FANCD2 restrains fork progression and prevents fragility at early origins upon re-replication

Domain: neuroscience
Published: 2026-02-07
Editorial outcome: major_revision

Abstract
DNA replication is tightly regulated to ensure a single round of chromosome duplication per cell division. DNA licensing restricts origin firing to once-per-cell-cycle while aberrant licensing promotes re-replication and genome instability. Here, we investigate the mechanisms that protect genome integrity following re-replication induced by depletion of the licensing inhibitor Geminin. We find that re-replicating cells require FANCD2 to prevent genome instability. FANCD2 is rapidly recruited to chromatin upon Geminin loss, where it limits unrestrained fork progression and prevents single strand DNA gap accumulation and fork breakage. Genome-wide analyses reveal that upon re-replication, FANCD2 localizes to early origins within highly transcribed regions prone to accumulate R-loops and enriched in early replicating fragile sites. Importantly, reducing transcription and R-loops alleviates re-replication-induced genome fragility, whereas PARP inhibition exacerbates it. Our study uncovers a role for FANCD2 in safeguarding genome integrity during re-replication, offering avenues for selective targeting of cancer cells.

Introduction
Accurate DNA duplication is essential during somatic cell division to ensure the inheritance of a complete copy of parental DNA by the offspring. In eukaryotes, replication initiates from discrete genomic regions known as origins, which are spatially distributed along the chromosome and become activated in clusters that follow a defined temporal program to achieve the complete replication of the genome1,2. Replication initiation is mediated by two major steps that are temporarily spaced by the different activity of cyclin-dependent kinases: origin licensing (helicase loading) in G1 and origin firing (helicase activation) in S-phase. Origin licensing refers to the assembly of pre-replicative complexes (pre-RCs) containing the origin recognition complex (ORC), CDC6 and CDT1, and the inactive form of the mini-chromosome maintenance 2–7 (MCM) helicase onto each origin of replication 3–7, whereas origin firing entails the recruitment of additional replication factors for the conversion of pre-RCs into active replisomes8–10. Uncoupling between licensing and firing is crucial to ensure that no genomic region is left un-replicated while limiting single origin firing to once per cell cycle11,12. Among all the licensing proteins involved in pre-RC formation, CDT1 undergoes the greatest control mechanisms as it recruits soluble replicative helicases from the nucleus to ORC and CDC6-bound origins13,14. Consistently, earlier work showed that aberrant CDT1 activity induces re-replication and DNA breakage across species. Over-expression of CDT1 in fission yeast cells leads to re-replication and gene amplification15. Accordingly, addition of CDT1 to X. laevis eggs or over-expression of the ortholog of CDT1 in Drosophila is sufficient to induce re-replication and DNA damage16,17. In mammalians, ectopic expression of CDT1 promotes re-replication and tumorigenesis18–22, and inhibition of its ubiquitin-mediated proteolysis has demonstrated to induce apoptosis in several cancer models23,24. Besides transcriptional control of CDT1 expression and fine-tuning over its activity through the ubiquitin proteasome system25, timely regulation of CDT1 depends on Geminin, a small protein only present in metazoans that acts as an inhibitor of pre-RC assembly by direct binding to CDT1 in S and G226–29. Control over DNA licensing by Geminin is considered a redundant mechanism in metazoa to prevent any non-degraded and nuclear-free CDT1 from initiating illegitimate licensing of origins outside of G1. According to this, Geminin loss induces re-replication and DNA damage selectively in cancer cells30,31, which usually exhibit an abnormal expression of several licensing factors and compromised tumor barriers32. These results are consistent with analysis in vivo, where Geminin ablation has demonstrated to favor tumorigenesis and induces cell death in neural stem cells during development33,34. The dependency of highly proliferative cells on Geminin could be a genomic trait of therapeutic significance in cancer, and small molecules phenocopying the effects of inhibiting Geminin activity are being currently characterized with promising results35,36. The increasing evidence suggesting that re-replication is a driving force of tumorigenesis argues for a better understanding of the mechanisms detecting and preventing re-replication associated DNA lesions.

Here, we show that cells depleted of Geminin require FANCD2 to promote cell survival and prevent deleterious levels of DNA damage. FANCD2 is pivotal during repair of inter-strand crosslinks (ICLs)37, but recent studies have demonstrated that it also plays a fundamental role during replication stress by promoting fork stability38. Our results indicate that Geminin depletion induces FANCD2 recruitment onto chromatin early during the first round of replication, which is required to limit the unrestrained progression of re-replicating forks. Consequently, loss of FANCD2 in Geminin-deficient cells triggers the accumulation of single strand DNA (ssDNA) gaps and leads to massive fork breakage, which ultimately results in increased genomic instability. Genome-wide analysis of FANCD2 distribution on chromatin upon Geminin depletion demonstrates its enrichment at early DNA origins localized within highly transcribed genes, which are characterized by increased levels of transcription-replication conflicts (TRCs) and DNA damage. The results in this study reveal a function of FANCD2 in the maintenance of genome stability upon re-replication and provide insights into the clinical potential of targeting Geminin in FA/BRCA-deficient cancers.

Results
High-content screening identifies FANCD2 as essential to prevent DNA damage in Geminin-depleted cells
To identify mechanisms involved in the detection and repair of DNA lesions associated with aberrant licensing, we examined the effects of Geminin depletion on re-replication and DNA damage across different cell lines. We performed flow cytometry on Geminin-depleted and EdU pulse-labeled cells at different time points, which allowed us to measure the prevalence of cells with a DNA content greater than G2/M that were incorporating EdU as a metric for re-replication. As an alternative approach to quantify the levels of re-replication, we also measured nuclear areas39, while we analyzed γH2AX and 53BP1 signals as proxies of DNA double-strand breaks (DSBs). Depletion of Geminin in U2OS cancer cells resulted in an initial accumulation of cells in S phase within the first 24 h, with a small proportion undergoing re-replication (>4C DNA content) (Supplementary Fig. 1a). A high prevalence of re-replicating cells was observed by 48 hs, at which point approximately 30% of cells showed >4C DNA content and active EdU incorporation (Supplementary Fig. 1a). These results were also reproduced by measuring nuclear areas (Supplementary Fig. 1b) and were also accompanied by a significant increase in γH2AX and 53BP1 signals (Supplementary Fig. 1c). Co-depletion of CDT1 partially suppressed these phenotypes (Supplementary Fig. 1d), indicating that these effects are dependent on active DNA replication. In contrast, depletion of Geminin in non-cancerous hTERT-RPE1 cells had minimal effects on re-replication and DNA damage (Supplementary Fig. 1e-g), supporting the notion that Geminin prevents re-replication selectively in cancer cells30,31. Accordingly, Geminin depletion increased DNA damage in MCF7 cancer cells but had a minor effect in the non-tumorigenic MCF10A cell line (Supplementary Fig. 1h).

The mild effects observed in hTERT-RPE1 cells 48 h after Geminin depletion provided an ideal system to achieve low-level re-replication and prompted us to employ this cell line to perform a high-content screening for factors required to maintain genome stability in cells primed for re-replication. To that end, we used a custom-designed siRNA library targeting 300 genes associated with the DNA damage response (DDR) and DNA repair pathways (Supplementary Data 1). After transient transfection with the siRNA library, control and Geminin-depleted hTERT-RPE1 cells were pulse-labelled with EdU for 1 h prior to fixation, and total cell counts, as well as EdU and γH2AX signal intensities, were measured as markers of cell viability and DNA damage (Fig. 1a).

The results of the screen yielded two distinct classes of candidates, consisting of genes whose depletion was toxic to both cell lines (Geminin-depleted and control), which served as a control to validate the methodology of the screening, and a second class, comprising genes whose depletion was particularly toxic to Geminin-depleted cells, which were identified as hits. As expected, among the top-scoring genes whose downregulation increased DNA damage and compromised survival of both Geminin-depleted and control cells, we identified critical regulators of the cell cycle and cell cycle-checkpoint pathways40–42 (i.e. PLK1, WEE1 or CHK1) (Fig. 1b and Supplementary Data 2). Concentrating on the candidates whose downregulation increased DNA damage or interfered with survival specifically in Geminin-depleted cells, we identified BRWD3, a histone reader and substrate of the Cullin 4-DDB1 E3 ubiquitin ligase complex43, and the pro-survival and BCL2-related protein BCL2L244 as top hits (Fig. 1b and Supplementary Data 2). Importantly, among the top candidates increasing DNA damage specifically in Geminin-depleted cells, we also found FANCD2 (Fig. 1b and Supplementary Data 2), a central component of the FA pathway traditionally associated with repair of ICLs37. Interestingly, other FA signaling components which cooperate with FANCD2 during ICL repair (such as BRCA1/2 or ERCC1) did not score positive in our screening (Supplementary Data 2), suggesting that the role of FANCD2 during re-replication is independent from its canonical ICL repair function. These results, together with the growing evidence supporting a critical role for FANCD2 in maintaining replication fork stability38, led us to prioritize this candidate for further analyses. Thus, we validated our results from the screening with a different set of siRNAs, which confirmed increased γH2AX intensities and 53BP1 foci (Fig. 1c) and revealed a significant reduction in long-term cell survival, as measured by clonogenic assays (Fig. 1d), in double Geminin and FANCD2 knockdowns when compared to control or single-depleted cells.

A similar effect was observed in cancer cells with high levels of re-replication. Specifically, this was confirmed in U2OS cells in which Geminin was depleted by siRNA or through a doxycycline-inducible shRNA (Supplementary Fig. 2a, b, and Supplementary Fig. 3a, b), as well as in HCT116 cells containing an auxin-inducible degron (AID)45 for total inactivation of Geminin (Supplementary Fig. 3c). Of note, co-depletion of Geminin and FANCD2 impaired long-term clonogenic survival but did not induce early apoptotic cell death, as demonstrated by analysis of Annexin V+-PI and cleaved Caspase-3 (Supplementary Fig. 2c, d). These results are in line with the limited effects observed in cell counts during the screen (Fig. 1b and Supplementary Data 2) and exclude the possibility that the effects are due to acute cytotoxicity.

Additionally, these results were further supported by assessing the effect of Geminin depletion in FANCD2-KO PD20 cells derived from a FA-D2 patient. As expected, silencing of Geminin induced significant levels of DSBs and reduced cell viability in PD20 cells (Supplementary Fig. 3d, e). Importantly, introduction of the corrected wild-type (WT) FANCD2 rescued these phenotypes to basal levels (Supplementary Fig. 3d, e), altogether suggesting that Geminin-depleted cells require FANCD2 to limit extensive DNA damage and promote cell survival.

FANCD2 is recruited at re-replicating forks prior to massive DNA breakage
To explore the potential contribution of FANCD2 in preventing DSBs during re-replication, we initially examined its localization on chromatin. Subcellular fractionation of U2OS cells depleted of Geminin revealed increased FANCD2 recruitment to chromatin (Fig. 2a). Consistently, Geminin depletion also induced the formation of FANCD2 foci, which localized near γH2AX sites (Fig. 2b). To determine whether FANCD2 activation is a general response to re-replication, we also analyzed the formation of FANCD2 foci following mild over-expression of CDT1. Since transient overexpression of this protein induces massive re-replication in cancer cells46, we generated a stable clone expressing a CDT1-GFP fusion integrated into the genome of hTERT-RPE1 cells. This clone exhibited cell cycle-regulated CDT1 expression at levels 2-3 times higher than endogenous CDT1, sufficient to induce DNA damage without massively altering re-replication, as demonstrated by analysis of γH2AX intensities and nuclear areas (Supplementary Fig. 4a, b). As expected, over-expression of CDT1 in hTERT-RPE1 cells led to an increase in the proportion of cells with FANCD2 foci when compared to GFP-NLS control cells (Supplementary Fig. 4b), suggesting that FANCD2 is actively recruited to chromatin upon re-replication. Moreover, analysis of publicly available (TCGA) data (https://www.cbioportal.org) revealed an upregulation of FANCD2 in subgroups of bladder, uterine and breast cancers with high CDT1 expression (Supplementary Fig. 4c), consistent with an association of FANCD2 with aberrant DNA licensing.

Given that FANCD2 recruitment on chromatin is considered a consequence of replication fork blockage47–50, and our results in U2OS cells demonstrating an initial accumulation of cells in S prior to high levels of re-replication (Supplementary Fig. 1a), we hypothesized that Geminin depletion could trigger FANCD2 recruitment at stalled forks prior to the formation of widespread DSBs. To test this hypothesis, we measured the presence of FANCD2 foci and the increase in γH2AX nuclear intensities in Geminin-depleted U2OS cells in a time-course experiment. FANCD2 foci were observed as early as 12 h after siRNA transfection whereas γH2AX did not significantly accumulate until 48 h (Fig. 2c). Consistent with the analysis of FANCD2 foci, we did not observe a significant increase in the nuclear area until 12 h after transfection (Supplementary Fig. 5a), suggesting that upon Geminin depletion, FANCD2 recruitment on chromatin occurs early upon re-replication and precedes DNA breakage.

To further explore the dynamics of FANCD2 recruitment during re-replication, we performed a synchronization experiment in Geminin-depleted U2OS cells. Cells were arrested at G2/M by a sequential block with thymidine and the CDK1 inhibitor RO-3306, depleted of Geminin and then released into the next cell cycle (Supplementary Fig. 5b). To follow FANCD2 recruitment as cell progressed through S and G2 phases, cells were pulse-labelled with EdU and immunostained against FANCD2, together with the replication stress marker RPA32, and the G2/M marker pH3 (H3-Ser10Pho). While control cells showed the characteristic early, middle and late S-phase EdU patterns as cells progressed through S-phase, in Geminin-depleted cells a population with a particular diffused EdU pattern was evident from 14 h post-release, termed “R” from re-replicating (Fig. 2d). This cell population, which was actively synthesizing DNA, increased progressively and accumulated at the G2/M boundary, as shown by EdU incorporation in pH3-positive cells (Supplementary Fig. 5c). In parallel to the appearance of re-replicating cells, FANCD2 and RPA32 foci increased significantly in Geminin-depleted cells from 14 h post-release and accumulated at later time-points (Fig. 2e). Importantly, FANCD2 and RPA32 foci accumulation was evident specifically in re-replicating cells, as judged by the EdU pattern, and persisted as cells accumulated at G2/M (Supplementary Fig. 5c, d), indicating that re-replication upon Geminin loss begins within the first round of DNA synthesis and leads to early FANCD2 recruitment and RPA32 accumulation.

Next, to investigate whether FANCD2 is actively recruited at the vicinity of re-replicating forks, we pulse-labelled Geminin-depleted cells with EdU and monitored FANCD2 interaction with the newly synthesized DNA by proximity ligation assays (PLA). Geminin depletion significantly increased the number of EdU-FANCD2 PLA foci when compared to control cells (Fig. 3a). These results were also reproduced when we quantified the number of interacting PLA foci between FANCD2 and other replisome factors such as MCM2 or PCNA (Fig. 3b and Supplementary Fig. 5e). Notably, the interaction between the replisome and FANCD2 not only occurred in S-phase cells but became evident in those cells exhibiting the characteristic EdU incorporation pattern of re-replicating cells (Fig. 3b). Altogether, these results suggest that re-replication upon Geminin loss begins within the first round of DNA synthesis and induces FANCD2 recruitment to active forks prior to G2/M arrest and high levels of DNA damage.

FANCD2 limits fork progression in Geminin-depleted cells upon checkpoint activation
Given its accumulation at re-replicating forks, we next sought to determine whether FANCD2 was required to limit the extent of re-replication. Interestingly, co-depletion of Geminin and FANCD2 in U2OS cells did not increase the percentage of EdU positive cells with a DNA content >4C nor the nuclear size above that of Geminin-depleted cells, but on the contrary slightly reduced it (Fig. 3c, d). These results were also reproduced in hTERT-RPE1 cells (Supplementary Fig. 6a), excluding the possibility that FANCD2 restricts re-replication upon Geminin-depletion.

To further explore the dynamics of replication in cells depleted of Geminin and FANCD2, we then analyzed the progression of forks by DNA fiber assays. Double Geminin and FANCD2-depleted cells significantly increased fiber tract length and fork asymmetry (Fig. 3e), suggesting that FANCD2 limits fork speed and prevents fork stalling in the absence of Geminin. Although we could not measure origin distances due to DNA fiber analysis limitations, we observed that Geminin-depleted cells showed a slight increase in origin firing (Fig. 3f), according to the idea that over-licensing can also lead to ectopic origin firing32. However, downregulation of FANCD2 did not significantly alter the percentage of origin initiation (Fig. 3f), arguing against a primarily role for this protein in preventing ectopic firing of origins in the context of re-replication. Additionally, we investigated the effect of co-depleting Geminin and FANCD2 on cell cycle progression, using EdU incorporation and pH3 as markers of S and G2/M phases, respectively. According to the results obtained during synchronization experiments, Geminin depletion led to an increased proportion of cells retaining active DNA synthesis in G2, while they also exhibited high levels of DNA damage (Fig. 3g and Supplementary Fig. 6b). Geminin-depleted cells also demonstrated increased levels of phosphorylated Chk1 (Fig. 3h), in agreement with re-replication triggering checkpoint activation51–53. Indeed, inhibition of the G2/M checkpoint by incubating Geminin-depleted cells with a low dose of the VE-821 ATR inhibitor or with the general checkpoint inhibitor caffeine suppressed re-replication and the associated DNA damage phenotypes (Supplementary Fig. 6c, d). This suggests that overriding the G2/M checkpoint forces Geminin-depleted cells into mitosis prematurely, leaving insufficient time for extensive re-replication to occur. Consistent with this, incubation of Geminin-depleted cells with VE-821 resulted in the accumulation of inherited DNA lesions, as shown by increased micronuclei, aberrant mitosis, and reduced cell survival (Supplementary Fig. 6e, f). Importantly, co-depletion of Geminin and FANCD2 further increased the percentage of G2-arrested and EdU-positive cells, while it retained checkpoint activity (Fig. 3g, h), suggesting that FANCD2 is not required for checkpoint activation in response to re-replication. Collectively, these results indicate that depletion of FANCD2 accelerates fork speed and causes DNA damage upon checkpoint activation in Geminin-depleted cells.

Loss of FANCD2 promotes the accumulation of ssDNA gaps and genomic instability in Geminin-depleted cells
Unrestrained fork progression promotes the accumulation of ssDNA gaps54,55. These gaps may lead to excessive RPA loading and subsequent depletion of the available RPA pool, leading to fork breakage and genome instability56–58. To determine whether the abnormal progression of forks in double Geminin and FANCD2-depleted cells correlated with an accumulation of ssDNA gaps, we analyzed the formation of RPA foci. Depletion of Geminin and FANCD2 triggered the accumulation of RPA foci exclusively in re-replicating cells, as shown by colocalization of pre-extracted RPA32 and pH3 positive cells (Fig. 4a and Supplementary Fig. 7a). Notably, the intensity of RPA foci was also significantly increased in double-depleted cells (Supplementary Fig. 7b), likely due to longer ssDNA gaps. As an alternative way to visualize ssDNA, we also labeled the cells with the nucleotide analogue BrdU, which was then detected by immunofluorescence under native conditions. Consistent with the results observed during analysis of RPA32 foci, Geminin depletion induced the formation of BrdU foci while the proportion of cells exhibiting BrdU foci was further increased upon co-depletion with FANCD2 (Fig. 4b). To further confirm the presence of ssDNA gaps, we performed a DNA fiber assay in cells depleted of Geminin and FANCD2 and incubated with the S1 endonuclease. Labelled nascent DNA tracks were shorter and thus sensitive to the S1 enzyme in double Geminin and FANCD2-depleted cells compared to single-depleted and control cells (Fig. 4c). The shortening in tract length observed upon incubation of Geminin and FANCD2-double depleted cells to S1 nuclease was similar to that observed in cells incubated with the PARP inhibitor Olaparib (Fig. 4d). This is consistent with previous findings showing that PARPi sensitivity correlates with fork acceleration and gap formation58,59. Indeed, cells depleted of Geminin slightly increased the levels of parylation (Fig. 4d), whereas PARP inhibition in Geminin-depleted cells induced sensitivity to S1 nuclease activity and accumulation of RPA32 and BrdU foci, together with increased DSBs and reduced cell viability (Fig. 4e and Supplementary Fig. 7c-e). This reinforces the idea that an excess of ssDNA gaps underlies the high levels of genomic instability observed in re-replicating cells upon Geminin loss.

Finally, we tested whether increased formation of ssDNA gaps and accumulation of DNA damage in Geminin and FANCD2 double-depleted cells would result in chromosome breakage and high levels of genomic instability. As anticipated, metaphase spreads from Geminin-depleted cells already exhibited an increased number of chromosome breaks and aberrations (Fig. 4f). However, depletion of FANCD2 in Geminin-defective cells further exacerbated these phenotypes (Fig. 4f). Additional signatures of persistent genomic instability, such as 53BP1 nuclear bodies or micronuclei were also increased in Geminin and FANCD2 double-depleted cells across different cell lines (Fig. 4g and Supplementary Fig. 7f, g). Altogether, the results suggest that FANCD2 prevents the accumulation of ssDNA gaps and deleterious levels of DNA damage upon Geminin loss.

Contribution of FANCD2 mono-ubiquitination to cell survival during re-replication
The role of FANCD2 during replisome surveillance occurs independently of its mono-ubiquitination by the FA core complex, while it is required to preserve fork stability upon acute replication stress38,60. To explore the requirement of FANCD2 mono-ubiquitination for its function in sustaining the survival of re-replicating cells, we depleted FANCA - a key subunit of the E3 ubiquitin-ligase FA core complex - in Geminin-depleted cells. Consistent with the results obtained during chromatin fractionation, depletion of Geminin induced mono-ubiquitination of FANCD2, while depletion of FANCA reduced total FANCD2 levels (Fig. 5a), as previously described54. Our analyses also revealed increased DSBs (Fig. 5b) and accumulation of RPA foci (Fig. 5c) in double Geminin and FANCA-depleted cells in comparison to single Geminin-depleted cells. Moreover, clonogenic assays showed reduced survival of Geminin-deficient cells upon FANCA depletion (Fig. 5d), although this effect was less pronounced than that observed upon FANCD2 loss. Consistent with this partial effect, FANCD2 foci, although significantly reduced, could still be detected on chromatin in Geminin and FANCA double-depleted cells (Fig. 5e).

To further explore the contribution on FANCD2 mono-ubiquitination for survival of re-replicating cells, we evaluated the clonogenic capacity of FANCD2-KO U2OS cells reconstituted with either FANCD2-WT or the ubiquitination-resistant FANCD2-K561R mutant to Geminin depletion (Fig. 5f). As anticipated, depletion of Geminin in FANCD2-KO cells induced massive cell death (Fig. 5g) in respect to control cells. Interestingly, while introduction of FANCD2-WT significantly improved cell viability in Geminin-depleted cells, depletion of Geminin in cells expressing the FANCD2-K561R mutant showed an intermediate clonogenic outgrowth (Fig. 5g). These results are consistent with residual levels of chromatin-bound, non-ubiquitinated FANCD2 partially rescuing replication fork restart and cell proliferation upon replication stress60–62. Thus, our data indicate that FANCD2 mono-ubiquitination contributes to preserve cell survival upon re-replication and suggest an additional ubiquitination-independent role protecting genome integrity.

FANCD2 limits R-loop-associated fragility in Geminin-depleted cells
Intragenic origins mapping within highly transcribed genes have been shown to fire early during S phase upon oncogene-induced replication stress72. Furthermore, early replicating regions have been associated with transcriptionally active sites and accessible chromatin configuration73,74. Thus, we wondered whether Geminin depletion leads to fork stalling during early replication. Based on the analysis of publicly available Repli-seq datasets from the ENCODE project, which map replication timing across individual S subphases at specific genomic sites75, we observe a significant increase in BrdU intensity during early S phase, and a reduction in mid-to-late replication timing in those origins identified to accumulate FANCD2 specifically in Geminin-depleted cells when compared to control cells (Fig. 6f), suggesting these origins replicate early.

Transcriptional activity at early replicating regions has been linked to DNA damage and DNA-RNA hybrids at early replicating fragile sites (ERFSs)76,77. Thus, we analyzed the levels of DNA damage and DNA-RNA hybrids from available γH2AX ChIP-seq78 and DRIPc-seq79 data of wild-type K562 cells. γH2AX signal was higher in those genes specifically enriched in FANCD2 upon Geminin loss (Fig. 7a). In addition, and despite DNA-RNA hybrid distribution profiles were similar, these genes exhibited higher DRIPc-seq signal levels (Fig. 7b). These results suggest FANCD2 accumulation upon Geminin depletion occurs at R-loop containing regions that are prone to break even under unperturbed conditions, suggesting they could correspond to ERFSs. By lifting over ERFS data from synchronized mouse B cells76 employing the LiftOver (UCSC) alignment tool, we analyzed FANCD2 coverage over potential human ERFSs. Strikingly, metagenomic analyses showed a significant accumulation of FANCD2 at ERFSs in Geminin-depleted cells (Fig. 7c). Altogether, these results suggest that re-replication upon Geminin loss leads to the expression of ERFSs, supporting a model in which active transcription and R-loops cause fork stalling close to early replicating regions, thus contributing to genome instability. To test this hypothesis, we used the adenosine analog Cordycepin and we over-expressed RNaseH1 to examine the effect of inhibiting general transcription or degrading DNA-RNA hybrids in replication fork stalling (FANCD2 foci), ssDNA gaps (RPA foci) and DSBs (γH2AX foci) upon Geminin depletion. Cordycepin treatment effectively reduced EU incorporation (Supplementary Fig. 8f), supporting its role as a general transcription inhibitor. However, we were able to observe that it also caused a reduction in EdU incorporation (Supplementary Fig. 8g), which could partially impact S-phase progression. This was not the case for cells over-expressing RNaseH1, where bulk EdU incorporation remained unaffected (Supplementary Fig. 8g). Importantly, both treatment with Cordycepin and RNaseH1 over-expression significantly reduced the number of γH2AX, FANCD2 and RPA foci of Geminin-depleted cells (Fig. 7d, e). These results support that aberrant origin licensing and re-replication upon Geminin loss promote fragility of ERFSs, which is prevented by FANCD2 recruitment.

Methods
Cell lines and drug treatment
All cell lines were grown at 37°C under standard cell culture conditions (humidified atmosphere, 5% CO2) and regularly tested for the detection of potential contamination with mycoplasma. U2OS, MCF7, 293T and GMNN mAID HCT116 cells were incubated in Dulbecco’s modified Eagle’s medium (DMEM) (Gibco™, 11962) supplemented with 10% heat inactivated fetal bovine serum (FBS). Human hTERT-RPE1 cells were grown in DMEM-F12 (Sigma-Aldrich, D8437), supplemented with 10% FBS. MCF10A cells were grown in DMEM-F12 supplemented with 5% horse serum, 20ng/ml EGF, 0.5μg/ml hydrocortisone, 10μg/ml insulin and 100ng/ml choleratoxin. K562, PD20 FANCD2-KO and PD20 FANCD2-WT cells were cultured in RPMI (Gibco™, 11875093) supplemented with 10% FBS. The following compounds were used in this manuscript at the indicated final concentrations unless stated otherwise: Hydroxyurea (1mM, Sigma-Aldrich, H8627), Caffeine (5mM, Sigma-Aldrich, C0750), VE-821 (ATRi, 2 μΜ, Sigma-Aldrich, SML1415), Olaparib (PARPi, 0,5 μΜ, Selleckchem, S1060), Aphidicolin (0,5μM, Sigma-Aldrich, A0781), Nocodazole (100ng/ml, Sigma-Aldrich, M1404), Cordycepin (100μM, Sigma-Aldrich C3394).

Generation of stable CDT1-GFP hTERT-RPE1 cells
To generate a CDT1-GFP stable cell line, hTERT-RPE1 cells were infected with lentiviral particles expressing GFP-NLS or CDT1-GFP. The second-generation lentiviral packaging plasmid mix was used for lentivirus production as previously described108. Briefly, 293T cells were co-transfected using Turbofect (Fermentas) with the expression vector carrying either GFP-NLS (pLVDest-GFP) or CDT1-GFP (pLVDest-CDT1-GFP) and the plasmids encoding for the structural proteins of the virus (psPAX2, Addgene 12260) and the proteins of the viral envelope (pMD2.G, Addgene 12259). The supernatant was harvested 48 h after transfection, filtered and used to infect hTERT-RPE1 cells with 5μg/mL of hexadimethrine bromide (polybrene, Sigma-Aldrich). Limiting dilution and single colony picking were used to generate stable cell lines.

Generation of inducible Geminin knockdown cell lines
To generate a U2OS cell line with doxycycline-inducible depletion of Geminin, shRNA sequences targeting human Geminin (GMNN) were cloned into the Tet-pLKO-puro “all-in-one” lentiviral system for tetracycline-inducible shRNA expression (Addgene #21915). shRNA sequences were selected from the RNAi Consortium collection (MISSION®, Sigma-Aldrich) and synthesized oligonucleotides for each shRNA were annealed and ligated into the Tet-pLKO-puro vector following enzimatic digestion followed by transformation into competent bacteria. Lentiviral particles were produced by co-transfecting HEK-293T cells with the individual shRNA-containing Tet-pLKO-puro plasmids, along with the packaging plasmids psPAX2 (Addgene #12260) and pMD2.G (Addgene #12259), using a suitable transfection reagent. Viral supernatant was collected 48 h post-transfection, filtered through a 0.45 μm filter, and stored at −80°C. To generate stable cell lines, U2OS cells were transduced with the lentiviral particles in the presence of 5 μg/ml Polybrene for 48 h before selection and maintainance in media containing 2 μg/ml puromycin and Tet-System Approved FBS. To induce Geminin knockdown, stable cells were treated with 1 μg/ml doxycycline for 48 h.

Generation of mAID-tagged HCT116 cells
For the generation of the HCT116-Gem-mAID auxin inducible degron cells, the mAID tag was added in the C-terminal of the GMNN protein. Guide RNAs targeting the 3’ UTR of the GMNN (Geminin) gene were designed with Bbs1/Bpi1 overhangs and ligated into pX330-U6-Chimeric_BB-CBh-hSpCas9 (#42230, Addgene). Homology directed repair templates were assembled using the GeneArt™ Gibson Assembly HiFi Master Mix (Invitrogen): ~900 bp homology arms flanking the GMNN 3’ UTR overwriting the stop codon, either mAID-mClover-Hygro or mAID-mCherry-NeoR pieces and the respective backbones cut by BamHI-HF from plasmids #72828 and #72830 (Addgene), respectively. Plasmids expressing the Cas9 and the targeted gRNAs and plasmids expressing the homology arms and the mAID cassettes were transfected into HCT116-CMV-OsTIR1 cells using FuGENE-HD (Promega) according to manufacturer’s instructions. 24 h after transfection, cells were reseeded in low density and grown in antibiotics (Hygromycin 100 μg/ml, G418 700 μg/ml, and Puromycin 2 μg/ml) for >10 days for colonies formation. Single colonies were selected and propagated independently. Correct clones were confirmed for the presence of the mAID cassette, the fluorescent markers, and the expression of the tagged protein by PCR, immunofluorescence microscopy and immunoblotting (in-house developed Geminin-specific antibodies).

Immunostaining
Immunofluorescence experiments were performed 72 h after siRNA or plasmid transfection, unless otherwise indicated in the experiment. Optionally, for the detection of proteins closely bound to chromatin, cells were pre-extracted with 0,2% Triton X-100 in PBS for 2 min at RT. Cells were fixed in 4% paraformaldehyde (PFA) solution for 10 min at RT, washed three times in PBS, permeabilized with 0,3% Triton X-100 for 5 min at RT and washed three times in PBS. Cells were then incubated with a blocking buffer containing PBS, 10% FBS and 3% BSA for 1 h at RT, before incubation with primary antibodies for 16 h at 4°C in a humid chamber. After washed with PBS-0,1% Tween, coverslips were incubated with secondary antibodies for 1 h at 25°C in a dark and humid chamber. After three subsequent washes with PBS-0,1% Tween, coverslips were incubated with Hoechst 33342 for 15 min at RT for nuclei staining. Finally, cells were washed with PBS and mounted with MOWIOL® 4–88. (Sigma-Aldrich). A list of the antibodies used in this study can be found in Supplementary Data 1.

EdU labeling
5-Ethynyl-20-deoxyuridine (EdU, Invitrogen, Cat#C10340) labelling was performed using BaseClick EdU-HTS 488 kits (Sigma-Aldrich, Cat#BCK-HTS488-2). First, cells were cultured in complete medium supplemented with 10 μM EdU for 30 min at 37°C. Optionally, to detect proteins closely bound to chromatin, cells were treated with a pre-extraction solution (0,2% Triton X-100) for 2 min at RT. Cells were then fixed by incubation with a 4% paraformaldehyde (PFA) solution for 10 min at RT. After that, cells were washed for 3 subsequent times with 1x PBS for 5 min each and then were permeabilized by incubation with a 0,3% Triton X-100 solution for 5 min at RT. At this point, detection of EdU was performed on the coverslips according to manufacturer’s instructions.

Detection of nascent ssDNA by native BrdU assay
To detect nascent ssDNA, native BrdU assay was performed as previously described109. Briefly, U2OS cells in coverslips, were pulse-labeled with 10 μM BrdU (Sigma-Aldrich, B5002) for 24 h. After washing with PBS, cells were permeabilized with freshly made extraction buffer (10 mM PIPES, 100 mM NaCl, 300 mM sucrose, 1.5 mM MgCl2 and 0.5% Triton X-100) for 2 min on ice. Subsequently, cells were fixed with 4% PFA for 10 min on ice and washed 3 times in PBS. Cells were then incubated with a blocking buffer containing PBS and 10% FBS for 1 h at RT, before incubation with mouse monoclonal anti-BrdU (B44 clone) (BD Biosciences, 347580, 1:50) for 16 h at 4°C in a humid chamber. After washing with PBS, coverslips were incubated with goat anti-mouse secondary antibody (Life Technologies, A11031) for 1 h at 25°C in a dark and humid chamber. After three subsequent washes with PBS, coverslips were incubated with Hoechst 33342 for 15 min at RT for nuclei staining. Finally, cells were washed with PBS and mounted with MOWIOL® 4–88 (Sigma-Aldrich).

Cell synchronization for monitoring DNA synthesis along S and G2
For synchronization of U2OS cells to monitor DNA synthesis along S and G2, cells were plated onto coverslips and incubated with 2 mM of thymidine (Sigma-Aldrich, T9250) for 20 hours, then washed 2 times with 1x PBS and released in normal media for an additional period of 6 h. During the release period, cells were transfected with a siRNA oligo targeting Geminin or against a control sequence, as mentioned above. 5 h after transfection, the medium was removed, and cells were incubated with 10 μΜ of CDK1 inhibitor RO-3306 (Sigma-Aldrich, SML0569) for 8 h. Cells were then washed 2 times with 1x PBS and were subsequently released for 12 h prior to collection of time points every 2 h for a total time-course of 30 h. 30 min before each timepoint, cells were incubated with 10 μΜ EdU (Invitrogen, Cat#C10340) to monitor replication. Fixed cells were then permeabilized and immunofluorescence analyses coupled with EdU detection were performed, as mentioned above.

PLA assay
PLA was performed using Duolink PLA Technology (Merck). Cells were incubated with 25 mM EdU for 15 min. Samples were then incubated with 0.1% formaldehyde in PBS for 5 min, and pre-extracted in CSK buffer (10 mM Pipes pH 7, 0.1 M NaCl, 0.3 M sucrose and 3 mM MgCl2), prior to fixation. Click Reaction (100 mM Tris-HCl pH 8, 100 mM CuSO4, 20 mg/mL sodium-L-ascorbate and 10 mM azide-biotin) was performed according to the manufacturer’s instructions for 30 min at 37°C. Duolink Blocking Solution was replaced by 5% BSA, 10% Donkey serum in PBS. First and secondary antibody binding, ligation and amplification reactions were performed according to the manufacturer’s instructions. The PLA reaction was performed using the following reagents: PCNA (PC10, sc-56, Santa Cruz), MCM2 (D7G11, 3619, Cell Signaling), FANCD2 (NB100-182, Novus Biologicals), anti-Biotin (200-002-211, Jackson ImmunoResearch), Duolink in situ PLA probe anti-rabbit PLUS (Merck, DUO92002), Duolink in situ PLA probe anti-mouse MINUS (Merck, DUO92004) and Duolink-Detection Reagents Red (Merck). Finally, nuclei were stained with DAPI and mounted with MOWIOL® 4–88. (Sigma-Aldrich). Antibodies were used at 1:500 dilution. PLA foci were automatically quantified using Metamorph v7.5.1.0 software (Molecular Probes).

Flow cytometry
For 2D EdU/PI flow cytometry, cells were pulse-labeled with 10 μM EdU for 30 min before harvest. Afterwards, cells were trypsinized, washed in PBS, resuspended in 70% ice-cold ethanol and optionally stored at 4°C. Cells were then washed with PBS 1% BSA and incubated with 0.5% Triton X-100 in PBS for 10 min. EdU staining was performed using the BaseClick EdU-HTS 488 kit (Sigma-Aldrich, BCK-HTS488-2) according to the manufacturer’s instructions. The genomic DNA was stained with propidium iodide (Biotium, 40017) in combination with RNAse (AppliChem, A2760). For Annexin V/PI flow cytometry Annexin V Apoptosis Detection Kit (BD Pharmingen™, 556547) was used, according to manufacturer’s instructions. A BD FACSCantoTM Flow Cytometry System with FACSDiva software was used and FlowJo 10 was used for cell cycle analysis.

siRNA transfections
Transient gene depletions were carried out using the Lipofectamine RNAiMax transfection reagent, according to the manufacturer’s instructions, and a final concentration of 20 nM, unless stated otherwise. Silencer select negative control #1 siRNA was used as a control siRNA at the same concentration of the most concentrated siRNA used in the same experiment. For each experiment, cells were plated at approximately 70-80% confluency in 35 mm dishes. Cells were incubated with the mixture of Opti-MEM, RNAiMax and siRNA for 5 h at 37oC and then the transfection medium was removed and full DMEM 10% FBS was added to cells. Control over transfection efficiency was monitored 24–48 h post-transfection by Western Blot (WB) analysis. The following Silencer Select siRNAs were used: siGMNN (s27306), siFANCD2 (s533670), siFANCA (s528717), siCDT1 (s37722).

siRNA-based high content screening
Phenotypic image-based siRNA screens were performed in 384-well format as previously described110. Briefly, a custom designed siRNA library consisting of three individual siRNAs targeting 300 DNA repair genes (Silencer Select, Thermo Fisher Scientific, Supplementary Data 1) was pre-printed into 384 wells. From stock siRNA libraries, in which every well contained a single-targeting siRNA oligo at a concentration of 0.16 μM, 5 μl per well were transferred onto screening plates with the use of a multichannel pipette, in which a mixture of 5 μl of Opti-MEM and 0.09 μl of RNAiMAX was then added. Lipid mixture was incubated with the siRNAs for 20 min at RT and 40 μl of cells in suspension at a concentration of 1×103 cells per well were then added to the lipid mixture to reach a final concentration of siRNA of 16 nM in a total volume of 50 μl. Cells were incubated with the transfection solution for 48 h and before fixation staining of cells. During automated high-content screening, a total of 9 pictures per well were acquired for the analysis of over 1000 cells using a 20x air lens using an Olympus ScanR wide-field microscope equipped with DAPI, FITC, Cy3 and Cy5 filters. Automated image quantification and analysis were performed in the ScanR station software (Olympus ScanR Image Analysis Software version 3.0.0 & 3.0.1) based on mean intensities of fluorescent signal. The analysis pipeline used the DAPI channel for the detection of the nuclei and the application of a mask that served to quantify pixel intensities for EdU incorporation and γΗ2ΑΧ for each individual cell. ScanR data was processed in Excel for heatmap generation, normalization of systematic plate effects and calculation of percentage of negative controls. For analysis of significance and hit identification based on γH2AX and EdU, positive cells or mean intensity values were normalized for systematic plate (column-wise “CNORM”) effects prior to calculate the percentage of control values and standardize these values by robust Z-score transformation. For analysis of cell counts, raw values were used to calculate the percentage of control and robust Z-score transformation. Only siRNAs with a mean z-score ± 2 standard deviations to negative controls were considered significant. To ensure on-target specificity, a candidate was selected as a true hit only if a z-score greater than 2 was achieved by a minimum of two out of three individual siRNA oligos.

Plasmid transfection
Transient transfections of plasmid DNA were carried out using the Lipofectamine 2000 transfection reagent, according to the manufacturers’ instructions, and a final concentration of 1 μg/ml of the corresponding plasmid DNA. For each experiment, cells were plated at ~70–80% confluency in 35 mm dishes. Cells were incubated with the mixture of Opti-MEM, Lipofectamine 2000 and plasmid DNA for 5 h at 37°C, and then the transfection medium was removed and full DMEM 10% FBS was added to cells. Control over transfection efficiency was monitored 24–48 h post-transfection through fluorescence-microscopy (IF) and by Western Blot (WB) analysis.

DNA fiber assay and S1 nuclease DNA fiber assay
For experiments with siRNA treatment and drug treatment, exponentially growing cells were pulse-labeled with 25 μM IdU (5-Iodo-2’-deoxyuridine, Millipore Sigma) for 30 min, washed 3 times with pre-warmed PBS, then pulse-labeled with 250 μM CldU (5-Chloro-2’-deoxyuridine, Millipore Sigma) for 30 minutes, followed by 3 washes with cold PBS. Cells were harvested, pelleted at ~300×g for 5 min at 4°C, and resuspended in PBS for a final concentration of 7,5*105 cells/ml. For the DNA fiber assay with the ssDNA-specific S1 nuclease, cells were permeabilized with CSK100 (100 mM NaCl, 10 mM MOPS pH 7, 3 mM MgCl2, 300 mM sucrose and 0.5% Triton X-100 in water) after the CldU pulse for 10 min at R.T., washed once with PBS and then treated with the S1 nuclease (18001-016, Thermo Fisher Scientific) at 20 U/mL in Nuclease S1 buffer for 30 min at 37°C, and collected in PBS-0.1% BSA with cell scraper. Nuclei were then pelleted at 4600 xg for 5 min at 4°C, then resuspended in PBS (nuclei cannot be quantified, so initial number of cells plated should be considered when resuspending to a final concentration of 1500 nuclei/μl).

For both the standard DNA fiber assay and the S1 fiber assay, 6 μl of cells were mixed with 7 μl of lysis buffer (200 mM Tris-HCl pH 7.5, 50 mM EDTA, 0.5% SDS in water) on top of a positively charged glass slide, pre-coated with a freshly prepared solution of methanol: glacial acetic acid at 3:1 for 10 min. After 3 min incubation at RT, slides were tilted at 15–20 angle to spread the fibers at a constant, low speed. After air drying for 10–15 min at RT, DNA was fixed onto the slides with a freshly prepared solution of methanol: glacial acetic acid at 3:1 for 5 min, dried, and stored at 4°C overnight. For immuno-staining of DNA fibers, DNA was denatured with 2.5 M HCl for 80 min at RT. Slides were then washed with PBS three times and blocked with 5% BSA at RT for 20 min. DNA fibers were immuno-stained against BrdU (1:400, Bu1/75 Ab6326, Abcam and 1:25, 347580, BD Biosciences) for 2 h at RT, washed 3 times with PBS for 5 min each, and then incubated with stringency buffer (10 mM Tris-HCl pH 7, 400 mM NaCl, 0,02% Tween, 0,02% NP-40) for 10 mins. After 3 washes with PBS of 5 min each, DNA fibers were incubated with anti-rat Alexa Fluor 488 (1:750, A21470) and anti-mouse Alexa Fluor 546 (1/1000, A21123) and then washed 3 times with PBS for 5 min each, before mounting with 100 μl Mowiol®. Images were acquired using the LAS AF software using TCS SP5 confocal microscope (Leica) with a 63x oil immersion objective and each experiment was repeated at least two times independently. At least 15 images were taken across the whole slide using only one channel to select the regions for the images, to avoid any potential bias. At least 100-150 individual tracts were scored for each dataset. For all DNA fiber experiments, we measured both IdU and CldU tracts only on forks characterized by contiguous IdU-CldU signals. The length of each tract was measured manually using the straight-line tool on ImageJ software. Pixel values were converted into μm using the scale bar generated by the microscope software. Size distribution of tract length or ratios from individual DNA fibers was plotted as scatter dot plots with the center line representing the median. Data were pooled from independent experiments. Statistical differences in DNA fiber tract length were determined by the Mann-Whitney U test.

Chromatin fractionation
Cells were scrapped from the surface of the plate with ice-cold PBS and 1:3 of the volume was kept as total protein isolates (WCE), boiled for 5 min at 95°C and stored at −80°C until further analysis. Samples were centrifuged for 5 min at 500g at 4°C and the supernatant was discarded and replaced by 100 μl of Buffer A (10 mM Hepes pH=7.9, 10 mM KCl, 1.5 mM MgCl2, 340 mM Sucrose, 10% Glycerol, 1 mM DTT, 0.1% Triton X-100, 1x Protease inhibitor cocktail – Complete EDTA-free). Samples were then centrifuged for 5 min at 500g at 4°C and the supernatant was transferred to new tubes and stored in FSB-DTT as the cytoplasmic fraction. Sample pellets were then resuspended in 75 μl of Buffer B and kept on ice for 10 min before being centrifuged for 5 min at 1700g at 4°C. The supernatant was transferred to new tubes and stored in FSB-DTT as the nuclear fraction. Insoluble chromatin was collected by washing cell pellets with 50 μl of Buffer B (5 mM Hepes pH=7.9, 3 mM EDTA, 0.2 mM EGTA, 1 mM DTT, 1x Protease inhibitor cocktail – Complete EDTA-free) and by sonication at 6 cycles of 5s ON/OFF and 30% amplitude before adding FSB-DTT.

Western blotting
Cells were cultured in 35mm dishes and whole cell extracts were obtained by lysis of pelleted cells in 1x FSB-dTT. Samples were boiled at 95°C for 5 min and centrifuged at 4°C for 5 min at 15.500g. Lysates were loaded in a 4%-20% SDS gel, according to each experiment with the use of a marker for the molecular weight (MWP04, Nippon Genetics), following standard procedures. After the protein separation, proteins were then transferred from the polyacrylamide gel to an Immobilon-P PVDF membrane (Millipore, IPVH00010) using a wet blotting system (Biorad). Membrane sections were blocked for 1 hour at RT using a blocking buffer containing 5% milk in PBS – 0.1% Tween, unless used for the immunodetection of p-Chk1 or Chk1, when the membrane was blocked with a blocking buffer containing 3% BSA in 1x PBS – 0.1% Tween. Primary antibodies were incubated for 16 hours at RT in blocking buffer. Membranes were washed thoroughly with PBS − 0.1% Tween (2 quick washes, 1x15min, 3x5min), before incubation with secondary HRP-conjugated antibodies for 1 h at RT. Membranes were washed again with 1x PBS − 0.1% Tween (2 quick washes, 1×15min, 3×5min) and bound antibodies were detected through a chemiluminescent reaction with the use of Clarity™ ECL, by using the ChemiDoc imaging system (Biorad). A list of the antibodies used in this study can be found in Supplementary Information. Uncropped scans are provided in the Source Data.

Clonogenic assays
For cell survival assays, after the specific treatment was performed, cells were seeded onto 6-well plates in technical triplicates for each condition (750–1000 cells per well). Cells were cultured for 10–14 days, with a change of media in-between, before fixation with the fixative solution (0,5% crystal violet, 20% methanol in water). Percentage of survival was measured by manual colony counting after methanol/0.5% crystal violet staining.

RNA isolation, reverse transcription, and quantitative PCR (qPCR)
Total RNA was extracted from cells using the NucleoSpin RNA kit (740955.50, Macherey-Nagel) according to manufacturers’ instructions. cDNA was synthesized by M-MLV Reverse Transcriptase (28025013, Invitrogen) and PCR was performed using 10 ng of cDNA as a template, using the KAPA SYBR® FAST qPCR reagent kit (Kapa Biosystems) in a StepOne™ Real-Time PCR System (Applied Biosystems) according to manufacturers’ instructions. The results were calculated according to the 2-ΔΔCt methodology and are shown as relative expressions to the correspondent control. Results were then analyzed using the REST-MCS beta software.

Metaphase spreads
Mitotic cells were collected by incubation with 100 ng/ml Nocodazole for 12 h at 37°C and mitotic shake-off. After centrifugation, cell pellets were incubated in a hypotonic solution (0.075 M KCl) for 20 min at 37°C and fixed in a freshly prepared mix of methanol: acetic acid (3:1). For chromosome spreading, nuclear preparations were dropped onto pre-fixed slides and stained with DAPI. Image acquisition was performed using a Nikon Eclipse TE2000-U or Olympus IX83 widefield microscopes at 63x magnification. An appropriate number of metaphase spreads were imaged so to have representative numbers for subsequent analyses on the number of DNA fragments and chromosomal aberrations. After image acquisition, chromosomes were analyzed using ImageJ/Fiji to account for DNA fragments, chromosome breaks or abnormal chromosome structures. Only metaphase spreads clearly isolated were analyzed.

Chromatin immunoprecipitation ChIP
For chromatin immunoprecipitation (ChIP-qPCR), 1–2×107 cells were transfected with the indicated siRNAs for 48 h and were then fixed by incubation in 1% formaldehyde for 10 min at RT. Fixed cells were then quenched with 200 mM glycine, washed in PBS and frozen at −80°C for later processing. Lysates were generated by serial incubation of cell pellets in lysis buffers and chromatin was sheared by sonication using a Bioruptor Picosonicator (Thermo Fisher Scientific) until the average sheared size was 500 bp. Sheared chromatin was then cleared by high-speed centrifugation and used for immunoprecipitation (IP). For each IP, 40 μl of Protein A Dynabeads (Thermo) were washed, blocked in PBS + 0.5% BSA and conjugated with 1 μg of anti-FANCD2 or anti-IgG and then incubated with 30 μg of cleared, sonicated lysate. Beads were washed three times in RIPA buffer and DNA was eluted, de-crosslinked and purified for qPCR analysis in order to validate the assay. Reactions were run using Universal SYBR Green Supermix (Bio-Rad), and all qPCRs were performed using a PCR 7500 Fast reader (primers used are available upon request). The dilution factor was adjusted, and the percentage of the input signal was calculated.

ChIP-seq and genome-wide data analysis
After ChIP was conducted using the standard protocol, sequencing libraries were built using the ThruPLEX DNA-Seq Kit (Takara Bio) and size-checked with a Bioanalyzer DNA High sensitive chip. Samples were then sequenced in a NextSeq500 Illumina platform using a 2x75bp paired-end read high output flow cell with high success (92.91 % >=Q30). Sequencing data were first subjected to quality control filtering, demultiplexing and adapter trimming using the BaseSpace SeqHub from Illumina. Downstream processing of ChIP-seq data was performed using the European Galaxy Platform v18.05-20.05 (https://usegalaxy.eu), UNIX command line (GNU bash, version 4.2.46(2)) and R studio (2022.07.0 Build 548) with R 4.2.1 version; Reads were mapped to the canonical version of human reference genome hg38 (H. sapiens, GRCh38 no-alt analysis set; (https://bowtie-bio.sourceforge.net/bowtie2/manual.shtml) with Bowtie2111 and further processed using SAMtools112 (up to Galaxy Version 2.0.3/ UNIX 1.9). PCR duplicates were removed and only properly paired primary alignments with a mapping quality (MAPQ)>20 used for downstream analysis. Coverage data were computed using RPKM normalization for each file with deepTools113 (up to Galaxy Version 3.3.2.0.0/ UNIX 3.5.1). Then, mean coverages from the two biological replicates were calculated and the Input subtracted from IP signal, for better visualization of the results. Peak calling was performed with MACS2 package114 (up to Galaxy Version 2.1.1.20160309.6) allowing broad peak identification (--broad option) and setting p-value and broad cutoffs <0.01. Biological replicates were then intersected using BEDtools115 (up to Galaxy Version 2.29.2 / UNIX 2.29.2) and only peaks detected in both replicates considered for further analysis. Comparative analysis was conducted using BEDtools to determine condition-specific peaks (only appearing in one condition). We used HOMER79 (UNIX v4.11) for standard genome annotation of the peak datasets with the default annotation dataset and a homemade annotation dataset to cross the peak datasets with genome features of our interest: common fragile sites (CFS), centromeres and telomeres (retrieved from UCSC), rDNA, R-loops79 and DNA replication origins66. We applied contingency tests to calculate log₂ odds ratios and evaluate the statistical significance of differences between conditions. We crossed peaks with chromatin features retrieved from UCSC116 (bedGraphToBigWig) (UNIX v4) by intersecting genomic regions using BEDtools. Geecee package117 was used to calculate GC content. Integrative Genome Viewer (IGV)118 (v2.15.2) for visualize genome-wide data and deepTools for metaplot analysis. Additional genome-wide tracks used in this work were retrieved from sources as determined in refs. 79,119, except K562 Repli-Seq data which was obtained from the ENCODE project (Replication Timing Series: ENCSR591OXO) and average coverage of Repli-seq data from each S subphases measured at specific sites using deepTools. Human ERFSs were inferred by lifting over ERFS data from synchronized mouse B cells data using LiftOver (UCSC)120 (Kent, W.J., Sugnet, C.W., Furey, T.S., Roskin, K.M., Pringle, T.H., Zahler, A.M., & Haussler, D. (2002). The Human Genome Browser at UCSC. Genome Research, 12(6), 996-1006. https://doi.org/10.1101/gr.229102), as previously done76.

TCGA data analysis
Data were accessed through the public TCGA data portal and the cBioPortal for Cancer Genomics (http://www.cbioportal.org). For each cancer type, samples with both expression and CNA data were divided in quartiles based on the mRNA expression of CDT1. FANCD2 expression was assessed in tumors with high CDT1 expression (samples in the first quartile of the CDT1 distribution) and high CDT1 expression (samples in the fourth quartile).

Image acquisition, statistics, and reproducibility
Images obtained from immunofluorescence experiments, DNA fibers and metaphase spreads were acquired with Nikon Eclipse TE2000-U, an Olympus IX83 inverted widefield microscope or a Leica TCS SP5 confocal scanning microscope. Acquisition of images for colocalization assays and EdU incorporation was performed using a Leica TCS SP5 confocal scanning microscope. Images were analyzed with ImageJ/FIJI software (64-bit, National Healthcare Institute, USA). Statistical analysis and graph were generated using GraphPad Prism version 7 or 9 and Metamorph (Molecular Devices). Statistical significances (p values) of the observed differences between conditions are indicated in figure panels and figure legends.

Reporting summary
Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability
ChIP-seq data have been deposited in NCBI’s Gene Expression Omnibus with GEO Series accession number GSE285033. A full analysis of the Microscopy data is available at Zenodo (https://doi.org/10.5281/zenodo.18000043). Raw image files and reagents are available from the corresponding author. Source data are provided with this paper.

Code availability
The custom ImageJ/Fiji and CellProfiler macros used for the analysis of nuclear areas, signal intensity, foci quantification, and colocalization have been deposited in a public GitHub repository [Link: https://github.com/ElenaKarydi/Image-analysis-pipelines/releases/tag/v1.0.0].

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "Major Comments:\n1.The main approach in the study to measure re-replication is nuclear area measurements. Re-replication is associated with\nincreased origin firing. While the authors do perform DNA fiber studies, there is no mention of % initiation or inter origin\ndistance which is very peculiar. This needs to be addressed.\n2.It is unclear why authors haven’t assessed copy number changes that are very indicative of re-replication by sequencing or\nDNA microarrays. In addition, re-replication can cause aneuploidy which is easy to detect and report.\n3.Does Geminin loss or co-depletion with FANCD2 induce cell death? This needs to be addressed.\n4.The entire manuscript hinges on one molecular approach, siRNA mediated transient depletion to study the role of Geminin\nin re-replication. An alternate approach to inhibit such as the small molecule inhibitor targeting CDT1/Geminin protein\ncomplex (PMID: 35548337) and CRISPR to stably knockdown Geminin and(or) CDT1 is required to strengthen the findings.\n5.Authors mention “significant increase in the nuclear area and the proportion of cells with >4N DNA content as well as\nincreased γH2AX and 53BP1 signals” – In Fig S1a, below the immunoblot, there are 2 cell cycle histograms without labels.\nAssuming that the first one is control cells and the second one is GMNN-depleted cells, it doesn’t make sense that there are\nalmost no cells in S-phase considering that the cells have accumulated DNA damage. The profile only shows a clear G2\narrest after transiting s-phase successfully. This needs to be addressed. A EdU PI plot would be better suitable to assess\nDNA content.\n6.Fig. S1b – similar methods of H2AX and 53BP1 measurement should be used. γH2AX is a very clean and standard IF\nantibody where data quantification is almost always presented as number of foci per nucleus. It is unclear why the authors\nchoose to present this analysis as signal intensity when the gold standard is foci/nucleus.\n7.In Fig1a, if H2AX is labelled in red, what does the green signal indicate?\n8.The authors say “Importantly, among the top candidates we also found FANCD2…” There is no clear explanation for why\nthe authors chose to proceed with FANCD2 when there were a number of other more highly significant hits. This needs to be\nexplained and clarified in the introduction. Why not PLK1 or WEE1 or BRCA1/2?\n9.The authors move in and out of S-phase phenotypes and G2/M phenotypes in every figure without clearly distinguishing\nbetween them. This adds a lot of confusion and ambiguity to the data presented.\n10.In page 5 the authors mention that they used cell counts to measure cell survival and in the methods they report using\nclonogenic survival assays over 14 days. It is unclear how cell survival was measured and plotted, especially considering\nthey have primarily used siRNA mediated transient depletion for Geminin for most of the experiments. Why not use an assay\nlike Cell titer Glo for reporting survival? Example: In Fig S2c and Fig S2f it is unclear at what time point the cell survival was\nmeasured and how it was measured, and the times need to be marked on the figure.\n11.Authors state “Subcellular fractionation of U2OS cells depleted of Geminin revealed increased FANCD2 recruitment to\nchromatin (Fig. 2a)”. This is shown by a fractionation blot. The quantification lacks error bars from 3 independent\nexperiments.\n12.Authors state “Consistently, Geminin depletion also induced the formation of FANCD2 foci, which localized near γH2AX\nsites (Fig. 2b).”. This statement cannot be made without a co-localization analysis after confocal microscopy or atleast after\nobtaining z-stacks of the different planes.\n13.From figure 2, the authors conclude that “FANCD2 is recruited to chromatin and it is upregulated upon aberrant DNA\nlicensing.”. The chromatin recruitment of FANCD2 is always increased upon DNA damage so it is not clear whether this has\nanything to do with origin licensing.\n14.Assessment of proteins recruited to nascent DNA by iPOND or an analogous approach incorporating some timepoints\nwould be critical in establishing the timing of FANCD2’s recruitment to the chromatin. Is FANCD2 at the origins or is it at\ntraveling forks?\n15.Authors state “Double Geminin and FANCD2-depleted cells significantly increased fiber tract length and fork asymmetry”.\nThis experiment is very confusing. Since the co-depletion is carried out 72h prior to the fiber pulsing, one would expect both\nIdU and CldU lengths to be equal and if the co-depletion were to increase fork lengths, that would have to have an effect on\nboth IdU and CldU lengths equally. Why would the IdU and CldU tracts not be of equal lengths? Why do the authors expect\nasymmetry when there is no drug treatment involved? Additionally, why is the IdU tract unchanged but the CldU longer? The\nauthors also need to present IdU lengths atleast in the supplemental data.\n16.In figure 3e, the authors jump directly to assessing the G2/M checkpoint while investigating replicative abnormalities and\nDNA damage, without really addressing the intra-S checkpoint. The only mention of anything related to the intra-S\ncheckpoint is the assessment of pCHK1 in Fig 3f but the authors do not address the significantly reduced total CHk1 levels\nin the co-depleted cells. There is also no assessment of ATR protein expression/activity before their inhibition with VE-821.\nIn addition, FANCD2 depletion is not complete.\n17.Authors state “Importantly, G2-arrested cells in Geminin-depleted cells retained active DNA synthesis, as demonstrated\nby EdU incorporation and PCNA-staining, while exhibited high levels of DNA damage”. In Fig 3e and S3f, the authors don’t\nquantify the PCNA or DNA damage.\n18.Foci intensity measurements are tricky because they are completely dependent on slide prep, staining and imaging\nconsistencies. Whenever possible as in the case of H2AX, 53BP1, FANCD2 and in Figure 4 RPA32, it is essential to\npresent foci/cell nucleus rather than image intensity.\n19.Again, in Figure 4 there is extensive inconsistencies in measurement of RPA. Some experiments use intensity\nmeasurements while others use foci/nucleus measurements. Authors should stick to foci/nucleus for all measurements. In\nFig 4, authors could detect ssDNA foci by incubating the cells with a nucleotide analog (e.g., BrdU under non-denaturing\nconditions) to directly track ssDNA foci, which would provide more comprehensive evidence.\n20.In the metaphase analysis presented in Fig 4e, significance comparisons need to be presented for all cell lines groups.\nImportantly, in addition to aberrations, since re-replication results in aneuploidy, this needs to be reported.\n21.Figure 5 is missing an immunoblot that shows loss of FANCD2 monoubiquitylation. In addition, since this is a very\nimportant point, depleting Geminin in a K561R*-FANCD2 expressing the monoubiquitylation deficient FANCD2 should\nprovide critical insights.\n22.ChIP-qPCR experiment in S5b needs replicates to generate error bars. If this experiment was just to show that the\nFANCD2 antibody works for ChIP, then it can just be excluded from the manuscript. These fragile sites can very well be\nassessed from the ChIP-seq data. It is unclear why the qPCR data is being presented.\n23.The authors state “However, we noticed that FANCD2 was specifically enriched at replication origins in Geminin-\ndepleted cells when compared to control cells (Fig. 6c).”. Where are origin locations being annotated from? However, even if\nit is truly the sites of origin firing, the difference between control and Geminin-depleted cells is just not evident from the bar\ngraph presented. The difference seems very modest and definitely not significant. Following up using a second approach to\nactually map origins and correlate with FANCD2 ChIP-seq data is essential to make this conclusion.\n24.The authors state “Furthermore, analysis of the functional elements of FANCD2-enriched genes showed a significant\naccumulation of FANCD2 at intergenic regions and introns in control and Geminin-depleted cells (Fig. 6e), consistent with\nFANCD2 being recruited to repetitive sequences and the coding region of long genes contained within CFSs62,63”. The\nimportant thing to note is the lack of difference between control and Geminin depleted cells. The authors are over stating a\nconclusion that is yet to be drawn. They need to provide a list of genes that are either long and(or) are known for their\nrepetitive DNA content from their ChIP-seq analysis to make this statement. A volcano plot depicting log fold change in\ngenes versus log p-value or FDR could help with this.\n25.Based on the data presented, how can we conclude that FANCD2 enrichment at promoter-transcription start sites (TSSs)\nin Geminin depleted cells is significant.\n26.The authors state “Consistent with peak analysis (Fig. 6e), more than 90% of the origins enriched in FANCD2 specifically\nin Geminin-depleted cells colocalized with genes (Supplementary Fig. 5d), thus indicating these origins lie within genes.”.\nWhere are these origins being mapped from? There have been no experiments conducted to map origins in this study. The\ndata is extremely correlative at best. The authors need to map origins in their cell lines because this is a critical point in the\nmanuscript.\n27.The authors state “Thus, Geminin loss leads to FANCD2 enrichment at replication origins within highly transcribed genes\ngenome-wide.”. A list of these highly transcribed genes from their ChIP-seq data set would be a very important inclusion to\nthe manuscript.\n28.The authors state “Thus, we next analyzed the replication timing of origins identified to accumulate FANCD2 specifically\nin Geminin-depleted cells. Using publicly available genome-wide Repli-seq data from the ENCODE project 72, we observed\na significant increase in BrdU intensity during early S phase and a reduction in mid-to-late replicating origins compared to\ncontrol cells (Fig. 6f), suggesting these origins are predominantly early replicating.”. It is unclear whether the authors\nperformed the repli-seq to generate the data or analyzed existing Repli-seq data from the ENCODE project. It is very\nstraightforward to check if the existing list of ERFs (PMID: 23352430 and PMID: 36099913-for BRCA1 and BRCA2) and the\nlongest genes in the human body are both enriched for FANCD2 in the Geminin depleted cells from their ChIP-seq data.\nWhy has this not been prioritized?\n29.For the Cordycepin experiments: The decreases observed in co-depleted cells although significant are very modest.\nTranscription inhibition by Cordycepin often shuts down DNA replication as well which would be highly counterproductive\nfor the study focused on re-replication. EdU incorporation in Cordycepin treated cells needs to be presented to address this.\n30.The Model presented in Fig. 7 is too simple and doesn’t convey the message of the paper with any clarity.\n31.The title ”FANCD2 limits unrestrained fork progression and prevents fragility of early replicating sites upon aberrant origin\nlicensing” is not very accurate. The paper does not address unrestrained fork progression in the absence of FANCD2 or\nfragility of early replicating sites by cytogenetics using FISH probes and there is definitely no measure of aberrant origin\nlicensing anywhere in the paper.\nMinor:\n1.Page 4: “co-depletion of CDT1 partially restored these phenotypes”. – change restored to rescued\n2.In Fig S1a, the cell cycle plot labels need to be bigger to match the other fonts.\n3.Supplementary 2d-f say KO FANCD2 which is inaccurate. It needs to mention that they are PD20 patient cells completing\nlacking FANCD2.\n4.In every bar graph presented, the individual experimental means need to be shown as dots representing the 3\nindependent experiments.\n5.Significance representations and comparisons should be presented for all groups and not selected ones.\n6.Figure 3f Chk1 protein is misspelled on the gel.\n7.Need to add a sentence introducing caffeine as a checkpoint inhibitor\n8.Fig 4d needs to be sub divided into smaller figures\n9.Typos in figure legends need to be carefully rectified.", "reviewer_2": "In this work the authors analyse how cells deal with the deregulated licensing of origins of replication caused by the\ndepletion of Geminin. Unlike previous studies, where defective licensing was driving large amounts of re-replication in\ncancer cells, this study focuses on milder levels of DNA re-replication seen in immortalised cells, such as hTERT-RPE1\ncells.\nThrough an siRNA screen, the authors identify FANCD2 as key in suppressing DNA damage and supporting cell\nproliferation upon Geminin depletion. They then show that FANCD2 loss causes faster fork speed and accumulation of S1-\nnuclease sensitive stretches of DNA, likely ssDNA gaps. Genomic analysis reveals that in Geminin depleted cells, FANCD2\nis not only at difficult and late replicating DNA regions, but it is also seen at early replicating regions, in the proximity of\nhighly transcribed genes and clusters of early firing replication origins.\nSurprisingly they find that dampening of transcription or R-loops digestion significantly supresses DNA damage in Geminin\ndepleted cells.\nThis study suggests a new mechanism for the generation of genomic instability upon reactivation of replication origins within\nthe same cell cycle, which now directly implicates transcription and collision between the “re-replication” and the\ntranscription machineries. Such a mechanism is likely to occur in pre-malignant cells leading to tumorigenesis.\nThe experiments are well performed and controlled, combining both classical cell biology and genomics approaches; the\nresults are clear-cut and important. The manuscript is well written and easy to read.\nThis reviewer has few minor comments that may improve the manuscript:\n1. In supplementary Fig S1, it would be nice to show the efficiency of the co-depletion of Geminin and CDT1by western blot\nand assess if CDT1 and CDT1+Geminin depleted cells are still efficiently replicating their DNA under these experimental\nconditions.\n2. In many instances nuclear area is used as surrogate for re-replication. I understand that such approach may not be\nparticularly sensitive, thus creating some confusion when the authors argue that geminin depletion has sometime “mild” or\n“no” effects in immortalised cells. How does this assay compare to classical DNA content measurement by flow cytometry or\nother techniques?\n3. Please provide legends for Supplementary tables 1 and 2\n4. The siRNA screen appears to also identify proteins that, when depleted, cause DNA damage which is prevented by\ngeminin depletion e.g. PLK3, NEK1, CDK7 and few others. If this is correct, can the authors mention and discuss it?\n5. Fig 3C: does siFANCD2 increases fork speed compared to siCtrl?\n6. Line 211 and Fig S3c-d: it is not clear why low doses of ATR inhibitors suppress re-replication. It is known that ATR\ninhibition facilitates origin activation, thus I would expect the opposite phenotype. Is this a dose-dependent effect and is it\nrelated to the override of mitotic checkpoint thus not leaving sufficient time for large quantities of DNA to accumulate? (see\nalso point 2 related to sensitivity of the nuclear area assay).\n7. Line 337: how effective is Cordycepin in inhibiting general transcription and. in particular. in inhibiting transcription of the\ngenes located in the proximity of clusters of early firing origins?\n8. Similarly, would Cordycepin treatment or RNAse1 overexpression also suppress FANCD2 binding at siGEMNN specific\npeaks determined in fig 6b?", "reviewer_3": "Initiation of eukaryotic DNA replication is tightly controlled and coordinated with cell cycle progression to ensure only a\nsingle round of DNA synthesis. In metazoa this occurs primarily through careful regulation of CDT1 and Geminin levels.\nDefective regulation of CDT1/Geminin can lead to re-replication of the genome which is mutagenic, toxic, and is implicated\nin oncogenesis. However, the exact molecular consequences of CDT1/Geminin dysregulation are unclear. Badra-Fajardo et\nal have investigated the consequences of Geminin dysregulation and identified an important role for FANCD2 in preserving\ngenome stability. They find that loss of FANCD2 causes DNA damage, chromosomal aberrations, and possibly cell death in\nGeminin-deficient cells. Mechanistically, these consequences are proposed to arise from accumulation of single-stranded\nDNA gaps and formation of DNA breaks that are a downstream consequence of re-replication. Furthermore, these effects\nmay be unequally distributed throughout the genome. Overall the authors have connected FANCD2 to defects in Geminin\nfunction but there are major concerns about the mechanism proposed, the interpretation of the data, and the overall novelty\nof FANCD2 function supported by the data.\nMajor concerns\n1. The manuscript seems to imply a unique role for FANCD2 in responding to Geminin loss. However, the upstream\n(FANCA in Fig 5) and downstream (RAD51, BRCA1, BRCA2 in Fig 1) components of the Fanconi Anemia pathway behave\nsimilarly. It would thus seem that the role described for FANCD2 here reflects the Fanconi Anemia pathway’s function and\nnot something more unique about FANCD2. Because a major role of the FA pathway is to coordinate DNA breaks in a\ncontrolled manner (by XPF-ERCC1) this would seem to contradict the model that FANCD2 is preventing breaks. The data\nwould certainly support a model that FANCD2 prevents break accumulation, but not that it prevents break formation.\n2. It is well established that cancerous cells are much more readily undergo re-replication following Geminin/CDT1\ndysregulation. hTERT-RPE1 cells are examined in Figure 1 but the remainder of the manuscript focuses on cancerous cells\n(U2OS, MCF7, 293T, HCT116). It’s unclear how much of the observed effects are specific to cancerous cells.\n3. The manuscript focuses on re-replication but this is only directly measured once (Fig. S1A). For the remainder of the\nmanuscript, nuclear size is used as a proxy. Not only is nuclear size very indirect but it is unclear whether it serves a\nreasonable proxy: in Fig S1A the fraction of cells that exhibit increased nuclear area is much greater than the fraction of cells\nthat exhibit >4n DNA content. The lack of direct measurements of re-replication and concerns about the suitability of cell size\nas a proxy for re-replication call into question most of the authors’ conclusions that relate to re-replication.\n4. Geminin knockdown is likely to have effects other than re-replication of the genome. The mechanism of re-replication by\ngeminin is loading of additional MCM complexes followed by origin firing on the already-replicated DNA. There is no reason\nthis could not also occur in the unreplicated region of the genome, which could manifest as unscheduled/excess origin firing\nand may account for some of the effects observed (e.g. PMID29466339). Taken together with point (1), above, it seems very\npossible the authors are documenting breaks induced by replication-transcription collisions and repaired by the Fanconi\nAnemia pathway.\n5. The is an ~24 hour delay between FANCD2 and gamma H2AX focus formation (Fig. 3A). Because the cell cycle in U2OS\ncells (used in Fig 3A) is approximately 24 hours it would seem very unlikely that the FANCD2 levels and damage are both\noccurring within S phase or G2, as would seem to be implied by the model that FANCD2 protects forks from excessive\nbreakage. The H2AX foci would at the very least need to arise from the following S phase, but the simpler explanation would\nbe that they arise from breaks during mitosis.\n6. In several figures the authors report increased number of foci of different DNA damage response proteins following\nGeminin knockdown (Figs 1c, 2d, 3a, 4a, 4d, 5a, 5d, 7d-e). Because Geminin increases nuclear area there would be\nexpected to be some increase in foci per nucleus independent of any effects on DNA damage. The same issue may also be\ntrue for nuclear signal intensity (depending on how it was calculated). As the data stands, many of the increases in foci (and\npossibility nuclear signal) could be a consequence of increased nuclear size.\n7. Similar to the concerns about nuclear size, re-replication is expected to result in more DNA per nucleus and thus an\nincrease in replication proteins on DNA (E.g. RPA in Fig 4A-B). The same would also be true for DNA damage foci because\nmore replication would increase the level of spontaneous DNA damage. It is thus not clear how meaningful increased DNA\nreplication/repair foci are under conditions where re-replication may be taking place.\n8. The genome wide distribution of FANCD2 presented in Figure 6 may not be particularly meaningful because there is no\ncorrection for the prevalence of different genomic features. For example, the prevalence of FANCD2 peaks at intergenic\nregions and introns in Fig 6E could simply be because this constitutes most of the genome. Similarly, the approximately 50%\noverlap between FANCD2-bound genes in Fig. 6D is very similar to what would be expected for any two randomly selected\nsets of genes. As things stand, many of the conclusions drawn from genomic analysis may not be meaningful.\n9. The effects of Geminin and FANCD2 knockdown on cell survival seem to be purely additive (Fig. 1D), in contrast to the\neffects on the DNA damage response which are clearly synergistic (Fig. 1C). The DNA damage data fit with the authors’\nmodel of FANCD2 responding to Geminin knockdown but the viability data do not. It is not clear whether Fig 1D supports a\nrole for the authors’ proposed mechanism in ensuring cell viability.\nMinor concerns\n10. The legend for Figure 1 refers to panel c but appears to actually be referring to b.\n11. For Figure 2A it is not obvious that Geminin knockdown causes increased FANCD2 recruitment to chromatin. The\ndifference is modest and total cellular FANCD2 is increased so the increased binding could just reflect higher protein levels.\n12. It is unclear what the asterisk in Fig 2C corresponds to.\n13. The analysis of Cbio data in Fig. 2E cannot be used to draw a causal conclusion because this data only demonstrates\ncorrelation. Also it is unclear how this analysis constitutes n=3 as the legend would seem to indicate.\n14. When discussing Figure 5 the authors conclude that Geminin loss “promotes activation of canonical FA signaling and\nrecruitment of monoubiquitinated FANCD2”. They did demonstrate a role for FANCA following Geminin knockdown and can\ninfer importance of the FA pathway following Geminin knockdown, but they cannot infer any altered signaling (especially\nbecause they did not measure signaling).\n15. For Fig 7d-e how do the authors know that the RPA signal is due to gaps and not single-stranded DNA arising directly\nfrom R-loops?\n16. In the results section, the authors state “Finally, we tested whether increased formation of ssDNA gaps in Geminin and\nFANCD2 double-depleted cells would result in chromosome fragility and high levels of genomic instability.” There is no\ndirect measurement of gaps and the intervention (Geminin depletion) is highly indirect so they cannot make this conclusion.\n17. For Fig S1A the graphs of cell counts should be labeled to indicate which is control vs which is Geminin depleted"}, "author_response": "FANCD2 restrains fork progression and prevents fragility at early replicating origins upon re-replication\nWe sincerely thank the reviewers for the detailed evaluation of our manuscript. The\ninsightful and encouraging comments helped us to perform additional experiments and\na thorough revision of the manuscript. In this response letter, we address the reviewers’\ncomments point-by-point and outline the revisions we have made to enhance the quality\nand clarity of our study.\nPoint-by-point response to the reviewers’ comments:\nReviewer #1 (Remarks to the Author):\nMajor Comments:\n1.The main approach in the study to measure re-replication is nuclear area\nmeasurements. Re-replication is associated with increased origin firing. While the\nauthors do perform DNA fiber studies, there is no mention of % initiation or inter origin\ndistance which is very peculiar. This needs to be addressed.\nWe thank the reviewer for this consideration and agree that analyzing the percentage of\norigin firing was an important question to address in the context of aberrant DNA\nlicensing. In the revised manuscript, we included the percentage of origin firing in respect\nto the total replicative structures of DNA fibers obtained for each condition. The analysis\nof three replicates shows that Geminin depletion slightly increases the rate of origin\ninitiation, although this increase is not further enhanced by FANCD2 loss (see new Fig.\n3e). Thus, we believe that although Geminin loss might promote some activation of\ndormant origins as a consequence of aberrant CDT1 activity and/or to counteract the\nperturbations on replication -especially as cells undergo prolonged re-replication-\nFANCD2 primarily functions in reducing the progression rate of re-replicating forks rather\nthan directly controlling origin firing. These observations are in line with the idea that re-\nreplication occurs primarily at constitutive origins that are used during normal replication,\nwhich also tend to be early replicating1 where we indeed observe an accumulation of\nFANCD2 (see Fig. 6). These results are included and further discussed in the submitted\nrevised manuscript (page 8).\n2.It is unclear why authors haven’t assessed copy number changes that are very\nindicative of re-replication by sequencing or DNA microarrays. In addition, re-replication\ncan cause aneuploidy which is easy to detect and report.\nWe acknowledge the reviewer’s concern and agree that another approach to measure\nre-replication was important to strengthen our results. Although examining copy number\nchanges is an interesting metric, these changes have only been observed upon massive\nlevels of re-replication2,3. Geminin depletion typically induces partial re-replication and\ncell to cell heterogeneity, making it difficult to observe such differences by accounting for\nnet copy number changes. As for aneuploidy, this metric requires that cells pass through\nmitosis, however origin re-licensing after Geminin depletion activates checkpoint\nsignaling and promotes cell arrest in G2/M, which indeed we show is required for\nextensive re-replication. This, together with the fact that U2OS cells already present\nextensive levels of aneuploidy4, made it difficult to use this assay to quantify levels of re-\nreplication.\nAs an alternative and following the reviewers suggestions (including reviewer #1 in major\npoint number 5), we implemented flow cytometry analyses in U2OS and hTERT-RPE1\ncells depleted of Geminin at different timepoints to examine the kinetics of the cell cycle\nand the prevalence of cells with DNA content above G2/M cells incorporating EdU\nas a metric of re-replication. Our analyses in U2OS cells show that Geminin depletion\ninduces an initial accumulation of cells in S phase at 24 hs, while there is already a small\nproportion of cells over-replicating (>4 C) when compared to control cells (New\nSupplementary Fig. 1a). However, the high prevalence of cells with DNA content above\nG2/M levels occurs at 48 hs, where about 30% of cells are re-replicating, consistent with\nthe results observed by measuring nuclear areas (New Supplementary Fig. 1b) and\nreinforcing this timepoint as the appropriate for the remainder of the study. As expected,\nthere were no evident signs of re-replication in hTERT-RPE1 cells depleted of Geminin\n(New Supplementary Fig. 1e, f). Importantly, in the revised manuscript we also included\na new set of experiments using this approach to show the effect of depleting FANCD2 in\nre-replicating cells. According to the results obtained by measuring the nuclear areas,\ndepletion of FANCD2 did not increase the levels of re-replication but rather slightly\nreduced them (New Fig. 3c). We provide cell cycle distributions for each timepoint and\ncell line from 3 independent experiments together with gates applied in the Source data.\n3.Does Geminin loss or co-depletion with FANCD2 induce cell death? This needs to be\naddressed.\nWe thank the reviewer for this comment, and we are glad to further clarify the results\nfrom our work. Throughout the study, we performed clonogenic assays to focus on the\nlong-term proliferative potential of re-replicating cells, either in cells solely depleted of\nGeminin, or upon double silencing of Geminin and FANCD2 or the alternative targets\nindicated in each case (see New Fig. 1d, New Supplementary Fig. 2b, d and e, New\nSupplementary Fig. 3, New Fig. 4, New Fig. 5c, f and New Supplementary Fig. 5f). Rather\nthan survival assays -usually employed to reveal acute cytotoxicity at a short time\nintervals- we conducted clonogenic assays, as this approach can capture the ability of\nindividual cells to proliferate and form colonies over longer periods5-8. As we describe in\nthe results section, our analyses show that depletion of Geminin slightly reduces cell\nsurvival in normal cells (New Fig. 1d, while this effect is significantly more acute in cancer\ncells (New Supplementary Fig. 2b, d and e). Co-depletion FANCD2 renders Geminin-\ndepleted cells hypersensitive across cell lines (Fig. 1 and Supplementary Fig. 2). Cell\ndeath is not detected at early time points, when our analysis of molecular pathways takes\nplace, by either cytometry analyses (sub G1 population, New Supplementary Fig. 1 and\nNew Fig. 3), cell counts or MTT (data not shown). This rules out apoptotic pathways as\nthe cause of the observed molecular phenotypes.\n4.The entire manuscript hinges on one molecular approach, siRNA mediated transient\ndepletion to study the role of Geminin in re-replication. An alternate approach to inhibit\nsuch as the small molecule inhibitor targeting CDT1/Geminin protein complex (PMID:\n35548337) and CRISPR to stably knockdown Geminin and(or) CDT1 is required to\nstrengthen the findings.\nIn our initial submission, we compared the effect on cell survival of siRNA-mediated\ndepletion of Geminin across different cells lines, but we additionally used an auxin-\ninducible degron system integrated in HCT116 cells for the rapid degradation of Geminin\nand observed increased sensitivity to silencing of FANCD2 (New Supplementary Fig.\n2e). We performed additional analyses in these cells by quantifying the proportion of cells\nwith FANCD2 foci or γH2AX nuclear intensities. Geminin depletion alone elicits very high\nlevels of DNA damage in this system (Fig. R1a). This makes the detection of even higher\nγH2AX levels by co-depletion of FANCD2 technically challenging (Fig. R1b). We provide\nthese data, which are consistent with our findings, below. To further strengthen our\nresults, during the revision period we generated a stable U2OS cell line containing a\ndoxycycline (dox)-dependent inducible short hairpin (shRNA) targeting human Geminin.\nIn the revised version of this manuscript, we included a new set of data using this system\nand validating the increased sensitivity of Geminin-depleted cells to loss of FANCD2 in\nterms of DNA damage and cell survival (New Supplementary Fig. 2c, d).\n400\n300\n200\n100\n0\ntshceoH\nXA2Hγ\n)2mµ(\naera\raelcuN\nDMSO Auxin 24hs Auxin 48hs\nDMSO Auxin 24hs\ntshceoH\n2DCNAF\n40\n30\n20\n10\n0\n)%(\nicof\n2DCNAF\n01>\nslleC\n****\n****\n1.5x106\nns\n1x106\n5x105\nAux - + - +\nsiCtrl siFANCD2\n)UA(\nseitisnetni\nnaem\nXA2Hγ\na\nb\nFig. R1. (a, b) Representative images and quantification of asynchronously growing\nHCT116 cells treated with 1μg Doxycycline for 24hs prior to addition of Auxin at 300μM\nand stained against FANCD2, and γH2AX. For quantification of nuclear area, plot shows\nthe distribution of individual values of a representative replicate. For quantification of\ncells with >10 FANCD2 foci, plot shows the mean ± SD of biological replicates. For\nquantification of γH2AX nuclear intensities, plot shows the distribution of individual\nvalues of a representative replicate. These experiments were performed in two biological\nreplicates (n=2) and a minimum of 250 cells were counted per replicate for each\ncondition. DNA staining, Hoechst. Scale bar, 10μm. ns, non-significant, **** p < 0.0001\n(two-tailed t-test).\n5.Authors mention “significant increase in the nuclear area and the proportion of cells\nwith >4N DNA content as well as increased γH2AX and 53BP1 signals” – In Fig S1a,\nbelow the immunoblot, there are 2 cell cycle histograms without labels. Assuming that\nthe first one is control cells and the second one is GMNN-depleted cells, it doesn’t\nmake sense that there are almost no cells in S-phase considering that the cells have\naccumulated DNA damage. The profile only shows a clear G2 arrest after transiting s-\nphase successfully. This needs to be addressed. A EdU PI plot would be better\nsuitable to assess DNA content.\nWe thank the reviewer for pointing out this observation and suggesting this important\nexperiment. As previously mentioned, we performed the recommended experiment,\nwhich it thoroughly discussed in the response to comment number 2 of #Reviewer1. As\nthe reviewer reasonably points out, there is an initial accumulation of cells in S-phase\n24hs after Geminin depletion (New Supplementary Fig. 1a), which agrees with a potential\ndelay of S in response to re-firing of origins (New Supplementary Fig. 1). However, we\ncan observe that at 48hs the percentage of S-phase cells decreases as cells start\nmassively re-replicating (New Supplementary Fig. 1a).\n6.Fig. S1b – similar methods of H2AX and 53BP1 measurement should be used. γH2AX\nis a very clean and standard IF antibody where data quantification is almost always\npresented as number of foci per nucleus. It is unclear why the authors choose to present\nthis analysis as signal intensity when the gold standard is foci/nucleus.\nWe thank the reviewer for drawing our attention to this issue. We could not apply the\nsame methods to detect γH2AX and 53BP1 due to the differential staining of these\nmarkers; Quantification of γH2AX and 53BP1 foci relies on the visualization of discrete\nsites of localized DNA double-strand breaks. In our analysis however, depletion of\nGeminin triggered a pan-nuclear γH2AX staining rather than distinct foci in several cells\n(Fig. R2). This pan-nuclear γH2AX has previously been observed in cases of high DNA\ndamage. This diffuse staining lacked the spatial resolution required to identify foci and\ntherefore we could only compare nuclear intensities in our experimental system.\nFig. R2. a. Representative images of HCT116 mAIDGMNN cells (left), treated with Auxin\nand the indicated siRNA oligos. b. U2OS cells (right) treated with the indicated siRNA\noligos, and immunostained against γH2AX. Cells encircled in white depict a pan-nuclear\nγH2AX staining. DNA staining, Hoechst. Scale bar, 10μm. \n7.In Fig1a, if H2AX is labelled in red, what does the green signal indicate?\nWe thank the reviewer for pointing out this detail. The green signal accounts for EdU\nincorporation, which we used during primary screens as a marker of the proliferative\ncapacity of the cells. In the revised version of the manuscript, we also included the robust\nz-scores of EdU in respect to negative controls in the Supplementary Table 2.\n8.The authors say “Importantly, among the top candidates we also found FANCD2…”\nThere is no clear explanation for why the authors chose to proceed with FANCD2 when\nthere were a number of other more highly significant hits. This needs to be explained\nand clarified in the introduction. Why not PLK1 or WEE1 or BRCA1/2?\nAs the reviewer indicates, there were other candidate genes that were also highly\nsignificant in our primary screen in addition to FANCD2 (provided in New Table1, New\nTable2 and Source data). Some of these targets, however, were included in the\nscreening already as positive controls for cell death (e.g. PLK1) and/or DNA damage\n(e.g. WEE1, CHEK1 or CDT2). In some other cases, the levels of DNA damage were\nhigh in both control and Geminin-depleted hTERT-RPE1 cells (e.g. BRCA1/2). Since the\nfocus of our study was the identification of factors required to promote the survival\ninduced specifically by Geminin loss, these factors were not selected for further analysis.\nWe have now clarified further this point in the result section of this manuscript (page 5).\n9.The authors move in and out of S-phase phenotypes and G2/M phenotypes in every\nfigure without clearly distinguishing between them. This adds a lot of confusion and\nambiguity to the data presented.\ntshceoH\nXA2Hγ\nHCT116 mAIDGMNNN\nsiCtrl siFANCD2 GMNN\nAUX - + - + siRNA Ctrl GMNN FANCD2 FANCD2\ntshceoH\nXA2Hγ\na b\nU2OS\nWe thank the reviewer for highlighting this section of the results that could be improved.\nWe understand that the potential confusion might have arisen mainly due to the fact that\nre-replicating cells undergo DNA synthesis outside of S, that is, upon cell cycle arrest at\nG2. Initially, we tried to demonstrate this through immunofluorescence, showing that the\n“giant” nuclei with >4C DNA content also incorporate EdU. Following the reviewer’s\nrecommendation, in the revised manuscript we introduced early in the text the new data\nfrom the flow cytometry analysis using EdU and PI, which reveals a distinct cell\npopulation exhibiting >4C DNA content and high levels of EdU incorporation\n(Supplementary Fig. 1a), which we identify and refer to as re-replicating cells for the\nremainder of the manuscript. Additionally, we tried to re-structure the results included in\nthe study to thoroughly address the potential ambiguity regarding the characteristics of\nre-replicating cells.\n10.In page 5 the authors mention that they used cell counts to measure cell survival and\nin the methods they report using clonogenic survival assays over 14 days. It is unclear\nhow cell survival was measured and plotted, especially considering they have primarily\nused siRNA mediated transient depletion for Geminin for most of the experiments. Why\nnot use an assay like Cell titer Glo for reporting survival? Example: In Fig S2c and Fig\nS2f it is unclear at what time point the cell survival was measured and how it was\nmeasured, and the times need to be marked on the figure.\nThe primary screen was imaging-based; hence, we could not use CellTiter-Glo to\nmeasure cell survival without interfering with the fluorescent markers. As an alternative\nand since we plated the same number of cells per well, for the screening we used cell\ncounts as a preliminary measure of viability. We validated the robustness of this metric\nin small-scale pilot screens by transfecting the cells with cell death-siRNA controls or\nwith CDT2, which causes massive re-replication (New Supplementary Table 1 and New\nSupplementary Table 2). Consistently, the screening data also showed reduced cell\ncounts when using these controls or after silencing essential genes such as PLK1 or\ncyclin D (New Supplementary Table 1 and New Supplementary Table 2). As mentioned\npreviously, for subsequent analyses we employed clonogenic assays to assess the long-\nterm viability of the cells. We have included the workflow of the clonogenic assay\nindicating the time point of fixation and analysis as the reviewer suggested (New Fig. 1d)\nand have clarified these points in the text (page 6).\n11.Authors state “Subcellular fractionation of U2OS cells depleted of Geminin revealed\nincreased FANCD2 recruitment to chromatin (Fig. 2a)”. This is shown by a fractionation\nblot. The quantification lacks error bars from 3 independent experiments.\nWe thank the reviewer for this suggestion. In the revised manuscript, we included the\nstatistical analysis with respective error bars.\n12.Authors state “Consistently, Geminin depletion also induced the formation of FANCD2\nfoci, which localized near γH2AX sites (Fig. 2b).”. This statement cannot be made without\na co-localization analysis after confocal microscopy or at least after obtaining z-stacks of\nthe different planes.\nThe representative image shown in this figure was a maximum-intensity projection of\nmultiple z-stacks acquired by confocal microscopy. Following the reviewer’s\nrecommendation, we used Cell Profiler to measure the percentage of colocalization\nbased on the individual planes and the results are included in the revised version of the\nmanuscript (New Fig. 2b).\n13.From figure 2, the authors conclude that “FANCD2 is recruited to chromatin and it is\nupregulated upon aberrant DNA licensing.” The chromatin recruitment of FANCD2 is\nalways increased upon DNA damage so it is not clear whether this has anything to do\nwith origin licensing.\nWe thank the reviewer for this comment. Although it is true that FANCD2 binds to\nchromatin upon DNA damage, in our study we demonstrate that recruitment of this\nprotein to chromatin occurs during the first 24hs and prior to high levels of DNA breakage\n(see New Fig. 2c), which rules out the possibility that its recruitment is simply due to the\noccurrence of DNA breaks. Nevertheless, as a mean to strengthen our results showing\nthat FANCD2 binds to chromatin because of re-replication, in the revised manuscript we\nincluded a new set of data employing the SIRF assay (Single-cell Assay for in situ Protein\nInteraction with Nascent DNA Replication Forks). This technique combines EdU labeling\nof nascent DNA with a proximity ligation assay (PLA) to detect proteins within ~40 nm of\nnewly synthesized DNA9 and thus allowed us to directly visualize FANCD2 at nascent\nDNA. Using this approach, we observed a significant increase in the number of FANCD2-\nEdU interacting foci compared to control cells following Geminin depletion, (New Fig.\n3a). These results were also validated using proximity ligation assays (PLA) for\nendogenous PCNA-FANCD2 and MCM2-FANCD2 (New Fig. 3b and New\nSupplementary Fig. 4c). Importantly, this approach allowed us to observe that FANCD2\ninteraction with the replisome not only occurs in S-phase but becomes also evident in\nre-replicating cells exhibiting the characteristic EdU incorporation pattern of G2. Thus,\nour data supports a model in which FANCD2 is recruited at the vicinity of active,\nelongating replication forks. This point has been further discussed in the results and\ndiscussion sections of the manuscript. Of note, we have replaced the term “aberrant\norigin licensing” to “re-replication” along the entire study, which is more appropriate for\nthe results we observe, as the reviewer indicates.\n14.Assessment of proteins recruited to nascent DNA by iPOND or an analogous\napproach incorporating some timepoints would be critical in establishing the timing of\nFANCD2’s recruitment to the chromatin. Is FANCD2 at the origins or is it at traveling\nforks?\nWe thank the reviewer for this suggestion and agree that it was important to further clarify\nthe precise location of FANCD2 recruitment to chromatin. As described in comment\nnumber 13, in the revised manuscript we addressed this point by performing SIRF and\nstandard PLA assays, in which we demonstrate FANCD2 recruitment at the vicinity of\nactive, elongating replication forks (New Fig. 3a, b and New Supplementary Fig. 4c). This\npoint is now discussed in the results and discussion sections of the manuscript.\n15.Authors state “Double Geminin and FANCD2-depleted cells significantly increased\nfiber tract length and fork asymmetry”. This experiment is very confusing. Since the co-\ndepletion is carried out 72h prior to the fiber pulsing, one would expect both IdU and\nCldU lengths to be equal and if the co-depletion would to increase fork lengths, that would\nhave to have an effect on both IdU and CldU lengths equally. Why would the IdU and\nCldU tracts not be of equal lengths? Why do the authors expect asymmetry when there\nis no drug treatment involved? Additionally, why is the IdU tract unchanged but the CldU\nlonger? The authors also need to present IdU lengths atleast in the supplemental data.\nWe appreciate the reviewer’s comment, which allowed us to clarify this point. As the\nreviewer correctly noted, both IdU- and CldU-labeled fibers are indeed of the same\nlength. In a standard DNA fiber experiment, the first analog marks the replication forks\nthat are actively synthesizing DNA during the initial pulse, while the second analog\nreflects the progression of these forks during the subsequent pulse. This is the reason\nwe measured the length of the second analog (in this case CldU) to compare lengths\nbetween conditions, as it serves as the actual readout of replication fork progression. As\nfor asymmetry, we measured the ratio of the length of the second analog between two\nsister forks emanating from the same origin. Replication forks exhibit asymmetry under\nreplication stress (endogenous or exogenous)10,11. In our case, our data indicate that this\nasymmetry is linked to the replication stress arising from uncontrolled progression of\nreplicating forks in the absence of Geminin and FANCD2. We believe these points are\nnow clear in the revised manuscript. As requested by the reviewer, the raw data\ncontaining IdU and CldU fiber lengths and CldU ratios for fork asymmetry are included\nin the Source data.\n16.In figure 3e, the authors jump directly to assessing the G2/M checkpoint while\ninvestigating replicative abnormalities and DNA damage, without really addressing the\nintra-S checkpoint. The only mention of anything related to the intra-S checkpoint is the\nassessment of pCHK1 in Fig 3f but the authors do not address the significantly reduced\ntotal CHk1 levels in the co-depleted cells. There is also no assessment of ATR protein\nexpression/activity before their inhibition with VE-821. In addition, FANCD2 depletion is\nnot complete.\nWe thank the reviewer for bringing up this interesting point which we are glad to further\ndiscuss. As the reviewer states, the ATR-CHK1 axis controls both the intra-S and the\nG2/M checkpoint pathways during replication stress. Except for the experiment in which\nwe follow the kinetics of FANCD2 and γH2AX, throughout the entire study we focused\non the effects of Geminin and/or FANCD2 depletion at 48hs after transfection, a\ntimeframe in which there was a reduced proportion of cells in S and a high prevalence\nof cells re-replicating at the G2/M boundary (see New Supplementary Fig. 1a). During\nthe revisions period, we included the kinetics for the cell cycle profile of Geminin-depleted\nat 24 and 48hs, and we could indeed observe that at 24hs there was an increase in the\nproportion of cells incorporating EdU (see New Supplementary Fig. 1a). This supports\nthe reviewer’s argument that there is an initial activation of the intra-S checkpoint, which\nusually slows down the progression of S phase upon DNA damage. However, it seems\nthat activation of the intra-S does not impede the accumulation of re-replicating cells,\nwhich occurs during G2.\nRegarding the levels of pChk1, we provide the quantification of three independent\nreplicates of the immunoblot to show that the reduction in co-depleted cells is non-\nsignificant (New Fig. 3g and Fig. R3). Total Chk1 levels are also not significantly affected\n(please note loading control). We did not assess ATR expression since our point is to\nlook directly for the downstream effector of the checkpoint (pChk1) and quantify in\nrespect to the total Chk1 levels. As for FANCD2 depletion, we provide the quantification\nshowing that its depletion is significant (Fig. R3).\nD2 C N A F\nFig. R3. Left: Immunoblot detection of indicated proteins in U2OS cells transfected with\nindicated siRNAs. α-Tubulin, loading control. Right: Quantification of pChk1 and\nFANCD2 levels in each experimental condition. Plot shows the mean ± SD of all\nbiological replicates (n=3). In all cases: ns, non-significant, *p < 0.05, ** p < 0.01, *** p <\n0.001, **** p < 0.0001 (two-tailed t-test).\n17.Authors state “Importantly, G2-arrested cells in Geminin-depleted cells retained active\nDNA synthesis, as demonstrated by EdU incorporation and PCNA-staining, while\nexhibited high levels of DNA damage”. In Fig 3e and S3f, the authors don’t quantify the\nPCNA or DNA damage.\nWe thank the reviewer for noticing the missing information in this figure. Indeed, the plot\nwas already accounting for pH3-PCNA-EdU triple positive cells. In the revised\nmanuscript, we have corrected the labels of this plot (see New Fig. 3f). Moreover, as\nsuggested by the reviewer, we have also included new data demonstrating that G2-\narrested cells in Geminin-depleted cells which actively synthesize DNA also exhibit\nincreased DNA damage (see New Supplementary Fig. 5b).\n18.Foci intensity measurements are tricky because they are completely dependent on\nslide prep, staining and imaging consistencies. Whenever possible as in the case of\nH2AX, 53BP1, FANCD2 and in Figure 4 RPA32, it is essential to present foci/cell nucleus\nrather than image intensity.\nWe understand the point of the reviewer and we agree that usually it is better to measure\nfoci rather than intensities to reveal discrete sites of DNA breaks or ssDNA gaps.\nHowever, we believe there is a meaningful biological difference between measuring the\n)UA(\nslevel\n2DCNAF\n6\nsiCtrl siGMNN\nsiFANCD2 - + - +\nFANCD2 150kDa\npChk1 50kDa\nChk1 50kDa\nGMNN 33kDa\nαTubulin 50kDa Ctrl G M N N A N C D2 N+F\nN M G\n)UA(\nslevel\n1khCp\nns 1.5 ** *** ** **\n*** 1.0\n4\n2 0.5\n0 0\nsiRNA C G trl M N A N N C D A 2 N C D 2 siRNA F F\n+ N N M\nG\nnumber of foci and measuring the intensity of those foci. As mentioned above in comment\nnumber 6, for some markers, such as γH2AX, measuring number of foci was not possible\ndue to the pan-nuclear staining of re-replicating cells. In the case of the remaining\nmarkers such as 53BP1, FANCD2 or RPA32, we followed the reviewer suggestion by\nalways including the proportion of cells with >10 foci, but in some cases, we also provided\nthe measurement of intensity of those foci. For instance, when quantifying RPA32 foci to\nanalyze the accumulation of single-stranded DNA (ssDNA) gaps, we could see that along\nwith an increased number of RPA32 foci (New Fig. 4a and New Supplementary Fig. 6a),\nthe intensity of those foci was also higher (New Supplementary Fig. 6b). The number of\nfoci typically reflects the number of discrete sites of ssDNA where RPA proteins is bound.\nHowever, the increase in the intensity of those foci reflects the amount of RPA protein\nloaded at each site, which correlates with the extent or length of ssDNA at the given\nfocus12,13.\n19.Again, in Figure 4 there is extensive inconsistencies in measurement of RPA. Some\nexperiments use intensity measurements while others use foci/nucleus measurements.\nAuthors should stick to foci/nucleus for all measurements. In Fig 4, authors could\ndetect ssDNA foci by incubating the cells with a nucleotide analog (e.g., BrdU under\nnon-denaturing conditions) to directly track ssDNA foci, which would provide more\ncomprehensive evidence.\nAs explained above in comment number 18, we measured both the number (New Fig.\n4a and Supplementary Fig. 6a) and intensity (New Supplementary Fig. 6b) of RPA32\nfoci, as they reflect different biological meanings. Concerning the BrdU under non-\ndenaturing conditions, we thank the reviewer for this excellent suggestion. We performed\nthe recommended experiment and analyzed the number of BrdU foci under non-denaturing\nconditions in cells depleted of Geminin and/or FANCD2. Consistent with the\nresults obtained during analysis of RPA foci and DNA fibers treated with the S1 nuclease,\nGeminin depletion triggered the extensive formation of BrdU foci, and the proportion of\ncells containing a high number of these was even higher upon co-depletion with FANCD2\n(New Fig. 4b). Additionally, we implemented this technic to validate the increased\nformation of ssDNA in cells depleted of Geminin and treated with Olaparib (New\nSupplementary Fig. 6d). The results of three independent biological replicates are now\nincluded in the revised version of the manuscript together with a description of the assay\nin the methods section.\n20.In the metaphase analysis presented in Fig 4e, significance comparisons need to be\npresented for all cell lines groups. Importantly, in addition to aberrations, since re-\nreplication results in aneuploidy, this needs to be reported.\nWe thank the reviewer for the observation and apologize that we missed the statistical\ncomparisons between some conditions in initial version of the submitted manuscript. In\nthe current version, we corrected this issue and the statistical comparisons between\nevery experimental condition are shown. As for measuring aneuploidy as a readout,\nplease see in comment number 2 to this reviewer the reasons we believe this approach\nwas not the ideal and refer to the flow cytometry experiments we performed to address\nthis concern.\n21.Figure 5 is missing an immunoblot that shows loss of FANCD2 monoubiquitylation.\nIn addition, since this is a very important point, depleting Geminin in a K561R*-\nFANCD2 expressing the monoubiquitylation deficient FANCD2 should provide critical\ninsights.\nIn the revised manuscript, we included the immunoblot requested by the reviewer.\nDepletion of Geminin increases the mono-ubiquitinated fraction of FANCD2 (New Fig.\n5a). FANCA reduces FANCD2 levels, as has been reported previously14. Regarding the\nreviewer’s comment on assessing depletion of Geminin in K561R*-FANCD2 cells, we\nindeed agreed with the recommendation and performed the suggested experiments. To\nthis aim, we employed FANCD2-KO and corrected (WT and mono-ubiquitination\nresistant K561R) U2OS cells generated by Dr. van de Kooij15. Consistent with our results\ndepleting the core complex component FANCA, depletion of Geminin in K561R cells had\na decreased ability to rescue viability in FANCD2-KO cells, when compared to corrected\nWT cells (New Fig. 5e, f). We have discussed these findings in the revised manuscript\n(page 10).\n22.ChIP-qPCR experiment in S5b needs replicates to generate error bars. If this\nexperiment was just to show that the FANCD2 antibody works for ChIP, then it can just\nbe excluded from the manuscript. These fragile sites can very well be assessed from the\nChIP-seq data. It is unclear why the qPCR data is being presented.\nAs recommended, we removed the validation of FANCD2 ChIP by qPCR for a better\nvisualization of the data.\n23.The authors state “However, we noticed that FANCD2 was specifically enriched at\nreplication origins in Geminin-depleted cells when compared to control cells (Fig. 6c).”.\nWhere are origin locations being annotated from? However, even if it is truly the sites\nof origin firing, the difference between control and Geminin-depleted cells is just not\nevident from the bar graph presented. The difference seems very modest and definitely\nnot significant. Following up using a second approach to actually map origins and\ncorrelate with FANCD2 ChIP-seq data is essential to make this conclusion.\nWe acknowledge the reviewer concern and apologize if he found misleading the\nreference to the source of DNA replication origin data. The source of these data from\nK562 cells16 was initially provided with a direct link in the material and methods section.\nIn the revised version of the manuscript, we have also included the reference in the\nresults section (page 11). Regarding the concerns of the reviewer about the significance,\nin the revised manuscript we have annotated FANCD2 ChIP-seq peaks using the golden\nstandard algorithm for genome annotation, HOMER. Additional information regarding the\nanalyses has been included in the new methods section. Briefly, we built a genome\nannotation datafile with genomic features of our interest and annotate peaks to them\nusing HOMER. We now provide enrichment signal and statistical significance of such\nenrichments (New Fig. 6c, e). In addition, we used contingency tests to calculate the log₂\nodds ratio and assess the statistical significance of the differences observed between\nsiCtrl and siGMNN-treated cells (New Fig. 6c, e).\n24.The authors state “Furthermore, analysis of the functional elements of FANCD2-\nenriched genes showed a significant accumulation of FANCD2 at intergenic regions\nand introns in control and Geminin-depleted cells (Fig. 6e), consistent with FANCD2\nbeing recruited to repetitive sequences and the coding region of long genes contained\nwithin CFSs62,63”. The important thing to note is the lack of difference between control\nand Geminin depleted cells. The authors are over stating a conclusion that is yet to be\ndrawn. They need to provide a list of genes that are either long and(or) are known for\ntheir repetitive DNA content from their ChIP-seq analysis to make this statement. A\nvolcano plot depicting log fold change in genes versus log p-value or FDR could help\nwith this.\nWe thank the reviewer for this point and agree that the manuscript conclusions could\nbenefit from further computational data analysis. Thus, we repeated the annotation of\nChIP-seq peaks using HOMER algorithm with the default annotation dataset, which\nprovided enrichment and statistical information on feature enrichments (New Fig. 6c, e).\nIn addition, we used contingency tests to calculate the log₂ odds ratio and assess the\nstatistical significance of the differences observed between siCtrl and siGMNN-treated\ncells. These new data is provided in New Fig. 6e, and shows clear significant\nenrichments in promoters, 5’UTR and exons, but not in introns or intergenic regions.\n25.Based on the data presented, how can we conclude that FANCD2 enrichment at\npromoter-transcription start sites (TSSs) in Geminin depleted cells is significant.\nThanks for this comment. As mentioned before, we have repeated ChIP-seq annotation\nusing HOMER, a well-known software for genome annotation, and now provide the\nstatistical significance of genome feature enrichments in New Fig. 6d, e.\n26.The authors state “Consistent with peak analysis (Fig. 6e), more than 90% of the\norigins enriched in FANCD2 specifically in Geminin-depleted cells colocalized with\ngenes (Supplementary Fig. 5d), thus indicating these origins lie within genes.”. Where\nare these origins being mapped from? There have been no experiments conducted to\nmap origins in this study. The data is extremely correlative at best. The authors need to\nmap origins in their cell lines because this is a critical point in the manuscript.\nWe apologize if the reference to these data was found misleading. As indicated earlier,\nthe source of DNA replication origin data was provided in the material and methods\nsection. In the revised manuscript, this information can also be found in the Results\nsection with a direct reference to replication origin data mapped in K562 cells.\n27.The authors state “Thus, Geminin loss leads to FANCD2 enrichment at replication\norigins within highly transcribed genes genome-wide.”. A list of these highly transcribed\ngenes from their ChIP-seq data set would be a very important inclusion to the\nmanuscript.\nWe thank the referee for this suggestion. These data can be obtained from open access\ngenome-wide data on K562 cells as well as from the Source data files provided.\n28.The authors state “Thus, we next analyzed the replication timing of origins identified\nto accumulate FANCD2 specifically in Geminin-depleted cells. Using publicly available\ngenome-wide Repli-seq data from the ENCODE project 72, we observed a significant\nincrease in BrdU intensity during early S phase and a reduction in mid-to-late\nreplicating origins compared to control cells (Fig. 6f), suggesting these origins are\npredominantly early replicating.”. It is unclear whether the authors performed the repli-\nseq to generate the data or analyzed existing Repli-seq data from the ENCODE\nproject. It is very straightforward to check if the existing list of ERFs (PMID: 23352430\nand PMID: 36099913-for BRCA1 and BRCA2) and the longest genes in the human\nbody are both enriched for FANCD2 in the Geminin depleted cells from their ChIP-seq\ndata. Why has this not been prioritized?\nWe apologize if this was not clear. As the reviewer indicates, we used Repli-seq data\nfrom the ENCODE project17. The reference to the source of these data can be found in\nthe material and methods section, but we have further clarified this analysis in the text\n(page 12). Regarding ERFS, the analyses of FANCD2 enrichment in Geminin-depleted\ncells at ERFS were indeed using the existing list of ERFs reported in PMID: 23352430,\nas recommended by the referee. We have further clarified this in the text (page 12) and\nin the new methods section.\n29.For the Cordycepin experiments: The decreases observed in co-depleted cells\nalthough significant are very modest. Transcription inhibition by Cordycepin often shuts\ndown DNA replication as well which would be highly counterproductive for the study\nfocused on re-replication. EdU incorporation in Cordycepin treated cells needs to be\npresented to address this.\nThis is an excellent observation, and we thank the reviewer for pointing out this potential\nissue that indeed needed further attention. As suggested, we performed a new set of\nexperiments using Cordycepin or over-expressing RNAseH1 and pulse-labelling the\ncells with EdU to assess the replication capacity of these cells. We could observe that\ntreating the cells with Cordycepin indeed reduced EdU incorporation, but this was not\nthe case after over-expressing RNAseH1 (Fig. R4), supporting our idea that active\ntranscription contributes to genomic instability upon re-replication. In the current version\nof the manuscript, we have clarified that Cordycepin is a general inhibitor of transcription\nand stressed the effect after degradation of the hybrids by over-expressing RH1 (page\n13).\n3000\n2000\n1000\nC\nFig. R4. Representative images and quantification of EdU intensity in U2OS cells, after\nincubation with Cordycepin (a), or overexpression of RNAseH1 (b). ns, non-significant,\n**** p < 0.0001 (two-tailed t-test).\n30.The Model presented in Fig. 7 is too simple and doesn’t convey the message of the\npaper with any clarity.\nWe thank the reviewer for this comment. We believe that, at its current version, the model\nclearly shows the mechanism by which re-replication upon Geminin depletion leads to\nfork fragility without misleading the reader with a busy figure. In the model, we intend to\nshow that re-firing of origins promotes the accumulation of FANCD2 at re-replicating\nforks to limit their advancement, thereby preventing the accumulation of ssDNA gaps\nand collisions with the transcription machinery (left side of the model). When FANCD2 is\nnot present (right side of the model), unrestrained replisome progression results in\ntranscription-replication conflicts and the generation of post-replicative ssDNA gaps that\ncan be subsequently converted into DSBs by advancing re-replicating forks. In the\nrevised manuscript, we made our model more specific by including that re-firing origins\nare early replicating (New Fig. 7f). Additionally, we improved substantially the figure\nlegend to transmit the results of our study with clarity.\n31.The title ”FANCD2 limits unrestrained fork progression and prevents fragility of early\nreplicating sites upon aberrant origin licensing” is not very accurate. The paper does not\naddress unrestrained fork progression in the absence of FANCD2 or fragility of early\nreplicating sites by cytogenetics using FISH probes and there is definitely no measure of\naberrant origin licensing anywhere in the paper.\nWe apologize if the reviewer considers the title is not highly suitable. We would like to\npoint out however, that we demonstrated the unrestrained progression of forks in re-\nreplicating cells depleted of FANCD2 by DNA fiber assays (New Fig. 3e), as well as\nprovided a mechanism by which the uncontrolled progression of the replisome is\nassociated to increase ssDNA gaps and fork breakage (New Fig. 4). We agree with the\nreviewer that the term “aberrant origin licensing” could be misleading. Thus, following the\nreviewer recommendation, we changed the title of the manuscript to “FANCD2 restrains\nfork progression and prevents fragility at early replicating origins upon re-replication”.\n)UA(\nseitisnetni\nUdE\negarevA\n2000\n1500\n1000\n500\n0\n)UA(\nseitisnetni\nUdE\negarevA\n**** ns\nP G F H 1\nR\nTU\n.droC\nEdU DAPI\nPFG\n1HR\na b\nEdU DAPI\nMinor:\n1.Page 4: “co-depletion of CDT1 partially restored these phenotypes”. – change\nrestored to rescued\nAs requested, we changed this section of the manuscript (page 4).\n2.In Fig S1a, the cell cycle plot labels need to be bigger to match the other fonts.\nWe apologize for showing such small labels. We replaced this cell cycle plot with\ncytometry analysis of PI and EdU (New Supplementary Fig.1), as suggested by the\nreviewer (discussed in major point number 2 to this reviewer).\n3.Supplementary 2d-f say KO FANCD2 which is inaccurate. It needs to mention that they\nare PD20 patient cells completing lacking FANCD2.\nWe appreciate the correction. We replaced KO for EV (empty vector) (New\nSupplementary Fig. 2f, g) and explained that PD20 cells are derived from a Fanconi\nanemia patient deficient in FANCD2 (page 6).\n4.In every bar graph presented, the individual experimental means need to be shown as\ndots representing the 3 independent experiments.\nIn the revised manuscript, we now show the individual data points from each biological\nreplicate as dots on the bar graphs for all conditions.\n5.Significance representations and comparisons should be presented for all groups and\nnot selected ones.\nWe apologize once more for missing the statistical comparisons between some\nconditions in initial version of the submitted manuscript. As we mentioned above, in the\ncurrent version we corrected this issue and included statistical comparisons between\nevery experimental condition.\n6.Figure 3f Chk1 protein is misspelled on the gel.\nWe thank the reviewer for noticing this error. This misspelling has been corrected in the\ncurrent version of the manuscript (New Fig. 3g).\n7.Need to add a sentence introducing caffeine as a checkpoint inhibitor\nWe thank the reviewer for pointing out this omission. We added in the manuscript a\nsentence introducing Caffeine as a checkpoint inhibitor (page 8).\n8.Fig 4d needs to be sub divided into smaller figures\nAs recommended, we adjusted the New Fig. 4 and distributed the results along different\npanels for a better visualization of the presented data.\n9.Typos in figure legends need to be carefully rectified.\nWe thank the reviewer for pointing out these typos. We have thoroughly corrected them\nthroughout the Figure legends in the revised manuscript.\nReviewer #2 (Remarks to the Author):\nIn this work the authors analyse how cells deal with the deregulated licensing of origins\nof replication caused by the depletion of Geminin. Unlike previous studies, where\ndefective licensing was driving large amounts of re-replication in cancer cells, this study\nfocuses on milder levels of DNA re-replication seen in immortalised cells, such as\nhTERT-RPE1 cells.\nThrough an siRNA screen, the authors identify FANCD2 as key in suppressing DNA\ndamage and supporting cell proliferation upon Geminin depletion. They then show that\nFANCD2 loss causes faster fork speed and accumulation of S1-nuclease sensitive\nstretches of DNA, likely ssDNA gaps. Genomic analysis reveals that in Geminin depleted\ncells, FANCD2 is not only at difficult and late replicating DNA regions, but it is also seen\nat early replicating regions, in the proximity of highly transcribed genes and clusters of\nearly firing replication origins.\nSurprisingly they find that dampening of transcription or R-loops digestion significantly\nsupresses DNA damage in Geminin depleted cells.\nThis study suggests a new mechanism for the generation of genomic instability upon\nreactivation of replication origins within the same cell cycle, which now directly implicates\ntranscription and collision between the “re-replication” and the transcription machineries.\nSuch a mechanism is likely to occur in pre-malignant cells leading to tumorigenesis.\nThe experiments are well performed and controlled, combining both classical cell biology\nand genomics approaches; the results are clear-cut and important. The manuscript is\nwell written and easy to read.\nThis reviewer has few minor comments that may improve the manuscript:\n1. In supplementary Fig S1, it would be nice to show the efficiency of the co-depletion\nof Geminin and CDT1by western blot and assess if CDT1 and CDT1+Geminin\ndepleted cells are still efficiently replicating their DNA under these experimental\nconditions.\nWe thank the reviewer for this suggestion and agree that showing the efficiency of co-\ndepletion of Geminin and CDT1 was important. Based on this comment, we performed\nthe suggested experiment, and we included this western blot in the revised manuscript\n(See Supplementary Fig. 1d). As for assessing whether CDT1- or double CDT1 and\nGeminin depleted-cells are replicating their DNA, depletion of CDT1 affects initiation of\nDNA replication by preventing origin licensing. It is very challenging experimentally to\nachieve restoring normal replication by co-depletion of CDT1+Geminin . We have now\nclarified this in the results section (page 5).\n2. In many instances nuclear area is used as surrogate for re-replication. I understand\nthat such approach may not be particularly sensitive, thus creating some confusion when\nthe authors argue that geminin depletion has sometime “mild” or “no” effects in\nimmortalised cells. How does this assay compare to classical DNA content measurement\nby flow cytometry or other techniques?\nWe acknowledge the reviewer’s concern (shared also by reviewer #1 and #3) and agree\nthat another approach to measure re-replication was important to strengthen our results.\nAs discussed above in our response to reviewer #1, in the revised manuscript we\nincluded a new set of experiments using flow cytometry to examine the cell cycle\ndistribution and the prevalence of cells with DNA content above G2/M incorporating EdU\nas a metric of re-replication. Our analyses in U2OS cells show that Geminin depletion\ninduces an initial accumulation of cells in S phase at 24 hs, while there is already a small\nproportion of cells over-replicating (>4 C) when compared to control cells (New\nSupplementary Fig. 1a). However, the high prevalence of cells with DNA content above\nG2/M levels occurs at 48 hs, where about 30% of cells are re-replicating, consistent with\nthe results observed by measuring nuclear areas (New Supplementary Fig. 1b) and\nreinforcing this timepoint as the appropriate for the remainder of the study. As expected,\nthere were no evident signs of re-replication in hTERT-RPE1 cells depleted of Geminin\n(New Supplementary Fig. 1e, f). Importantly, in the revised manuscript we also included\na new set of experiments using this approach to show the effect of depleting FANCD2 in\nre-replicating cells. According to the results obtained by measuring the nuclear areas,\ndepletion of FANCD2 did not increase the levels of re-replication but rather slightly\nreduced them (New Fig. 3c). We provide cell cycle distributions for each timepoint and\ncell line from 3 independent experiments together with gates applied in the Source data.\n3. Please provide legends for Supplementary tables 1 and 2\nWe thank the reviewer for pointing out this missing information. In the revised manuscript,\nwe included the legends for Supplementary Tables 1 and 2 in Supplementary information\nas well as within the Excel tables.\n4. The siRNA screen appears to also identify proteins that, when depleted, cause DNA\ndamage which is prevented by geminin depletion e.g. PLK3, NEK1, CDK7 and few\nothers. If this is correct, can the authors mention and discuss it?\nThe proteins the reviewer is referring to are scoring high in the parental control but not\nin the Geminin-depleted category (New Supplementary Table 2), and could indeed reflect\ninhibition by Geminin, though we have not validated or further investigated this category.\nTo avoid confusion, in the revised manuscript we have clarified the criteria of selection\nof the screening and the information about the hits in the result section (page 5).\n5. Fig 3C: does siFANCD2 increases fork speed compared to siCtrl?\nWe thank the reviewer for noticing this missing information in the initial version of the\nsubmitted manuscript. In the current version, we corrected this issue throughout our\nanalyses and included the statistical comparisons between every experimental condition.\nIndeed, loss of FANCD2 slightly, although significantly, increased fork speed (New Fig.\n3e), suggesting that it acts as a constitutive regulator of replication forks even during\nunperturbed conditions. Accordingly, although it was non-significant, we could also\nobserve a small increase in the percentage of new origins in cells depleted of FANCD2\n(New Fig. 3e) as observed previously18.\n6. Line 211 and Fig S3c-d: it is not clear why low doses of ATR inhibitors suppress re-\nreplication. It is known that ATR inhibition facilitates origin activation, thus I would expect\nthe opposite phenotype. Is this a dose-dependent effect and is it related to the override\nof mitotic checkpoint thus not leaving sufficient time for large quantities of DNA to\naccumulate? (see also point 2 related to sensitivity of the nuclear area assay).\nIndeed, we believe that what we observe is an override of the mitotic checkpoint, thus\nnot leaving sufficient time for re-replication in G2 and have now clarified this in the revised\nmanuscript (page 8). At the concentration used, this ATR inhibitor has been reported to\npromote replication activity19, and therefore we do expect certain activation of origins.\nHowever, we believe we are not able to see the consequences of increased firing on re-\nreplication nor DNA damage because ATR inhibition also abrogates cell cycle arrest at\nG2/M, and “pushes” re-replicating cells to prematurely enter mitosis before large\namounts of DNA can re-replicate. This is supported by suppression of Chk1\nphosphorylation, together with a decrease in the nuclear size and loss of DNA damage\nsignaling (New Supplementary Fig. 5c, d), while we see an elevated number of\nmicronuclei (New Supplementary Fig. 5e).\n7. Line 337: how effective is Cordycepin in inhibiting general transcription and in\nparticular in inhibiting transcription of the genes located in the proximity of clusters of\nearly firing origins?\nWe thank the reviewer for drawing our attention to this question. To assess the effect of\nCordycepin in transcription, we decided to use 5-ethynyl uridine (EU) incorporation as a\nproxy to measure global nascent RNA synthesis. As expected, the treatment with\nCordycepin led to a dramatic reduction in the mean fluorescence intensity of incorporated\nEU (Fig. R5). Given the potent global inhibition of transcription demonstrated with the\nEU staining data, and since the effect of Cordycepin is not dependent on any specific\nDNA sequence, we believe that inhibition occurs broadly across the genome, including\nat the loci specified by the reviewer.\n****\n800\n600\n400\n200\nFig. R5. Representative images and quantification plot of U2OS cells treated with\nCordycepin and incubated with EU. Plot shows the distribution of individual values of a\nrepresentative replicate. DNA staining, Hoechst. Scale bar, 10μm. **** p < 0.0001 (two-\ntailed t-test).\n8. Similarly, would Cordycepin treatment or RNAse1 overexpression also suppress\nFANCD2 binding at siGEMNN specific peaks determined in fig 6b?\nWe thank the reviewer for this comment. It is known that FANCD2 binds to stalled forks\nand R-Loops, which stimulates its mono-ubiquitination. We have now provided additional\ndata showing that FANCD2 is present at replication forks in re-replicating cells,\nemploying SIRF and Proximity Ligation Assays (see point 13 to Reviewer 1 above), and\nhave shown its mono-ubiquitination upon Geminin depletion. We have also provided\nstatistical evidence of FANCD2 enrichment at the vicinity of origins and promoter regions\nin re-replicating cells (see New Fig. 6c, e, and point 23 to Reviewer 1 above). Our results\nshowing that incubating Geminin-depleted cells with Cordycepin or overexpressing\nRNase H1 reduces the number of FANCD2 foci (see New Fig. 7d, e) suggests that\nrecruitment of this protein is at least partially dependent on the presence of active\ntranscription and the accumulation of R-loops, as previously shown20. Thus, taken\ntogether, our data strongly suggest that FANCD2 occupancy at the sites determined by\nour ChIP analyses would be reduced by Cordycepin/RNaseH, though we have not\ndirectly addressed this point. These points are discussed in the revised manuscript\ndiscussion.\nReviewer #3 (Remarks to the Author):\nInitiation of eukaryotic DNA replication is tightly controlled and coordinated with cell\ncycle progression to ensure only a single round of DNA synthesis. In metazoa this\noccurs primarily through careful regulation of CDT1 and Geminin levels. Defective\nregulation of CDT1/Geminin can lead to re-replication of the genome which is\nmutagenic, toxic, and is implicated in oncogenesis. However, the exact molecular\nconsequences of CDT1/Geminin dysregulation are unclear. Badra-Fajardo et al have\ninvestigated the\nconsequences of Geminin dysregulation and identified an important role for FANCD2 in\npreserving genome stability. They find that loss of FANCD2 causes DNA damage,\nchromosomal aberrations, and possibly cell death in Geminin-deficient cells.\nMechanistically, these consequences are proposed to arise from accumulation of single-\nstranded DNA gaps and formation of DNA breaks that are a downstream consequence\nof re-replication. Furthermore, these effects may be unequally distributed throughout the\ngenome. Overall the authors have connected FANCD2 to defects in Geminin function\nbut there are major concerns about the mechanism proposed, the interpretation of the\ndata, and the overall novelty of FANCD2 function supported by the data.\nMajor concerns\n1. The manuscript seems to imply a unique role for FANCD2 in responding to Geminin\nloss. However, the upstream (FANCA in Fig 5) and downstream (RAD51, BRCA1,\nBRCA2 in Fig 1) components of the Fanconi Anemia pathway behave similarly. It would\nseem that the role described for FANCD2 here reflects the Fanconi Anemia pathway’s\nfunction and not something more unique about FANCD2. Because a major role of the\nFA pathway is to coordinate DNA breaks in a controlled manner (by XPF-ERCC1) this\nwould seem to contradict the model that FANCD2 is preventing breaks. The data\nwould certainly support a model that FANCD2 prevents break accumulation, but not\nthat it prevents break formation.\nWe would like to thank the reviewer for this comment. It is indeed correct that some repair\nproteins, such as BRCA1/2, were found during our primary screen. However, we believe\nthat activation of these proteins after Geminin depletion, as it occurs with FANCD2, is\nassociated to their function during fork stability and not directly with inducing breaks in a\nregulated fashion, as is the case upon inter-strand crosslinks. If the observed FANCD2\nfunction during re-replication would be to promote DNA breaks (by XPF-ERCC1), we\nwould expect reduced DNA damage upon its depletion, which is not the case, as\nobserved by increased γH2AX or 53BP1 (Fig. 1c, Supplementary Fig. 2).\nIn the revised manuscript, we have performed a number of additional experiments to\nreinforce our data showing FANCD2 recruitment prior to formation of DNA breaks. Along\nwith the kinetics of FANCD2 and γH2AX in Geminin-depleted cells (New Fig. 2c), we\nincluded new results from a synchronization experiment in Geminin-depleted U2OS cells\n(see New Fig. 2.d and New Supplementary Fig. 4b). During the synchronization, we\ndepleted Geminin to ensure S-phase entry with no Geminin expression, and then\nreleased and subsequently monitored EdU incorporation, together with FANCD2 and\npH3, over a 30hs time course. Depletion of Geminin led to a time-dependent\naccumulation of cells that were actively synthesizing DNA at the G2/M boundary (EdU\nand pH3-double positive) (New Fig. 2d and New Supplementary Fig. 4b), sustaining the\nidea that FANCD2 recruitment occurs early and precedes high levels of DNA breaks.\nAdditionally, to strengthen our results showing that FANCD2 binds to chromatin upon re-\nreplication at replication forks, in the revised manuscript we included a new set of data\nemploying the SIRF assay, which combines EdU labeling of nascent DNA with a\nproximity ligation assay (PLA) to detect proteins at newly synthesized DNA9. Using this\napproach, we observed a significant increase in the number of FANCD2-EdU interacting\nfoci following Geminin depletion (New Fig. 3a). These results were also validated through\nthe analysis of other replisome factors by PLA (Fig. 3b and Supplementary Fig. 4c),\nwhich showed that this interaction is not only increased in S-phase cells but becomes\nevident in G2-arrested cells (New Fig. 3b), thus confirming that FANCD2 is directly\nrecruited at forks upon re-replication. We hope that the reviewer will now agree that our\nnew data strongly support our initial conclusions of a direct function of FANCD2 in\nprotecting re-replicating forks.\n2. It is well established that cancerous cells are much more readily undergo re-replication\nfollowing Geminin/CDT1 dysregulation. hTERT-RPE1 cells are examined in Figure 1 but\nthe remainder of the manuscript focuses on cancerous cells (U2OS, MCF7, 293T,\nHCT116). It’s unclear how much of the observed effects are specific to cancerous cells.\nWe thank the reviewer for this comment. We indeed mainly focused on U2OS cells to\naddress the molecular mechanism by which FANCD2 promotes the survival of re-\nreplicating cells. However, we have also provided validation of critical phenotypes in non-\ncancerous cell lines, such as the DNA damage phenotypes and the clonogenic assays\nin hTERT-RPE1 and the patient-derived PD20 cells (New Fig. 1 and New Supplementary\nFig. 2f, g). In the revised manuscript, we strengthen these data by including the analysis\nof additional signatures of genomic instability upon double-Geminin and FANCD2\ndepletion in hTERT-RPE1 and PD20 cells (See New Supplementary Fig. 6f, g), which\nshows that the observed effects are not only specific to cancerous cells.\n3. The manuscript focuses on re-replication but this is only directly measured once (Fig.\nS1A). For the remainder of the manuscript, nuclear size is used as a proxy. Not only is\nnuclear size very indirect but it is unclear whether it serves a reasonable proxy: in Fig\nS1A the fraction of cells that exhibit increased nuclear area is much greater than the\nfraction of cells that exhibit >4n DNA content. The lack of direct measurements of re-\nreplication and concerns about the suitability of cell size as a proxy for re-replication\ncall into question most of the authors’ conclusions that relate to re-replication.\nWe acknowledge the reviewer’s concern (shared also by reviewer #1 and #2) and agree\nthat another approach to measure re-replication was important to strengthen our results.\nAs discussed above, in the revised manuscript we included a new set of experiments\nusing flow cytometry to examine the cell cycle distribution and the prevalence of cells\nwith DNA content above G2/M incorporating EdU as a metric of re-replication. Our\nanalyses in U2OS cells show that Geminin depletion induces an initial accumulation of\ncells in S phase at 24 hs, while there is already a small proportion of cells over-replicating\n(>4 C) when compared to control cells (New Supplementary Fig. 1a). However, the high\nprevalence of cells with DNA content above G2/M levels occurs at 48 hs, where about\n30% of cells are re-replicating, consistent with the results observed by measuring nuclear\nareas (New Supplementary Fig. 1b) and reinforcing this timepoint as the appropriate for\nthe remainder of the study. As expected, there were no evident signs of re-replication in\nhTERT-RPE1 cells depleted of Geminin (New Supplementary Fig. 1e, f). Importantly, in\nthe revised manuscript we also included a new set of experiments using this approach\nto show the effect of depleting FANCD2 in re-replicating cells. According to the results\nobtained by measuring the nuclear areas, depletion of FANCD2 did not increase the\nlevels of re-replication but rather slightly reduced them (New Fig. 3c). We provide cell\ncycle distributions for each timepoint and cell line from 3 independent experiments\ntogether with gates applied in the Source data.\n4. Geminin knockdown is likely to have effects other than re-replication of the genome.\nThe mechanism of re-replication by geminin is loading of additional MCM complexes\nfollowed by origin firing on the already-replicated DNA. There is no reason this could not\nalso occur in the unreplicated region of the genome, which could manifest as\nunscheduled/excess origin firing and may account for some of the effects observed\n(e.g. PMID29466339). Taken together with point (1), above, it seems very possible the\nauthors are documenting breaks induced by replication-transcription collisions and\nrepaired by the Fanconi Anemia pathway.\nAs the reviewer rightly points out, ectopic licensing and thus recruitment of additional\nMCMs in Geminin-depleted cells is likely to affect un-replicated regions of the genome\nand increase transcription-replication collisions at those sites. Indeed, we performed\nadditional analysis and observed that Geminin loss increases origin firing (New Fig. 3e),\nsupporting that both re-firing as well as ectopic firing of origins take place when Geminin\nis inhibited. According to these findings, we demonstrate that inhibiting active\ntranscription or eliminating R-loops alleviates the phenotypes exhibited in Geminin\ndepleted cells (Fig. 7d, e). However, as we discussed in comment number 1 to this\nreviewer, our data strongly suggest that FANCD2 is not recruited following the break, but\nat re-replicating forks as a mean of stabilizing and limiting fork progression and thus\nprotecting them from breaks. This idea is supported by the kinetics of FANCD2 and\nγH2AX in Geminin-depleted cells (New Fig. 2c), together with the new set of experiments\nin synchronized cells (New Fig. 2d) and those employing SIRF and PLA assays, in which\nwe observed a significant increase in the number of FANCD2-EdU, as well as PCNA-\nFANCD2 and MCM2-FANCD2, interacting foci following Geminin depletion when\ncompared to controls (New Fig. 3a, b and Supplementary Fig. 4c).\n5. The is an ~24 hour delay between FANCD2 and gamma H2AX focus formation (Fig.\n3A). Because the cell cycle in U2OS cells (used in Fig 3A) is approximately 24 hours it\nwould seem very unlikely that the FANCD2 levels and damage are both occurring\nwithin S phase or G2, as would seem to be implied by the model that FANCD2 protects\nforks from excessive breakage. The H2AX foci would at the very least need to arise\nfrom the following S phase, but the simpler explanation would be that they arise from\nbreaks during mitosis.\nOur results show that, at least in the context of re-replication, DNA breaks do not arise\nupon mitosis but rather accumulate extensively when re-replicating cells are arrested at\nG2/M. These results are supported by the new set of data provided from flow cytometry\nanalysis of EdU and PI (see New Supplementary Fig. 1a) and synchronization\nexperiments (see New Fig. 2d); Geminin depletion induces an initial accumulation of\ncells in S phase within 24hs, while there is a small proportion of cells over-replicating.\nThe accumulation of cells in S probably reflects the activation of an intra-S checkpoint\nthat slows cell cycle progression and limits the proportion of re-replicating cells. This is\nconsistent with an early detection of FANCD2 foci but not γH2AX during the kinetics\nexperiment (New Fig. 2c). Over time, we observe how re-replicating cells continue\naccumulating at the G2/M boundary, showing a prevalence of about 30% of cells\nexhibiting >4C DNA content after 48hs (New Supplementary Fig. 1a, b). Importantly, this\npercentage highly fits with the proportion of cells showing increased nuclear area and\nDNA breaks in the kinetics of FANCD2 and γH2AX for asynchronous cells (New Fig. 2c),\nas well as phospho-Histone H3 and checkpoint activation (Fig. 3f and Supplementary\nFig. 4b) , as expected if cells re-replicate while being arrested at the G2/M. Consistently,\nabrogation of the checkpoint, which is expected to lead to entry into mitosis, resulted in\ndecreased re-replication, reduced DNA damage and aberrant mitotic figures (New\nSupplementary Fig. 5). These results support our model in which DNA break do not arise\nfrom mitosis but from massive re-replication after cells arrest at G2/M.\n6. In several figures the authors report increased number of foci of different DNA damage\nresponse proteins following Geminin knockdown (Figs 1c, 2d, 3a, 4a, 4d, 5a, 5d, 7d-e).\nBecause Geminin increases nuclear area there would be expected to be some increase\nin foci per nucleus independent of any effects on DNA damage. The same issue may\nalso be true for nuclear signal intensity (depending on how it was calculated). As the data\nstands, many of the increases in foci (and possibility nuclear signal) could be a\nconsequence of increased nuclear size.\nWe thank the reviewer for drawing our attention to this issue. We agree that certain\nincrease in nuclear intensity or the number of foci of DNA damage proteins is expected\nas the nuclear size becomes enlarged during re-replication. It should be noted, however,\nthat the highest levels of DNA damage are observed upon co-depletion of FANCD2,\nwhich does not increase, but rather reduces the nuclear size of re-replicating cells (Fig.\n3c, d and Supplementary Fig. 5a). Nevertheless, we acknowledge that measuring\nintensity per nuclear area was not the ideal approach to account for the levels of DNA\ndamage. Thus, in the revised manuscript, we replaced every analysis on nuclear\nintensities to raw mean intensities, a metric independent of the size of the nucleus.\nRegarding the number of foci, for the different markers assessed in our work, we re-\nanalyzed some of our data by correcting the number of foci in respect to the area.\nAs shown in Fig. R6, the results were comparable to those obtained by measuring the\nnumber of foci per nucleus, thus excluding the size of the nucleus as the reason for the\nobserved differences. In the manuscript, we use as a metric the percentage of cells with\nmore than 10 foci, as this is the metric that is most commonly used in the field.\n40\n30\n20\n10\noligos. *** p < 0.001, **** p < 0.0001 (two-tailed t-test).\n7. Similar to the concerns about nuclear size, re-replication is expected to result in\nmore DNA per nucleus and thus an increase in replication proteins on DNA (E.g. RPA\nin Fig 4A-B). The same would also be true for DNA damage foci because more\nreplication would increase the level of spontaneous DNA damage. It is thus not clear\nhow meaningful increased DNA replication/repair foci are under conditions where re-\nreplication may be taking place.\nAs mentioned above, we have addressed this question to rule out the possibility that the\neffects observed on DNA damage are simply due to increased nucleus/more DNA/more\nreplication. The differences we observe are large and highly significant and do not\nchange when correcting for nuclear size. In addition, we provide a number of other\nmetrics (such as for example intensity of single RPA foci and native BrdU staining– see\nresponse to Reviewer 1 above) and a number of orthogonal approaches (such as SIRF,\nPLA, DNA fiber analysis and ChIP seq data) which further strengthen our conclusions.\n8. The genome wide distribution of FANCD2 presented in Figure 6 may not be\nparticularly meaningful because there is no correction for the prevalence of different\ngenomic features. For example, the prevalence of FANCD2 peaks at intergenic regions\nand introns in Fig 6E could simply be because this constitutes most of the genome.\nSimilarly, the approximately 50% overlap between FANCD2-bound genes in Fig. 6D is\nvery similar to what would be expected for any two randomly selected sets of genes.\nAs things stand, many of the conclusions drawn from genomic analysis may not be\nmeaningful.\nWe thank the reviewer for this comment. Indeed, the manuscript benefited from further\ndata analysis. We have now reevaluated feature enrichment using HOMER, a golden\nstandard for genome annotation, and now provide the significance of these enrichments\n(New Fig. 6c, e). According to these results, , enrichment at introns and intergenic\nregions is not significant. while significant enrichment is observed at promoters, 5’UTR\n)%(\nicof\n1PB35\n01\n>\nslleC\n***\n****\nsiRNACtr G l M N F N A N N C + A FA N CA N M G\nsuelcun/icof\n1PB35\n**** ****\n40 **** 0.25 ****\n0.20\n30\n0.15\n20\n0.10\n10\n0.05\n0 0.00\nsiRNACtr G l M N G F N M A N N N C + A FA N CA siRNACtr G l M N G F N M A N N N C + A FA N CA\nN M G\nG\naera/icof\n1PB35\nand exons (New Fig. 6e). We have corrected this part of the text and further discuss our\nfindings (page 11).\n9. The effects of Geminin and FANCD2 knockdown on cell survival seem to be purely\nadditive (Fig. 1D), in contrast to the effects on the DNA damage response which are\nclearly synergistic (Fig. 1C). The DNA damage data fit with the authors’ model of\nFANCD2 responding to Geminin knockdown but the viability data do not. It is not clear\nwhether Fig 1D supports a role for the authors’ proposed mechanism in ensuring cell\nviability.\nWe thank the reviewer for raising this question. The effects on DNA damage among\ndouble-depleted and single depleted cells are indeed clearly synergistic along the study.\nDifferences in cell viability are clearly synergistic in normal cells, but it may be difficult to\nassess synergy by eye, in cancer cells, as Geminin depletion alone already triggers\nsignificant cell death . To address this, in the revised manuscript we prove the synergistic\neffect on viability by employing the Bliss independence model21. The model calculates\nan expected cell survival by multiplying the survival rates of the single knockdowns and\nsynergy is shown when the observed survival is significantly lower than the predicted\nvalue. This analysis was applied for each clonogenic performed along the study and we\nfound positive synergy scores. The synergy scores retrieved, together with the statistical\nsignificance are included in the Source data of the revised manuscript.\nMinor concerns\n10. The legend for Figure 1 refers to panel c but appears to actually be referring to b.\nWe thank the reviewer for noticing this error. We have now corrected this issue and\nthoroughly revise the Figure legends.\n11. For Figure 2A it is not obvious that Geminin knockdown causes increased FANCD2\nrecruitment to chromatin. The difference is modest and total cellular FANCD2 is\nincreased so the increased binding could just reflect higher protein levels.\nWe thank the reviewer for this observation. In the revised manuscript, we included the\nquantification of this chromatin fractionation experiment (New Fig. 2a).\n12. It is unclear what the asterisk in Fig 2C corresponds to.\nWe apologize for this omission. The asterisk refers to the appearance of a common\nunspecific band when we blot against CDT1. In the revised manuscript, we have included\nthis information in the figure legend of the corresponding figure (New Supplementary Fig.\n3a).\n13. The analysis of Cbio data in Fig. 2E cannot be used to draw a causal conclusion\nbecause this data only demonstrates correlation. Also it is unclear how this analysis\nconstitutes n=3 as the legend would seem to indicate.\nWe agree with the reviewer that the analysis of FANCD2 expression in tumors with high\nvs low CDT1 expression presented only demonstrates a correlation. In the revised\nmanuscript, we moved the entire section regarding mild CDT1-over expression to\nSupplementary information (see New Supplementary Fig. 3) and modified the text to\navoid confusion (page 7).\n14. When discussing Figure 5 the authors conclude that Geminin loss “promotes\nactivation of canonical FA signaling and recruitment of monoubiquitinated FANCD2”.\nThey did demonstrate a role for FANCA following Geminin knockdown and can infer\nimportance of the FA pathway following Geminin knockdown, but they cannot infer any\naltered signaling (especially because they did not measure signaling).\nWe acknowledge the reviewer’s point that based on the data presented, there was no\nevidence of altered FA signaling upon Geminin depletion. In the revised manuscript, we\nincluded an immunoblot showing an increase in the mono-ubiquitinated fraction of\nFANCD2 upon Geminin depletion (New Fig. 5a). FANCA depletion reduces total levels\nof FANCD2, as observed during assessment of FANCD2 foci in FANCA-depleted cells\n(New Fig. 5a) and reported previously14.\n15. For Fig 7d-e how do the authors know that the RPA signal is due to gaps and not\nsingle-stranded DNA arising directly from R-loops?\nWe thank the reviewer for this insightful question that has indeed been a matter of debate\nin the field. So far, there is no solid demonstration that RPA can bind to the displaced\nssDNA of the R-loop. Although it is possible that RPA could bind to the hybrid, we believe\nthat formation of large and intense RPA foci upon Geminin depletion are consistent with\nlong stretches of ssDNA that are usually generated upon post-replicative DNA gaps. The\ndisplaced strand within a canonical R-loop structure is often short and constrained,\nmaking it a less favorable substrate for the stable assembly of large RPA filaments.\nDuring the revision period, we performed additional experiments and analyzed the\nnumber of BrdU foci under non-denaturing conditions in cells depleted of Geminin and/or\nFANCD2. Consistent with the results obtained during analysis of RPA foci and DNA fibers\ntreated with the S1 nuclease, Geminin depletion triggered the extensive formation of\nBrdU foci, and the proportion of cells containing a high number of these was even higher\nupon co-depletion with FANCD2 (new Fig. 4b). Additionally, we implemented this technic\nto validate the increased formation of ssDNA in cells depleted of Geminin and treated\nwith Olaparib (new Supplementary Fig. 6d). Since the ssDNA derived from the displaced\nstrand of an R-loop is, by definition, parental DNA, it would not incorporate BrdU. We\nrecognize that it is possible that certain BrdU incorporated during S-phase could later\nappear in the displaced strand of an R-loop formed on the newly synthesized DNA but\nthis is highly unlikely due to the acute phenotype observed. Thus, we believe that the\nvast majority of the RPA foci we observe is a direct result of persistent post-replicative\ngaps, consistent with our DNA fiber analysis. The results of three independent biological\nreplicates for the BrdU assay are now included in the revised version of the manuscript\ntogether with a description of the assay in the methods section.\n16. In the results section, the authors state “Finally, we tested whether increased\nformation of ssDNA gaps in Geminin and FANCD2 double-depleted cells would result in\nchromosome fragility and high levels of genomic instability.” There is no direct\nmeasurement of gaps and the intervention (Geminin depletion) is highly indirect so they\ncannot make this conclusion.\nAs detailed above in comment number 15, we believe we demonstrate that Geminin\ndepletion, as well as Geminin and FANCD2 double-knockdown, promotes the\naccumulation of ssDNA gaps through the analysis of RPA foci, DNA fibers treated with\nthe S1 nuclease and by a new set of experiments included in the revised manuscript\ndemonstrating increased number of BrdU foci under non-denaturing conditions (see New\nFig. 4b). These metrics have been widely used in the DNA damage field to link\nuncontrolled replication to gap formation, DNA damage accumulation and chromosomal\ninstability5,18,22. Moreover, we also demonstrate a direct effect of Geminin loss on the\nnumber of chromosome breaks and aberrations, which is exacerbated after co-depleting\nFANCD2 (see New Fig. 4f). Nonetheless, we rephrased this section of the manuscript to\nbetter describe the results obtained (page 10).\n17. For Fig S1A the graphs of cell counts should be labeled to indicate which is control\nvs which is Geminin depleted\nWe thank the reviewer for pointing out this observation. As previously mentioned, we\nreplaced this figure with new results from flow cytometry analyses as recommended by\nthe reviewers (see New Supplementary Fig. 1a, e).\nReviewer #4 (Remarks to the Author):\nI co-reviewed this manuscript with one of the reviewers who provided the listed reports.\nThis is part of the Nature Communications initiative to facilitate training in peer review\nand to provide appropriate recognition for Early Career Researchers who co-review\nmanuscripts.\nAdditional references\nhttps://doi.org/10.1038/s41467-021-23835-0\nreplication reveals inhibitory controls that target multiple stages of replication initiation.\nMol Biol Cell 17, 2415-2423 (2006). https://doi.org/10.1091/mbc.e05-11-1037\ngene amplification. Science 329, 943-946 (2010).\nhttps://doi.org/10.1126/science.1190966\ncentrosome abnormalities. Cancer Genet Cytogenet 144, 91-99 (2003).\nhttps://doi.org/10.1016/s0165-4608(02)00929-9\nPromotes Alternative End-Joining DNA Repair. Cell Rep 15, 2488-2499 (2016).\nhttps://doi.org/10.1016/j.celrep.2016.05.031\ninhibitor resistance. Nat Commun 12, 3636 (2021). https://doi.org/10.1038/s41467-021-\n23463-8\nBRCA-deficient tumours. Nat Commun 15, 4430 (2024). https://doi.org/10.1038/s41467-\n024-48715-1\ninhibitors. Nature 628, 433-441 (2024). https://doi.org/10.1038/s41586-024-07217-2\nNascent DNA Replication Forks. Bio Protoc 9, e3377 (2019).\nhttps://doi.org/10.21769/BioProtoc.3377\nprevent R-loops that cause genome instability. Embo j 36, 3532-3547 (2017).\nhttps://doi.org/10.15252/embj.201797208\nremodeling complex helps resolve R-loop-mediated transcription-replication conflicts.\nNat Genet 53, 1050-1063 (2021). https://doi.org/10.1038/s41588-021-00867-2\nof RPA. Cell 155, 1088-1103 (2013). https://doi.org/10.1016/j.cell.2013.10.043\nprocessing limits DNA damage-induced mutagenesis. Nucleic Acids Res 53 (2025).\nhttps://doi.org/10.1093/nar/gkaf198\nactivation of s phase checkpoint signaling. Mol Cell 51, 678-690 (2013).\nhttps://doi.org/10.1016/j.molcel.2013.07.023\n15 van de Kooij, B. et al. The Fanconi anemia core complex promotes CtIP-dependent end\nresection to drive homologous recombination at DNA double-strand breaks. Nat Commun\n15, 7076 (2024). https://doi.org/10.1038/s41467-024-51090-6\ncombinations of chromatin marks in human cells. PLoS Genet 10, e1004282 (2014).\nhttps://doi.org/10.1371/journal.pgen.1004282\nhuman replication timing. Proceedings of the National Academy of Sciences 107, 139-\n144 (2010). https://doi.org/10.1073/pnas.0912402107\nreplication stress and genome instability in cells lacking BRCA2. Nat Struct Mol Biol 23,\n755-757 (2016). https://doi.org/10.1038/nsmb.3252\ntopoisomerase i inhibitors by disabling DNA replication initiation and fork elongation\nresponses. Cancer Res 74, 6968-6979 (2014). https://doi.org/10.1158/0008-5472.Can-\n13-3369\nremodeling complex helps resolve R-loop-mediated transcription–replication conflicts.\nNature Genetics 53, 1050-1063 (2021). https://doi.org/10.1038/s41588-021-00867-2\n26, 585-615 (1939). https://doi.org/https://doi.org/10.1111/j.1744-7348.1939.tb06990.x\ntherapy resistance primarily through its role in homology-directed repair. Mol Cell 84, 447-\n462.e410 (2024). https://doi.org/10.1016/j.molcel.2023.12.025\nFANCD2 restrains fork progression and prevents fragility at early replicating origins upon\nre-replication\nWe are grateful to the reviewers for their thorough evaluation of our manuscript and for providing\nvaluable feedback. We have fully addressed all points raised by conducting new experiments and\nrevising the text to enhance clarity and strengthen our conclusions throughout the manuscript.\nPoint-by-point response to the reviewers' comments:\nReviewers' comments:\nReviewer #1 (Remarks to the Author):\nThe inclusion of this analysis and discussion (new Fig. 3e, page 8) satisfactorily addresses my\nearlier comment about % origin activation. That said, my original comment also highlighted the\nabsence of inter-origin distance analysis, which remains unaddressed. If the authors are unable\nto perform this analysis at present, I would encourage them to clarify this limitation in the\ndiscussion.\nWe thank the reviewer for their careful reading and valuable feedback. We agree that an analysis\nof inter-origin distances would provide a more complete picture of replication dynamics. We want\nto clarify that our experimental approach, DNA fiber analysis, does not allow for a reliable\nmeasurement of inter-origin distances. Unlike DNA combing, DNA spreading results in a non-\nuniform distribution of DNA molecules on the slides, which makes a reproducible measurement\nof the distances between origins technically difficult. Given this limitation, we focused on the\npercentage of origin firing as a metric to address the reviewer's initial comment. We believe this\nprovides a meaningful and reproducible measure of replication origin activity given our\nexperimental setup. Following the reviewer’s suggestion, we added a sentence to the Results\nsection (Page 8) to explicitly state this limitation.\nI thank the authors for their detailed clarification. From the new data, it is clear that Geminin loss\nmodestly reduces clonogenic survival in normal cells, has a stronger effect in cancer cells, and\nthat co-depletion with FANCD2 further sensitizes cells. I also appreciate the point that no\nsignificant apoptotic cell death is detected at the early time points used for molecular pathway\nanalyses, and that the observed phenotypes are therefore not secondary to acute cytotoxicity.\nFor clarity, I would encourage the authors to explicitly state this distinction in the main text:\nnamely, that Geminin loss and co-depletion with FANCD2 impair long-term clonogenic survival\nbut do not induce early apoptotic cell death. This would help the reader reconcile the survival and\nmechanistic data more readily. I would still urge the authors to test apoptotic population using\nFACS for AnnexinV+ population.\nWe thank the reviewer for the suggestion and agree that clarifying this question was important.\nFollowing their recommendation, we used cleaved Caspase-3 and AnnexinV+ as markers of\napoptotic cell death. Our analysis shows that neither single- nor double-knockdowns trigger\napoptosis (see new Supplementary Fig. 2c), supporting our conclusion that Geminin loss and its\nco-depletion with FANCD2 impair long-term clonogenic survival without inducing early-stage\napoptosis. We have included these results and have clarified this distinction in the main text of\nthe revised manuscript (New Supplementary Fig. 2c and page 6 of the text).\nThe data from the Aux-degron system is consistent with siRNA approaches and is addressed by\nthe authors. However, they don’t address my previous comment about using pharmacological\ninhibitors. The rationale for not using pharmacological inhibitors needs to be clarified.\nWe thank the reviewer for their valuable feedback and apologize for not addressing this point in\nour initial revision. We are indeed working with the pharmacological inhibitor AF615 to target the\nCDT1/Geminin protein complex1. However, we would like to note that the mechanism of action of\nthis inhibitor is not yet fully characterized, and it is beyond the scope of the current manuscript.\n[REDACTED]\nreplicate from two independent experiments (n=2). c. Representative images and quantification\nof long-term clonogenic assays in U2OS and hTERT-RPE1 cells treated with 15μM of AF615.\nPlot shows the mean ± SD of all biological replicates (n=3). In all cases: ns, non-significant, *p <\n0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001 (two-tailed t-test).\nI thank the authors for their detailed response and the addition of SIRF and PLA experiments.\nThat said, I would encourage the authors to clearly acknowledge in the discussion that while\nFANCD2 recruitment precedes extensive DNA breakage, low-level DNA damage signaling may\nstill contribute to its localization in G2. However, I would like clarification on how the authors\ndistinguish between G2 re-replicating cells and late S-phase cells. The response indicates this\nwas based on EdU incorporation patterns, but such patterns can also be observed in late S. Were\nadditional markers (e.g., Cyclin B1) employed to confirm G2 identity? Without such controls, it is\ndifficult to unambiguously assign these cells to G2 rather than late S-phase.\nWe thank the reviewer for this well-taken comment. In the revised manuscript, we have clarified\nthat re-replication upon Geminin loss begins within the first round of DNA synthesis and induces\nFANCD2 recruitment to active forks prior to G2/M arrest and high levels of DNA damage (page\n8). We have also acknowledged in Discussion that DNA damage signaling could contribute to\nFANCD2 localization in G2 (Page 14). Regarding the question of the reviewer on how we\ndistinguish between G2 re-replicating cells and late S-phase cells, we would like to clarify that we\nused phosphorylated histone H3 (pH3) as a marker of G2/M cells. As demonstrated in our\nsynchronization experiment (Fig. 2d, e, and Supplementary Fig. 5c, d), we can reliably identify re-\nreplicating cells in G2 by three concomitant features: increased nuclear size, a diffused EdU\nstaining pattern (that indicates DNA synthesis), and positive pH3 staining (confirming the G2/M\nphase). This combined approach provides strong evidence that the re-replication events we\nobserve are indeed occurring in the G2 phase. In addition, re-replicating cells are distinguishable\nfrom normal S-phase cells as they show a characteristic diffused EdU pattern which is distinct\nfrom all S-phase EdU patterns (including late S-phase cells which exhibit a small number of large\nEdU foci). We have now further explained the synchronization experiment and the observed\nphenotypes in the revised manuscript (page 7 of main text).\nReviewer #2 (Remarks to the Author):\nUpon revision, this manuscript is greatly improved and the authors have satisfactory addressed\nmy comments. The inclusion of additional experiments by flow cytometry nicely complements the\nmeasurements of nuclear area as surrogate marker for DNA re-replication. DNA fibre analysis\ngives a good understanding on replication dynamics in presence and absence of Geminin and\nFANCD2 or both, and is consistent with the idea that damage occurs from re-replicating forks.\nAdditional comments:\n• Line 321: should be: depletion (and not co-depletion) of FANCA reduces the levels of FANCD2.\nWe thank the reviewer for noticing this error, which has now been corrected.\n• Fig 5f: it would be useful to indicate in the labelling of this panel the that the experiments were\ndone in FANCD2-KO cells unlike in all the other panels in the same figure.\nWe thank the reviewer for the suggestion. In the revised manuscript, we have added a label\nspecifying these are U2OS FANCD2-KO cells.\n• Figure legend 6f and text lines 393-396: please spell out better how the experiments was\nperformed or how existing datasets were used in the analysis.\nWe thank the reviewer for this comment. We have included more detailed information on the\nanalysis of publicly available genome-wide Repli-seq data in the revised manuscript.\n• As pointed out by Rev 1 and also reported ( https://doi.org/10.3892/ijo.2017.3862), long\ntreatments with high levels of Cordycepin can interfere with S-phase progression, so in principle\nsome of its effects in suppressing damage could be due because of a decreased in DNA rather\nthan RNA synthesis. It is unlikely that in the experimental conditions in which the experiments\nwere carried out (with only 4 hour exposure) this is a contributing factor but I believe useful to\ninclude information on EdU incorporation together with EU incorporation (currently Fig R5 in the\nrebuttal) after 4 hours of exposure to this drug.\nWe thank the reviewer for this helpful suggestion. As recommended, we have included the data\non EdU incorporation in cells treated with Cordycepin or overexpressing RNAseH1 and have\nclarified these results in the main text of the revised manuscript (New Supplementary Fig. 8f, g\nand page 13 of the main text).\nReviewer #3 (Remarks to the Author):\nWhile the authors have added much additional data to the manuscript, it does not change my\noverall assessment that the study is not suitable for publication and is majorly flawed. The\nexperiments presented by the authors are not set up to address the concerns I raised and in some\ncases further undermine the authors’ conclusions. I assessed the first six author responses to the\nconcerns I had raised and found these alone to be sufficient to recommend rejection of the\nmanuscript so I did not proceed further. Thus, my major concerns include but are not limited to:\n#1 XPF-ERCC1 depletion is required for repair of DNA interstrand crosslinks as well as in other\ncontexts. While it does make breaks the expected effect is removal of damage. The assertion that\nits depletion should reduce DNA damage is incorrect without also blocking the pathways that\noperate downstream of it (i.e. on the products of XPF-ERCC1 activity). Fig 2D is very complex\nand relies on the authors’ assertion that there was no Geminin expression upon entry into S-\nphase, which simply isn’t true (Fig. S4B shows it is dramatically reduced, not absent). Fig 2D\ndoesn’t analyze FANCD2 and there is no quantification for Fig. S4B so it’s unclear how this\ncorroborates Fig. 2D. The test the authors need to do is to assess whether the other Fanconi\nproteins they identify as behaving similarly to FANCD2 (RAD51, BRCA1, BRCA2 in Fig. 1B)\nbehave the same as FANCD2 or differently in other assays.\nWe believe there may have been a misunderstanding regarding the screening data, and we\napologize if we did not describe the results of the screen and the selection criteria with sufficient\nclarity.\nIn the previous version of the manuscript, candidate genes that increased γH2AX intensities or\nreduced cell counts were shown in Fig 1b for Geminin-depleted cells, without presenting z-scores\nin Geminin-depleted and control cells, for direct comparison. This likely led to the\nmisunderstanding that the genes shown specifically affected Geminin-depleted cells, which was\nnot the case. As shown in Supplementary Table 2, the depletion of BRCA1/2 or RAD51 did not\nsignificantly affect γH2AX nuclear intensities nor cell numbers in Geminin-depleted cells more\nthan it did in control cells. These genes were thus not identified as hits. In contrast, FANCD2\ndepletion showed a vulnerability that was specific to Geminin loss across our γH2AX analyses\n(New Fig. 1b and Supplementary Table 2), which is the reason we identified it as a hit and\nproceeded to further analyses. Though we had tried to explain in the text how FANCD2 was\nselected as a hit, we acknowledge that Fig. 1b needed to be interpreted in conjunction with\nSupplementary Table 2, which may have been confusing. We have now re-written this section of\nthe results (New Fig. 1b and page 5 of main text) and Materials and methods (page 23) while we\nhave provided a complete list of candidate genes and values in the Supplementary Table 2, to\nfully describe the results obtained from the screen and the criteria used to select hits and clarify\nany ambiguities. We hope that the reviewer will now agree with us that FANCD2 stands out as a\nhit in our screen and differs from other FA proteins.\nWe would also like to point out that during our screening we tested ERCC1 depletion using three\nseparate siRNA oligos (Supplementary Table 1). Our results show that ERCC1 depletion does\nnot increase DNA damage in Geminin-depleted cells (Supplementary Table 2). We have now\nperformed additional analyses in cells co-depleted of Geminin and ERCC1 and show that ERCC1\nloss causes a slight reduction in γH2AX nuclear intensities (Fig. R2). This contrasts to FANCD2,\nb\nwhich causes a significant increase, as presented in our manuscript.\n5.5x103\n****\n5x103\n4.5x103\n4x103\n3.5x103\nsiGMNN - +\nsiERCC1\nFig. R2. Quantification of siRNA-transfected hTERT-RPE1 cells immunostained against γH2AX.\nPlot shows a representative replicate out of two independent biological replicates (n=2). **** p <\n0.0001 (two-tailed t-test).\nWe hope that the reviewer will agree that these analyses strongly indicate that the role of FANCD2\nwe describe is distinct from its XPF-ERCC1-related function.\nIn addition, we employed several orthogonal approaches which demonstrate a function distinct\nfrom the XPF-ERCC1 pathway:\n- FANCD2 is recruited at re-replicating forks (SIRF and PLA with MCM and PCNA)\n- FANCD2 accumulates at origins located at active promoters/early fragile sites in re-\nreplicating cells (ChIP-seq data)\n)UA(\nseitisnetni\nraelcun\nXA2Hγ\n- Unprotected fork progression is observed in re-replicating cells depleted of FANCD2\n(Fiber assays)\n- Accummulation of ssDNA gaps is evident in re-replicating cells depleted of FANCD2 (S1\nnuclease fiber assay, native BrdU, RPA32 accumulation)\nWe hope the reviewer will agree that, taken together, our findings effectively rule out the possibility\nthat the role of FANCD2 during re-replication is merely through XPF-ERCC1 repair functions.\nInstead, our data strongly support its key role in the direct stabilization of re-replicating forks.\nRegarding the additional comment of the reviewer on Fig. 2d, we agree that stating \"no Geminin\nexpression\" when using an siRNA is an overstatement and apologize if the previous figure was\noverly complex. In the revised manuscript, we corrected the text to explicitly state that Geminin is\ndepleted (not absent) (page 7). Moreover, we have reorganized the synchronization experiment\nto enhance clarity and improve readability. We show the quantification of the different stages of\nS-phase based on EdU incorporation patterns in synchronized cells (Fig. 2d) and have now added\nthe quantification of FANCD2 foci over the same 30 hs time-course (Fig. 2e). Additionally, to\nprovide a more robust assessment of replication stress, we have included a new result showing\nthe quantification of RPA32 foci (a key marker of ssDNA) along the 30 hs time-course (Fig. 2e).\nOur analysis shows that, FANCD2 and RPA32 foci start accumulating early in Geminin-depleted\ncells, in parallel to the appearance of re-replicating cells. (New Fig. 2d, e and Supplementary Fig.\n5c, d). Importantly, Geminin-depleted cells continue to exhibit a high number of FANCD2/RPA32\nfoci at later time points while remaining EdU-positive and becoming pH3-positive, indicating that\nthese cells are arrested at the G2/M checkpoint and continue synthesizing DNA (Fig. 2e and\nSupplementary Fig. 5d). We thus demonstrate that re-replication upon Geminin loss occurs during\nthe first round of replication and leads to early FANCD2 and RPA32 recruitment.\n#2 There is synergy between Geminin knockdown and FANCD2 knockdown in terms of 53BP1\nfoci but, as noted by the authors, there’s no re-replication following Geminin knockdown in this\ncell type (>G2 population is less than the control for Fig. S1E). This data argues that the\nrelationship between FANCD2 and Geminin is independent of rereplication.\nWe are happy to clarify the phenotype of Geminin loss in hTERT-RPE1 cells, which forms the\nbasis of our screen. Our working hypothesis is that the non-cancerous hTERT-RPE1 cells, when\ndepleted of Geminin, do not overtly re-replicate (and thus we do not detect a high prevalence of\ncells with >4C DNA content), but they are primed for re-replication and thus rely on additional\nfactors for maintaining genome stability and viability2. As presented in the Results and Discussion\nsections (pages 5 and 17), the exact reason we selected this cell line for our screening was to\nfind candidates that selectively increase DNA damage and reduce cell survival in cells with low\nlevels of re-replication, which typically survive Geminin ablation. We must stress that we never\nargued that the functional relationship between Geminin and FANCD2 is related to FANCD2\naffecting levels of re-replication. On the contrary, we explicitly demonstrate that FANCD2 does\nnot protect re-replicating cells by preventing further re-replication (Fig. 3c, d and page 8 of the\nmain text). Instead, our data show that the increased DNA breaks we observe are due to re-\nreplicating forks—present because of Geminin depletion—breaking more frequently in the\nabsence of FANCD2, as a result of unprotected forks progressing faster, accumulating ssDNA\ngaps, and colliding with the transcription machinery. These points have been presented in detail\nin the Discussion.\n#4 The interpretation that H2AX foci form prior to FANCD2 foci assumes that the FANCD2 and\nH2AX signals are directly comparable in a quantitative manner. This is probably not the case and\nI think differences in antibody properties could easily explain the result. For example,\nphosphospecific antibodies are generally less specific than other poly- or mono-clonal antibodies,\nso the result in Fig. 2C could just reflect reduced sensitivity of the H2AX antibody. The data in\nFig. 2C could be considered supportive of the authors’ hypothesis that breaks occur after\nFANCD2 recruitment, but it cannot be the only piece of evidence for the temporal ordering as\nthings currently stand.\nWe want to clarify that we do not suggest that γH2AX foci form prior to FANCD2 foci. On the\ncontrary, we show that FANCD2 is recruited early upon Geminin depletion, before high levels of\nre-replication and extensive DNA breakage. We have now added additional synchronization data,\nshowing that FANCD2 is recruited from early/middle S-phase (from 14 hours post mitosis),\nprior to overt re-replication and accumulation of cells in G2. We would like to stress that Fig. 2c is by\nno means the only piece of evidence pointing to a direct function of FANCD2 in protecting re-\nreplicating forks. We have provided many lines of evidence demonstrating that FANCD2\nrecruitment is a protective response required to stabilize re-replicating forks against unrestrained\nprogression and collapse (rather than being recruited post double-strand breaks). The data\nsupporting this key mechanistic finding were already included in the previous round of revisions\n(see also above):\n• We demonstrated that FANCD2 is recruited at re-replicating forks by\nimmunofluorescence in asynchronous U2OS cells (Fig. 2c) and by SIRF (FANCD2-\nEdU) and PLA (FANCD2-MCM2 and FANCD2-PCNA) assays (Fig. 3a, b, and\nSupplementary Fig. 5e).\n• the synchronization experiments in Geminin-depleted cells demonstrate that FANCD2\n(and in the current version RPA as an alternative marker for the presence of ssDNA)\nstart accumulating from 14-16 hours following mitosis, in S phase (Fig. 2d, e and\nSupplementary Fig. 5c, d).\n• We demonstrated by DNA fibers that FANCD2 limits fork speed in Geminin-depleted\ncells (Fig. 3e).\n• We demonstrated that unrestrained fork progression in double Geminin and FANCD2-\ndepleted cells leads to the accumulation of ssDNA gaps by immunofluorescence\nagainst RPA32, DNA fibers treated with S1 nuclease and BrdU under non-denaturing\nconditions (Fig. 4a-e).\n• We demonstrated by ChIP seq that FANCD2 recruitment upon Geminin loss occurs at\nthe vicinity of early origins, which are characterized by increased replication-\ntranscription conflicts and fragility, and not at common fragile sites (CFSs), as in\ncontrol cells (Fig 6).\nWe believe the new data in this second revision, together with the above-mentioned experiments,\nstrongly support our hypothesis that FANCD2 recruitment occurs early and precedes high levels\nof DNA breakage, and we hope that the reviewer will now agree with us.\n#5 S phase accumulation is not sufficient to say that re-replication is taking place and the\nremaining arguments in this response are only correlative and thus do not directly address the\npoint.\nWe would like to point out that we demonstrate re-replication not through S-phase accumulation\nbut through the appearance of cells with >4C DNA content which incorporate EdU (Fig. 3c and\nSupplementary Fig. 1a), as well as overt increase in nuclear size (Supplementary Fig. 5a). We\nhope that the reviewer will agree that there is no doubt that overt re-replication is taking place in\nU2OS cells depleted of Geminin or Geminin and FANCD2 double-depleted cells (Fig. 3c and\nSupplementary Fig. 1a).\nThe point that the reviewer initially raised was whether the phenotypes we observe are taking\nplace within one cell cycle or they require passage through mitosis. To address this point, we\nperformed a synchronization experiment (Fig. 2d, e, described on page 7 of the revised\nmanuscript). In this experiment, presented with additional data in the new revised manuscript, we\ndepleted Geminin before S-phase entry and subsequently monitored EdU incorporation, together\nwith FANCD2, RPA32 as a marker of ssDNA as well as phosphorylated histone H3 (pH3) as a\nmarker of G2/M phase. As explained in comment #1 to this reviewer, Geminin-depleted cells show\na time-dependent increase in the number of FANCD2 foci and RPA32 foci as they enter S-phase,\nwhich coincides with the accumulation of re-replicating cells (diffused EdU staining) (New Fig. 2d,\ne and Supplementary Fig. 5c, d). Importantly, these cells continue to exhibit a high number of\nFANCD2/RPA32 foci at later timepoints, while they also stained positive for EdU and pH3 (New\nFig. 2d, e and Supplementary Fig. 5c, d) and show increased nuclear size, showing they are\narrested at the G2/M transition and continue to re-replicate. The synchronization experiment\ndirectly supports our model in which re-replication starts already from the first round of DNA\nsynthesis, and DNA breaks do not arise from mitosis -as the reviewer was suggesting initially- but\nrather occur after cells arrest at G2/M boundary and undergo excessive re-replication.\n#6 The magnitude of difference between FANCD2 depleted and control cells is small that it is not\nreasonable to conclude that FANCD2 depletion reduces nuclear size. While it is true that picking\nsome number of foci per cell is a standard measure in the field that does not address the issue of\nnuclear size increasing the expected number of foci – just because a metric is standard, it does\nnot mean it is appropriate in all cases.\nThe point of our response was not whether FANCD2 co-depleted cells have much smaller nuclei\nbut that their nuclei are not larger. Surely, the reviewer must agree that they are not larger. Thus,\nthe observation that they have a higher number of foci cannot be attributed to an increased\nnuclear size (as suggested by the reviewer). In any case, we have changed all figures in the\nmanuscript to show intensities per pixel (i.e. corrected for nuclear size) as requested by the\nreviewer. We would like to remind the reviewer that, following their suggestion, we re-analyzed\nall our data to show raw mean intensities, a metric that is completely independent on the size of\nthe nucleus. As for foci number, we refer the reviewer to the data presented in Fig. R7 (Fig. R3\nbelow), where we show that findings are the same whether we normalize the number of foci to\nthe nuclear area or not. We believe that we have shown beyond all doubt that the phenotypes we\ndescribe cannot be accounted for by increased nuclear size.\n40\n30\n20\n10\nquantification of cells with >10 53BP1 foci, plot shows the mean ± SD of all biological replicates.\nFor quantification of 53BP1 foci per nucleus, plot shows data for a representative biological\nreplicate. Unless specified otherwise, all experiments were performed at least in three biological\nreplicates (n=3) and a minimum of 250 cells were counted per replicate for each condition. In all\ncases: ns, non-significant, *p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001 (two-tailed t-test).\nAdditional references\nPromotes DNA Damage and Cell Death in Cancer Cells. Front Pharmacol 13, 860682\n(2022). https://doi.org/10.3389/fphar.2022.860682\np53. J Cell Biol 165, 473-482 (2004). https://doi.org/10.1083/jcb.200403106\n)%(\nicof\n1PB35\n01\n>\nslleC\n***\n****\nsiRNACtrl M N N N C A N C A G FA N+FA\nN\nM\nG\nsuelcun/icof\n1PB35\n**** ****\n40 **** 0.25 ****\n0.20\n30\n0.15\n20\n0.10\n10\n0.05\n0 0.00\nsiRNACtr G l M N F N A N N C + A FA N CA siRNACtr G l M N F N A N N N C + A FA N CA\nN M\nM G\nG\naera/icof\n1PB35\nFANCD2 restrains fork progression and prevents fragility at early origins upon\nre-replication\nPoint-by-point response to the reviewers' comments:\nWe are grateful to the reviewers once more for their thorough evaluation of our\nmanuscript and for their constructive comments.\nReviewer #1 (Remarks to the Author):\nThe authors have addressed all my comments.\nWe thank the reviewer for assessing our revision and for recommending our\nstudy for publication in Nature Communications.\nReviewer #5 (Remarks to the Author):\nIn my view, the authors have satisfactorily addressed most of the issues raised\nby the three reviewers. The third reviewer essentially reiterates their overall\nnegative assessment of the manuscript, but this seems to rest mainly on minor\nor interpretative criticisms. Importantly, none of the specific points raised\nappears to identify any substantive flaw in the study.\nPoint #1: The data convincingly demonstrate that FANCD2 prevents replication\nfork collapse in Geminin-depleted cells independently of the XPF-ERCC1\npathway. There is no substantive rationale to support the reviewer’s speculation\nthat re-replication would cause increased interstrand crosslinks. Even if a partial\ndependence of FANCD2 function on XPF were correct, this would represent a\nsecondary consideration rather than a major limitation.\nPoint #2: The reviewer infers that the absence of increased re-replication upon\nFANCD2 depletion implies an independence of FANCD2’s role from re-\nreplication. However, the authors’ objective was to identify factors that\nsafeguard genomic stability in Geminin-depleted cells, not necessarily to define\nfactors acting alongside Geminin to prevent re-replication. Clarifying this\nconceptual distinction is important for properly framing the study’s design and\nconclusions.\nPoint #4: This point concerns a minor methodological issue that has been\nadequately addressed by the authors and does not affect the main findings.\nPoint #5: The reviewer correctly states that S-phase accumulation alone does\nnot prove re-replication. However, the authors did not rely solely on this\nparameter. They provide data showing cells with over 4C DNA content and EdU\nincorporation, which are widely recognized as indicators of re-replication. The\nexpertise of the research team in this area is well established and lends\ncredibility to their interpretation.\nPoint #6: This issue also represents a minor concern that has been\nappropriately resolved by the authors and does not alter the study’s\nconclusions.\nWe are pleased with the reviewers' positive assessment of our manuscript and\ntheir recommendation for publication."}]

Decision letter