me3-EdU (e.g. not SIRF conditions with the important normalization) and with low 25uM EdU concentrations. We show that low EdU concentration is suboptimal for detecting reversed forks. Furthermore, as they do not normalize the data to total amounts of EdU, interpretations for the cause of PLA signal change are problematic.\nWe now note the important distinction between SIRF and EdU-PLA for reversed fork detection in the manuscript. We agree with the reviewer’s assessment that in hindsight, the increased EdU-PLA signals with high EdU concentration seen in our original JBC SIRF paper could have detected endogenous RFs.\n2) The interpretation of enrichment of RF-SIRF signal at the nuclear periphery. Line 266: “RF-SIRF signals show a preference for the nuclear periphery and are more prevalent during early/mid S-phase of cell cycle, a pattern shared with heterochromatin” -- this is potentially an overstatement or a confusing statement. The references given in its support describe relocation of heterochromatic DSBs and collapsed forks to nuclear periphery/pores. First, this relates to heterochromatin that is not already at the nuclear periphery, i.e., nucleolar heterochromatin. Second, more importantly, the authors’ observation that peak RF SIRF signal is in early/mid S phase (rather than, say, mid/late) is inconsistent with the assertion that these RFs come from replicating heterochromatin. Do the authors suggest that RF-SIRF detects a particular subset of early S phase-replicating, non-peripherally localized heterochromatin? This is a speculation I am not sure the authors need to make. The authors’ detection of H3K9me3 gain and H4K16ac loss at stalled forks mirrors the findings of Gaggioli et al (ref 52) on de novo assembly of heterochromatin-like state at stalled forks that supports their protection form degradation, as the authors point out in the discussion. However, at present there is no strong evidence in this manuscript or elsewhere, as far as I know, to suggest that all RFs, euchromatic or not, go to the nuclear periphery in mammalian cells, or that heterochromatin is more prone to RF formation than euchromatin. In fact, from a recent paper 10.1093/nar/gkae721 it can be inferred that RFs are degraded more and thus are shorter, in effect, less abundant in heterochromatin, though the paper did not examine specifically the RF formation step. This paper was not referenced in the manuscript. The enhanced peripheral RF SIRF alone is insufficient to draw conclusions about RF-SIRF and heterochromatin. If the authors want to make this conclusion they need more data, for example, use the Gaggioli et al de novo fork heterochromatization mutants to see if any of these disrupt peripheral localization of RF-SIRF signal. Does peripheral ring of RF-SIRF appear in early S cells?\nWe agree that this sentence was confusing. We reworded this section to clarify, thank you!\nThe nuclear rings of RF-SIRF are a noteworthy observation, not necessarily an inference regarding chromatin states, but a distinct one that can be further investigated in detail in future studies. In our initial analysis we used cell-cycle profiles to determine early/mid S-phase RF-SIRF ring localization. We now added S-phase pattern analysis, which confirm that the highest RF-SIRF signals are found in early S-phase coinciding with ring-patterns of RF-SIRF signals (added Fig.5f-h, Ext. Data. Fig 5g,h). Ring patterns are distinct for EdU-PLA as they do not follow EdU-Alexa488 patterns in early S-phase (see Fig. 5h). Moreover, PLA rings ensue into some mid-S-phase cell. However, also in these cells RF-SIRF signals do not simply follow Alexa488-EdU patterns (see Fig. 5h). e.g. as is seen in early S-phase cells, there is little or no RF-SIRF signal within the cell lumen also in mid-S-phase cells despite that there is ample EdU-Alexa488 incorporation there. This suggests that the RF-SIRF ring pattern is indicative of RF structures, rather than a reflection of mid-S-phase cells ring patterns. In contrast, PLA signals predominantly overlap with Alexa488-EdU in late S-phase cells. Notably, our data shows that RFs also form in late S-phase as the signals reduce with SMARCAL1 knockdown (Ext. Data Fig. 5h). However, in late S-phase RF-signals do not localize to the nuclear periphery but instead are exclusively within the cellular lumen, thus strikingly differ from early S-phase RFs.\nRegarding chromatin states, we have added multiple data figures, which we believe will better clarify the advances and discoveries (Fig. 7a-f and Ext. Data Fig. 7a-e). It is important to differentiate between global heterochromatin versus local epigenetic changes at stalled DNA replication forks, which our data shows includes both markers classically associated with transcriptional repressive heterochromatin and transcriptionally active euchromatin. Of note, our analysis at reversed forks critically extends the Gaggioli findings in 1) showing that both H3K9me3 (classically transcription repressing) and H3K4me3 (transcription activating, not tested in Gaggioli) increase at reversed replication forks, and show that these otherwise rare bi-valent marks are significantly increased at reversed forks, e.g. reduce with fork-reverser knock-outs (added Fig. 7a-b). Total H3-SIRF on the other hand does not increase at RFs (7c,d), suggesting epigenetic modification changes at stalled forks. Consistently, we find that epigenetic modifications are dependent on PTIP, which is MLL3/4-binding partner required for H3K4 tri-methylation, validating the epigenetic changes at stalled and reversed forks as new events.\n[REDACTED]\nMinor points:\nFigure 4C-E: please include siSMARCAL1 data for non-treated control.\nNow added (Ext. Data Fig. 4a-c)\nFigure 5b: RF-SIRF intensity color code was missing in the figure.\nNow added\nFigure 5c: how were cells selected for early vs late S phase? Was it based on the incorporation pattern or EdU incorporation vs DAPI content?\nFig. 5c was selected based on EdU vs DAPI. We now added also analysis based on EdU Pattern, which matches the previous analysis (Fig. 5f-h).\nReviewer #2 (Remarks to the Author):\nIn this work Roy et al present an original method named RF-SIRF based on proximity ligation assay to quantify reversed forks (RFs), they also suggest that an epigenetic code for replication stress may exist. The paper is well written with clear a clear introduction, the discussion may be improved by discussing more some results (see below).\nIn the recent years RFs has been observed in various contexts, however the lack of simple methods renders their study difficult. The state of art method to quantify them is electronic microscopy, however this approach is really tough and not accessible to most researchers of the field. Clearly there is a need for an easier method. Roy et al developed the RF-SIRF method that relies on Proximity Ligation Assay, a method accessible to most researchers that requires a low number of cells. The results are clear and the required controls are present, although I have comments (see below). It seems that the RF-SIRF is a robust method that will be really useful for many researchers in the field.\nI am little bit less convinced by the part regarding the spatio-temporal characteristics of RFs and the possible existence of a specific epigenetic code. To improve this part, I recommend experiments (see below) and new analysis and also strongly suggest the reviewers to discuss more the data.\nAltogether, this a very interesting paper, really useful for the community in term of methodology, that deserves to be published. In contrast the new results leave one hungry and should be improved before publication.\nMajor comments\nLines 137-138: I do not understand why CPT would decrease EdU-Alexa488 signal since CPT is added after EdU and therefore should not impact its incorporation. Does it mean there is DNA resection?\nIn fact they do write at line 168 that the loss of EdU signal is caused by loss of fork protection. Authors need to homogenize here.\nIn principle EdU changes after EdU wash out can be a reflection of either, degradation of EdU or residual EdU incorporation. In our experience, this depends at least in part on the stalling agent used (and of course the genetic background of the cell). In the case of CPT experiments, we find that there is some residual EdU incorporation even after EdU wash out. This is reduced in the presence of CPT and consistent with published data showing fork slowing with CPT (e.g. PMCID: 4347635). We have added these data (Ext. Data Fig. 1d). We also now better discuss the importance of proper controls and careful interpretation of the data. Whether EdU changes are caused by degradation or residual incorporation, we show that either can be mitigated by shortening the time after stalling, e.g. 15 minute conditions, which simplifies the experimental systems and aids to the robustness of the analysis.\nIn several figures (1C, 1D, 2A etc.) it is written norm. int., I supposed it stands for normalized intensities but it should be described in the legend. According to the methods section, they did a normalization procedure in order to pool together 2-4 experiments, it makes sense since PLA intensities varies between experiments. But it would be nice to show systematically the repetitions in Supdata like they did for Fig1 and Fig 3.\nWe added the definition to the legends. We also describe the normalization method in the method section. We also added the Supdata (Ext. Data Fig. 2d-o).\nIn Fig4 the authors show clear data demonstrating that, as expected, SMARCAL1 depletion decreases the formation of RFs using RF-SIRF. I think it would nice to see if this is also the case for the other translocases (HLTF and ZRANB3) known to be involved in fork reversal. It would strengthen the results.\nWe now have added data with 3KO U2OS cells obtained from the Cortez lab, which contain knock outs of all three fork remodelers, SMARCAL1, ZRAN3B and HTLF. We show a strong reduction of RF-SIRF signals (Fig. 4f and Ext. Data Fig.4 j,k), a reduction of DNAPKcs-SIRF signals (Fig. 4 g,h and Ext. Data Fig.4 l,m) and H3K4me3:H3K9me3 PLA signals (Fig. 7 a,b, and Ext. Data Fig. 7b,c) in 3KO cells with replication stalling, but not H3-SIRF signals (Fig. 7 c,d and Ext. Data Fig. 7d,e).\nOne of the most important issue of the paper is that the authors does not really consider S-phase patterns in their analysis (Fig5), I think it could provide a more precise description of the results. It is well established that the EdU patterns are diffused in early-S (euchromatin), more condensed in mid-S (facultative heterochromatin) and very condensed in late S (constitutive heterochromatin replication). For instance, the ring in Fig5D could correspond to heterochromatin replication and not to a relocalization of the fork to the nuclear membrane, the patterns of EdU presented in this figure could actually correspond to mid-S in my opinion. Actually, they do write on line 266 that RF-SIRF signals have a preference for early/mid S, but on Fig5C the just compare early-S and late-S. At the end I am a little bit lost. To increase the clarity of the results, it would be determinant to clearly decipher if RF-SIRF is occurring more in early-S or in mid-S. To do so the authors need to re-analyze the RF-SIRF data using the replication patterns and present the RF-SIRF level in Mid-S vs Early-S cells, this would complement their study based on dapi level in a more precise way.\nThank you for this suggestion! We re-analyzed the cell cycle data according to EdU-Alexa488 patterns. The highest RF-SIRF signal cluster into early S-phase cells with diffuse EdU patterns, coinciding with ring-patterns of RF-SIRF signals (Fig. 5g,h and Ext. Data Fig. 5g). Importantly, we added representative images that demonstrate that the RF-SIRF ring pattern does not correlate with the Alexa488-EdU patterns in neither early nor mid S-phase cells (Fig. 5g,h). e.g. there is no or little RF-SIRF signal within the cell lumen in early S-phase cells despite that there is ample EdU-Alexa488 incorporation there. While ring patterns ensue into some mid-S-phase cells, the RF-SIRF signals remain peripheral irrespective of luminal EdU-Alexa488. This suggests that the RF-SIRF ring pattern is indicative of RF structures, rather than a reflection of EdU patterns of mid-S-phase cells. Of note, our data shows that RFs also form in late S-phase as RF-SIRF signals reduce with SMARCAL1 knockdown (Ext. Data Fig. 5h). However, in late S-phase RF-signals do not localize to the nuclear periphery but instead are exclusively within the cellular lumen, thus strikingly differ from early S-phase RFs.\nThe data from Fig6 regarding the increase level of H3K9me3/H3K4me3 and decrease level of H4K16ac during fork reversal induced by CPT are interesting. However, I do not really understand the mechanism. Does it mean that somehow these modified histones are recruited or excluded during fork reversal? But how? Or does it mean that the enzymes required (such as SUV39H1 for H3K9me3 or MLL3/4 for H3K4me3) for these modifications are recruited to reversed fork? I think the authors should discuss more their data and provide a little bit of mechanistic otherwise it remains really descriptive.\nOn line 293 they write: “these data explain how MRE11 is recruited there”. I think the data provided in the current version of this manuscript are not sufficient to support completely this claim. As cited by the reviewer (Ray Chauduri 2016), MLL3/4 is required to recruit MRE11 to reversed forks. It would great if the authors could test by SIRF the impact of MLL3/4 inactivation on MRE11 recruitment and on H3K4me3 recruitment in the conditions used in Fig6.\nWe now show that H3K4me3 at reversed forks are dependent on PTIP, which is a MLL3/4-binding partner required for H3K4 tri-methylation (added Fig 7 e,f). This validates the epigenetic changes at stalled and reversed forks as new events. Total H3-SIRF on the other hand does not increase at RFs (7c,d). This supports epigenetic modification changes at stalled forks. Our previous data showed that both H3K9me3 (classically transcription repressing) and H3K4me3 (transcription activating) increase at reversed replication forks. We now show that these are otherwise rare bi-valent marks are significantly increased at reversed forks, e.g. reduce with fork-reverser knock-outs (added Fig. 7a-b), further supporting that there are replication stress specific epigenetic marks.\n[REDACTED]\nMinor comments\nI would put the SupFig1A instead of 1A because it simplifies the life of the reader to understand the principle of the method directly.\nDone.\nOn line 187 the write that the strongest effect of RF is H2O2, in Zelwegger 2015 JCB they also found that H2O2 has one of the strongest effects (with CPT). I would be curious to know authors hypothesis to explain this\nThere are several possible explanations both technical (e.g. the number of possible 8-Oxo-G lesions caused by H2O2 may be greater than the number of possible TopI crosslinks caused by CPT which is limited by TopI protein amounts) and biological (8-OxoG may be a more impactful lesion with atmospheric oxygen changes during metazoan evolution being one of the most impactful evolutionary drivers for the development in cellular protection systems), but all of them are speculations. We nevertheless describe the phenomenon as a noticeable event.\nOn Fig3 and SupFig3 it is written that data are derived from3 independent repeats, why only 2 repeats are shown for SupFig3 c, d and e?\nThank you for noticing the omission, we added the third repeat to the Ext. Data Fig.3\nIn Fig5 the gradient colors are missing.\nAdded.\nReviewer #3 (Remarks to the Author):\nReplication fork reversal is a protective mechanism during DNA replication stress that occurs when replication forks encounter an obstacle and reverse their course, forming a four-way \"chicken foot\" structure through the annealing of newly synthesized DNA strands, while the parental strands re-anneal. This structure has gained significant interest over the past decade, as it stabilizes the replication fork, prevents DNA damage, and allows time for repair. As stated by the authors, direct analysis of these structures requires complex electron microscopy-based techniques, mastered by only a few specialized laboratories worldwide. In this article, Roy and colleagues propose an elegant new approach to monitor fork reversal using a modified proximity ligation assay (PLA) that specifically detects EdU on the regressed arm of the reversed fork. The assay is based on the bold assumption that EdU epitopes incorporated into the two sister chromatids of a normal Y-shaped fork do not generate a PLA signal, due to their physical separation—even though they are held together by cohesin. Nevertheless, the authors convincingly demonstrate a specific RF-SIRF signal at reversed forks, normalized to total EdU incorporation. They further show that this signal increases upon treatment with drugs known to induce fork reversal and is lost following depletion of SMARCAL1, a key factor involved in fork reversal. Additionally, they show that this assay provides insights into the subnuclear organization of reversed forks, the presence of single-stranded DNA (ssDNA) at reversed forks, and their relative frequency during S phase. Altogether, RF-SIRF emerges as a promising assay that complements indirect methods such as DNA fiber analysis and serves as an accessible alternative to electron microscopy. This approach fulfills a clear need in the field and will be of great utility to the growing number of laboratories investigating replication stress responses and fork reversal. However, the authors should address the following points to strengthen their conclusions:\n1. Simultaneous click chemistry using biotin-azide and Alexa488-azide may compete for the single alkyne group on EdU. This competition could significantly impact data interpretation and may explain the substantial variability observed in EdU-PLA signals across biological replicates, as shown in Extended Data Fig. 1.\nVariability in PLA experiments is a well-known phenomenon and intrinsic to PLA technology (see eg. PMC8041061), but not specific to RF SIRF or to EdU reactions.\nYes, Alexa488 and biotin azides do compete for EdU moieties, which is by design. The azides are applied at the same time, so are in steady state equilibrium and have the same relative chances for chemical addition within a given experiment (e.g. same chance between the conditions tested). Given that Click-it is a chemical reaction, there seems to be no logical reason to believe it could explain variability in EdU-PLA signals.\n2. In the legend of Extended Data Fig. 1a, the authors mention anti-biotin primary antibodies. These should presumably be derived from two different species (e.g., mouse and rabbit). The authors should clarify this in the Methods section.\nYes, we utilize two anti-biotin antibodies from two different species. We now better clarify this in the methods section.\n3. In the ssDNA-SIRF assay, what is the minimum ssDNA length required for effective recognition by the anti-ssDNA antibody?\nThe antibody is raised against denatured calf thymus, so the minimum ssDNA length is not defined. We are not able to make a ssDNA size implication using our technique.\nHowever, we added data with structure specific ssDNA nucleases. Both ExoT (3’-5’ssDNA exonuclease) and RecJ (5-3’ssDNA exonuclease), which cannot act on dsDNA, degrade the RF structures detected by ssDNA-SIRF. This shows that both 3’ and 5’ ssDNA exists at these sites (added Fig 3e and Ext Data Figs.3h,i)\n4. Could the authors perform a time-course experiment to assess whether RF-SIRF signals increase during genotoxic treatment in the presence of nuclease inhibitors such as Mirin?\nTo further address how reversed forks form, we added a time course experiment for RF-SIRF. This validates the idea that it is predominantly the nascent DNA prior to DNA damage that forms RF-SIRF signals (Ext. Data Fig. 1h-j)\nMRE11 in the context of fork reversal formation is currently being actively investigated by several group. Inhibition with mirin would be insufficient to address any role for MRE11 during fork reversal, and this merits a separate full study that goes beyond the scope of this manuscript. Moreover, time course experiments with MRE11 are likely to heavily change the EdU concentrations, which will render interpretations difficult. We added discussion about the challenge in interpretation and the importance in considering total amounts of EdU to the discussion.\n.\n5. The authors present convincing evidence that ssDNA can be detected at reversed forks upon resection. It would be valuable to demonstrate that the RF-SIRF signal decreases as ssDNA-SIRF increases. A time-course experiment in BRCA2-deficient cells could illustrate these signal dynamics.\nTo our knowledge, there is no evidence for reversed forks containing blunt ends. Rather by the nature of their formation, reversed forks contain ssDNA overhangs at least initially, which are either 3’ or 5’ depending on if the leading or lagging strand are ahead.\nTo this end, we have added data demonstrating this using structure specific ssDNA nucleases. Both ExoT (3’-5’ssDNA exonuclease) and RecJ (5-3’ssDNA exonuclease), which cannot act on dsDNA, degrade the RF structures detected by ssDNA-SIRF, showing that both 3’ and 5’ ssDNA exists at these sites (Fig 3e and Ext. Data Fig. 3g,h).\nWhile we believe that RF-SIRF is a key advance in studying fork reversal and reversed forks, like any technique, it has its limitations. In this case, the signals are directly dependent on the total EdU molarity in addition to proximity. Both nascent DNA degradation (as expected in BRCA2 defective cells) and residual incorporation during extended periods of time (e.g. see added Ext. Data. Fig 1d, showing continued residual incorporation of EdU) can cause changes in EdU moieties. This multi-valent EdU changes make any RF-SIRF interpretation for a time course experiments in BRCA2 deficient cells speculative. We now better discuss RF-SIRF limitations in the discussion.\n6. The EdU-H3K9me and H4K16ac-EdU PLA experiments do not presently add substantial value, as the authors do not directly demonstrate that the observed signal arises from reversed forks. Unless supported by further experimental validation, this section should be removed.\nWe have added multiple data figures (Fig. 7 and Ext. Data Fig. 7) to better demonstrate and clarify the advances and discoveries. Our initial data showed that both H3K9me3 (classically transcription repressing) and H3K4me3 (classically transcription activating) increase with fork reversal (e.g. decrease with SMARCAL1 knockdown). From a simple transcriptional epigenetic point of view, these results appear contradicting. However, we now furthermore show that H3K9me3 and H3K4me3 are co-occurring upon for reversal (Fig. 7a,b): this suggests distinct specificity for these otherwise rare bi-valent epigenetic marks at reversed forks. Thus, these data corroborate our interpretation of a replication specific epigenetic language that is distinct from the transcriptional epigenetic code.\nTotal H3-SIRF on the other hand slightly decreases rather than increases at reversed forks, suggesting active epigenetic changes at stalled forks (Fig. 7c,d and Ext. Data Fig. 7b,c). Consistently, we find that these epigenetic marks at reversed forks are dependent on PTIP, which is a MLL3/4-binding partner required for H3K4 tri-methylation, validating the epigenetic changes at stalled and reversed forks as new events (Fig. 7e,f and Ext. Data Fig. 7d,e)\nThe advance in this data is that our results highlight replication-stress specific epigenetic changes that previously were not appreciated despite the clear understanding that single specific epigenetic marks recruit or repress distinct DDR proteins. Our data establish specific epigenetic marks or codes at stressed and reversed replication forks. Despite the fact that they are classically incompatible from a transcriptional view angle, these findings can explain how replication stress response proteins are recruited to stalled forks in the face of opposing epigenetic changes from a transcriptional view point.\n[REDACTED]\n7. The word \"telomere\" is misspelled multiple times in the manuscript and should be corrected throughout.\nThank you. Changed.\n[REDACTED]"}, {"round": 2, "reviewer_comments": {"reviewer_1": "The authors have addressed my concerns. I have no further suggestions and I believe the manuscript can be accepted for publication.", "reviewer_2": "The authors of the manuscript did a considerable amount of work that greatly improved the manuscript. Some experiments such as the one with RecJ and ExoT are really nice and strengthen the approach. They positively answered my comments and I am glad to see that the suggestion I made regarding the S-phase turned out to be really useful. I think now the relocalization of reversed forks to the nuclear periphery is supported by convincing results and open new questions. Also, the idea of epigenetic code for fork reversal is almost more solid. Therefore, I think the manuscript fully deserved to be published in Nat Com.\nHaving said that, there are few minor modifications that need to be done before publication:\n-Ideally the number of the repetition (1-2-3) should be indicated in extended fig 1-2-3. For instance, in extended Fig 1, the number of the experiment should be indicated for EdU-PLA/EdU-Alexa488/RF-SIRF to really show that the variation is similar in each replicate (paired data).\n-In extended fig 1 h/i/j it is written CTP instead of CPT.\n-In Fig2h & I they should comment on the 2 populations of cells in the legend since they have to cut the Y-axis\n-I believe the legend of the Y-axis of Fig4 g is not correct\n-there is a problem in the legend of the Y-axis in Fig 5g", "reviewer_4": "The authors have conducted new controls and experiments that substantially strengthen the manuscript. They now include cell cycle analyses, EdU chase assays, and time-course experiments, which confirm that RF-SIRF occurs only at high EdU concentrations—explaining why it was absent in their original SIRF protocol.They further validate reversed fork detection using structure-specific ssDNA exonucleases and triple-KO U2OS cells (SMARCAL1, ZRANB3, HLTF). They also provide stronger evidence of co-occurring H3K9me3 and H3K4me3 at reversed forks, supporting the intriguing idea of a unique chromatin landscape at these structures. In my view, the manuscript is now suitable for publication and should be of broad interest to the field."}, "author_response": "REVIEWERS' COMMENTS\nPlease see reviewer remarks in italic, and our responses marked in blue font.\nReviewer #1 (Remarks to the Author):\nThe authors have addressed my concerns. I have no further suggestions and I believe the manuscript can be accepted for publication.\nThank you\nReviewer #2 (Remarks to the Author):\nThe authors of the manuscript did a considerable amount of work that greatly improved the manuscript. Some experiments such as the one with RecJ and ExoT are really nice and strengthen the approach. They positively answered my comments and I am glad to see that the suggestion I made regarding the S-phase turned out to be really useful. I think now the relocalization of reversed forks to the nuclear periphery is supported by convincing results and open new questions. Also, the idea of epigenetic code for fork reversal is almost more solid. Therefore, I think the manuscript fully deserved to be published in Nat Com.\nHaving said that, there are few minor modifications that need to be done before publication:\n-Ideally the number of the repetition (1-2-3) should be indicated in extended fig 1-2-3. For instance, in extended Fig 1, the number of the experiment should be indicated for EdU-PLA/EdU-Alexa488/RF-SIRF to really show that the variation is similar in each replicate (paired data).\nNow provided\n-In extended fig 1 h/i/j it is written CTP instead of CPT.\nChanged thanks\n-In Fig2h & I they should comment on the 2 populations of cells in the legend since they have to cut the Y-axis\nClarified in Figure legend (for better visualization)\n-I believe the legend of the Y-axis of Fig4 g is not correct\nChanged, thank you\n-there is a problem in the legend of the Y-axis in Fig 5g\nChanged, thank you\nReviewer #4 (Remarks to the Author):\nThe authors have conducted new controls and experiments that substantially strengthen the manuscript. They now include cell cycle analyses, EdU chase assays, and time-course experiments, which confirm that RF-SIRF occurs only at high EdU concentrations—explaining why it was absent in their original SIRF protocol.They further validate reversed fork detection using structure-specific ssDNA exonucleases and triple-KO U2OS cells (SMARCAL1, ZRANB3, HLTF). They also provide stronger evidence of co-occurring H3K9me3 and H3K4me3 at reversed forks, supporting the intriguing idea of a unique chromatin landscape at these structures. In my view, the manuscript is now suitable for publication and should be of broad interest to the field.\nThank you"}]

Decision letter