HMCES corrupts replication fork stability during base excision repair in homologous recombination–deficient cells

Peña-Gómez et al., Sci. Adv. 11, eads3227 (2025) 26 March 2025 Science AdvAnceS | ReSeARch ARticle 1 of 17 MOLECULAR BIOLOGY HMCES corrupts replication fork stability during base excision repair in homologous recombination–deficient cells María José PeñaGómez1,2, Yaiza RodríguezMartín1,2, Marta del Rio Oliva1,2, Yodhara Wijesekara Hanthi3,4, Sara Berrada5,6, Raimundo Freire7,8,9, Jean Yves Masson5,6, José Carlos Reyes1, Vincenzo Costanzo3,4, Iván V. Rosado1,2* Apurinic/apyrimidinic (AP) sites and singlestrand breaks arising from base excision repair (BER) during the misincorporation of damaged nucleobases may hinder replication fork stability in homologous recombination–deficient (HRD) cells. At templated AP sites, crosslinks between the DNA and 5hydroxymethylcytosine binding, embryonic stem cell–specific (HMCES) regulate replication fork speed, avoiding cytotoxic doublestrand breaks. While the role of HMCES at the template DNA strand is well studied, its effects on nascent DNA are not. We provide evidence that HMCES–DNAprotein crosslinks (DPCs) are detrimental to the BERmediated removal of 5hydro xymethyl2′- deoxycytidine (5hmdC)–derived 5hydroxymethyl2′- deoxyuridine from replication forks. HRD cells have heightened HMCESDPCs, which increase further upon 5hmdC exposure, suggesting that HMCES binds both spontaneous and 5hmdCinduced AP sites. HMCES depletion substantially suppresses 5hmdCmediated replication fork defects, chromosomal aberrations, and cell death in HRD cells. This reveals that HMCESDPCs are a source of BERinitiated singlestranded DNA gaps, which indicates that endogenous DPCs contribute to genomic instability in HRD tumors. INTRODUCTION Chemical or physical modifications of DNA bases, in addition to uracil (and its derivative forms) misincorporation, are perhaps the major source of DNA lesions in cells. These harmful DNA lesions appear at an estimated frequency of 1×105 to 5×105 cell−1 day−1 (1). During the repair of those lesions, apurinic/apyrimidinic (AP) sites are formed by DNA glycosylase activity. To avoid their deleterious consequences due to their intrinsic chemical instability, AP sites are rapidly converted to singlestrand breaks (SSBs) by either the AP lyase or the AP endonuclease (APEX) activities of certain class of nucleases. Therefore, the repair of base alterations, AP sites, and SSBs is perhaps the most important process responding to oxidative and alkylating base damages (2,3). More intriguingly is how these damaged bases are repaired by base excision repair (BER) in the context of replication forks. This is of utmost relevance as AP sites and SSBs are potent replication fork–blocking lesions, which, if left unrepaired, hinder replication fork stability. Moreover, the persistence of singlestranded DNA (ssDNA) gaps behind replication forks is relevant in the clinic to eradicate homologous recombination–deficient (HRD) tumors, as they are efficiently targeted by poly(adenosine 5′- diphosphate–ribose) polymerase (PARP) inhibitors. For these reasons, understanding ssDNA gap formation caused by BER activity is of paramount interest in the context of HRD, as many factors involved in HR [breast cancer gene 1 (BRCA1), breast cancer gene 2 (BRCA2), radiation sensitive protein 51 (RAD51), fanconi anemia group D2 (FANCD2), etc.] are also crucial for replication fork maintenance. Misincorporation of damaged nucleotides and depurination of adducted bases are among the most common spontaneous base lesions in DNA (1,4–6), triggering canonical BER (7,8). During damaged base removal, AP sites generated by DNA glycosylases [e.g., single-strand selective mo nofunctional uracil DNA glycosylase (SMUG1) during 5hydroxymethyl2′- deoxyuridine (5hmdU) excision] are converted to SSBs by APEX1. Upon DNA end processing, DNA polymerase β or λ conducts gap filling, leaving nicked DNA ready for ligation by the xray repair crosscomplementing 1 (XRCC1)/DNA ligase 1 or ligase 3 complexes (9–11). Despite the “passingthebaton” model of BER, where DNA intermediates are passed along from one step to the next, AP sites or SSBs may persist and challenge replication fork progression, eliciting a replication stress (RS) response. Upon replication fork stalling by DNA intermediates, ataxia telangiectasia and Rad3related (ATR) accelerates recruitment and stabilization of critical components of the DNA damage response, such as the Fanconi anemia (FA) pathway and the HR factors FANCD2/FANCI-associated nuclease 1 (FAN1), structure-specific endonuclease (SLX4), xeroderma pigmentosum F (XPF), BRCA1/2, or RAD51. To fulfill DNA synthesis, the primasepolymerase (PRIMPOL) reprimes ahead of both the stalled replication fork and the lesion to limit replication fork instability, thus dampening RS (12–14). However, PRIMPOL activity generates potentially cytotoxic ssDNA gaps behind forks (15–18), which are subsequently filled in either by REV1POLζ translesion synthesis (TLS) polymerases or by POLQ (19– 21). Several mechanisms have been recently proposed as major contributors to HRD cell lethality. The unifying feature among them is the 1centro Andaluz de Biología Molecular y Medicina Regenerativa (cABiMeR) Universidad de SevillacSicUniversidad Pablo de Olavide, Seville 41092, Spain. 2departamento de Genética, Facultad de Biologia, Universidad de Sevilla, Seville 41012, Spain. 3iFOM, the AiRc institute of Molecular Oncology, Milan, italy. 4department of Oncology and hematologyOncology, University of Milan, Milan, italy. 5Genome Stability laboratory, chU de Québec Research center, hdQ Pavilion, Oncology division, 9 McMahon, Québec city, Qc G1R 3S3, canada. 6department of Molecular Biology, Medical Biochemistry and Pathology, laval University cancer Research center, Québec city, Qc G1v 0A6, canada. 7Unidad de investigación, hospital Universitario de canarias, instituto de investigación Sanitaria de canarias (iiSc), la laguna, Santa cruz de tenerife, Spain. 8instituto de tecnologías Biomédicas, centro de investigaciones Biomédicas de canarias, Facultad de Medicina, campus ciencias de la Salud, Universidad de la laguna, Santa cruz de tenerife, Spain. 9Universidad Fernando Pessoa canarias, las Palmas de Gran canaria, Spain. *corresponding author. email: ivrosado@ us. es copyright © 2025 the Authors, some rights reserved; exclusive licensee American Association for the Advancement of Science. no claim to original U.S. Government Works. distributed under a creative commons Attribution noncommercial license 4.0 (cc BYnc). Downloaded from https://www.science.org at Universidad de Sevilla on May 12, 2025 Peña-Gómez et al., Sci. Adv. 11, eads3227 (2025) 26 March 2025 Science AdvAnceS | ReSeARch ARticle 2 of 17 presence of persistent ssDNA gaps at nascent DNA strands, which correlates with the exquisite cytotoxicity to PARP inhibitors (e.g., olaparib) observed in HRD cells (22,23). However, the underlying molecular basis of this cytotoxicity still remains elusive. Recently, misincorporation of dU, 5′- chlorodU (5CldU), or 5hmdU has emerged as a previously unidentified source of replication fork impairment (24–27). Upon misincorporation, persisting DNA repair intermediate structures can be collided by the replisome, resulting in BERreplication conflicts (BRCs). These unresolved BRCs account for HRD cell death in the absence of proficient replication fork maintenance or HR pathways (24–27). In addition, modified DNA bases remaining undetectable throughout the cell cycle induce an ATRdependent RS and fork collapse response during the next cell cycle when present in template DNA (28,29). Whereas ssDNA gap accumulation by BRCs during the next cell cycle is well documented (12,30–33), how ssDNA gaps arise at nascent DNA during the current cell cycle remains largely unexplored (25,26). Damaged DNA bases are one of the main sources of AP sites. On template DNA, AP sites are exposed during DNA unwinding by Cdc45– MCM–GINS (CMG) helicase and are protected by the 5-hydroxymethylcytosine binding, embryonic stem cell–specific (HMCES) protein (34–39). Through its interaction with proliferating cell nuclear antigen (PCNA), HMCES associates to replication forks and binds ssDNA to shield AP sites (34,35). Through a thiazolidine bond between the Cys2 residue and the aldehydic conformation of the AP site (34–36,40), HMCES attaches covalently to AP sites, forming an HMCES–DNA-protein cross-links (DPC). HMCES-DPC formation on template ssDNA decreases replication fork speed while avoiding APEX1mediated cytotoxic doublestrand breaks (DSBs) at replication forks (35). HMCESDPCs are bypassed by a TLS step and subsequently removed from DNA by distinct mechanisms. Upon winding of DNA strands, HMCESDPC catalyzes a crosslink selfreversal reaction through its Glu127 residue, releasing HMCES from DNA (37,38). Alternatively, assisted by the protein denaturation activity of fanconi anemia group J (FANCJ) (39), SprT-like Nterminal domain (SPTRN) or proteasome degrades HMCES to allow replication fork resumption (35,39). While the consequences of HMCESDPCs on template DNA are recently being addressed, the effects on nascent DNA are unknown. Here, we provide evidence of a detrimental role of HMCESDPCs during nascent DNA synthesis. HRD cells display heightened chromatin levels of HMCES, which increase upon 5-hydroxymethyl-2′- deoxycytidine (5hmdC) exposure, suggesting that HMCES binds to spontaneous and 5hmdUinduced AP sites. HMCES loss rescues replication fork impairment, genomic instability, and cytotoxic phenotypes observed in HRD cells during 5hmdU misincorporation, indicating that HMCES is responsible for the 5hmdUmediated genetic instability and lethality of HRD cells. Genetic depletion of ssDNA gap–generating BER factors (e.g., SMUG1 or APEX1/2) largely suppresses these phenotypes, whereas loss of ssDNA gap–filling factors PARP1 or XRCC1 exacerbates them. Our findings demonstrate that HMCESDPCs on nascent DNA are responsible for the replication fork defects observed in HRDdeficient cells and place endogenous DPCs on nascent DNA as a novel source of genomic instability in HRD tumors. RESULTS Misincorporation of 5hmdU arising from deaminated 5hmdC causes genomic instability in HRD cells 5hmdU can arise from either the 5hmdC salvage pathway or the cytosine demethylation process (24,25,27). Previous data from our group and others have shown that HRD cells lacking FANCD2, methyl methanesulfonate and UV sensitive protein 81 (MUS81), BRCA1, or BRCA2 were sensitive to misincorporation of 5hmdU on genomic DNA (25,27). To investigate the molecular mechanisms accounting for 5hmdUmediated genotoxicity, we examined whether depletion of factors involved in the pyrimidine salvage pathway [e.g., deoxycytidine kinase (DCK) or 2′- deoxycytidine 5′- monophosphate deaminase (DCTD)] had any effect on the viability of 5hmdCtreated Fancd2−/− mouse embryonic fibroblasts (MEFs). Consistent with previous results in human HRD cells (25,27), 5hmdCmediated Fancd2−/− lethality was largely suppressed by knockdown of DCK or DCTD, whereas wildtype cells remained unaltered (Fig. 1A and fig. S1), confirming that deamination of 5hmdC to 5hmdU mediates HRD cell cytotoxicity. As PARP trapping by olaparib exacerbated 5hmdCdependent Fancd2−/− chromosomal instability and cell lethality (25), we sought to determine levels of nuclear PARylation and SSBs upon 5hmdC treatment. As expected, Fancd2−/− ethynyl2′- deoxyuridine–positive (EdU+) cells displayed a significant increase in nuclear PARylation (Fig. 1B), also confirmed in EdU unrestricted cells (fig. S2A), while it remained relatively unchanged in wildtype cells (Fig. 1B and fig. S2A). Next, we examined PAR levels associated with ongoing replication forks by “in situ protein interaction with nascent DNA replication forks” also known as SIRF assay. Compared to wildtype MEFs, Fancd2−/− cells displayed heightened EdUPAR foci, which were further exacerbated by 5hmdC treatment (Fig. 1C). The increase in nuclear PARylation or EdUPAR foci observed in Fancd2−/− cells was largely dependent on PARP1, as PARP1 knockdown significantly reduced both PARylation and EdUPAR foci (Fig. 1, B and C). Fancd2−/− MEFs also displayed significant spontaneous SSBs measured by alkaline comet assay, which were further increased by 5hmdC treatment, unlike in wildtype cells (Fig. 1D). These data suggest that 5hmdU misincorporation results in ssDNA gap accumulation in the absence of FANCD2, probably accounting for heightened PARylation at replication forks. Consistent with this, we also found that 5hmdC exposure caused increased replication fork asymmetry by DNA fiber assay (Fig. 1E), indicating that 5hmdUdependent replication fork impairment was not a consequence of a global checkpoint response. Moreover, 5hmdUmediated DNA damage increased sister chromatid exchanges (SCEs) in Fancd2−/− cells (Fig. 1F), suggesting that BRCs elicited an HRdependent repair mechanism. However, we did not observe any evidence of checkpoint activation measured as Ser345CHEK1 or Ser33RPA2 under these conditions (fig. S3). These data suggest that short 5hmdC exposure does not activate the DNA damage checkpoint despite increased SSBs and indicate that ssDNA gaps arising during 5hmdU removal trigger replication fork impairment and sister chromatid recombination in HRD cells. ssDNA gaps arising from the concerted activities of SMUG1 and APEX1/2 on misincorporated 5hmdU alter replication fork progression Previous studies have reported ssDNA gap accumulation as a major contributor to BRCA1/2 tumor cell death (18,31,41). As SMUG1 is the main BER DNA glycosylase responsible for 5hmdU elimination from genomic DNA (42), we sought to examine its contribution to ssDNA gap formation in nascent DNA. In agreement with a previous report (27), SMUG1 depletion in 5hmdCtreated Fancd2−/− cells significantly decreased nuclear PARylation to untreated levels in both total and EdU+ cells compared to wildtype cells (Fig. 2A and fig. S2B). SMUG1 depletion suppressed 5hmdCdependent EdUPAR Downloaded from https://www.science.org at Universidad de Sevilla on May 12, 2025 Peña-Gómez et al., Sci. Adv. 11, eads3227 (2025) 26 March 2025 Science AdvAnceS | ReSeARch ARticle 3 of 17 Fig. 1. Genomic instability induced by 5hmdC in HRD cells. (A) (3-(4, 5-dimethylthiazolyl-2)-2, 5-diphenyltetrazolium bromide) assay (Mtt) cell proliferation assay of 5hmdctreated wildtype or Fancd2−/− (Fd2−/−) cells knocked down for dctd (top) or dcK (bottom) exposed to the indicated dose for 3 days. (B) left: Representative PAR (red) immunofluorescence images of edU+ (green) wildtype, siPARP1, Fancd2−/−, and Fancd2−/− siPARP1 cells exposed to 5hmdc (10 μM) for 3 hours. 4′,6-diamidino2phenylindole (dAPi; blue) stains nuclear dnA. Right: Plot depicting PAR mean intensity signal per nucleus. A.U., arbitrary units. (C) left: Representative images of edU+- PAR foci (red) by SiRF analysis from wildtype, siPARP1, Fancd2−/−, and Fancd2−/− siPARP1 cells exposed to 5hmdc (10 μM) for 3hours. dAPi (blue) stains nuclear dnA and edU (green) stains Sphase cells. Right: Plot depicting edUPAR foci per nucleus (n=2). (D) left: Representative images of alkaline comet assay from wildtype and Fancd2−/− cells following 5hmdc treatment (10 μM) for 3hours. Right: Plot depicting comet tail moment per cell (n=4). (E) top left: Scheme of the dnA fiber origin symmetry. Bottom left: Representative images of dnA fibers from wildtype and Fancd2−/− cells upon 5hmdc treatment (40 μM) for 30 min. Right: Box plot of the asymmetry index. (F) top left: Scheme of Sce assay. Bottom left: Representative images of Sces from wildtype and Fancd2−/− cells exposed to 5hmdc (10 μM) for 40 hours. Right: Bar plot of Sce/chromosome per metaphase spread (n=50 of each of the two biological replicates; bar represents means±Sd). BrdU, 5bromo2′- deoxyuridine. Downloaded from https://www.science.org at Universidad de Sevilla on May 12, 2025 Peña-Gómez et al., Sci. Adv. 11, eads3227 (2025) 26 March 2025 Science AdvAnceS | ReSeARch ARticle 4 of 17 Fig. 2. 5hmdCmediated HRD genomic instability is dependent on SMUG1. (A) left: Representative PAR (red) immunofluorescence images of edU+ (green) wildtype, siSMUG1, Fancd2−/−, and Fancd2−/− siSMUG1 cells exposed to 5hmdc (10 μM) for 3hours. dAPi (blue) stains nuclear dnA. Right: Plot depicting PAR mean intensity signal per nucleus. (B) left: Representative images of edUPAR foci (red) by SiRF assay from wildtype, siSMUG1, Fancd2−/−, and Fancd2−/− siSMUG1 cells exposed to 5hmdc (10 μM) for 3hours. dAPi (blue) stains nuclear dnA and edU (green) stains Sphase cells. Right: Plot depicting edUPAR foci per nucleus (n=2). (C) left: Representative images of alkaline comet assay from wildtype, siSMUG1, Fancd2−/−, and Fancd2−/− siSMUG1 cells following 5hmdc treatment (10 μM) for 3hours. Right: Plot depicting comet tail moment per cell (n=2). (D) top left: Scheme of dnA fiber assay. Bottom left: Representative images of dnA fibers from Fancd2−/− and Fancd2−/− siSMUG1 cells untreated (Unt) or exposed to 5hmdc (40 μM) for 30 min. Right: Box plot representing the frequency of 5′- iododeoxyuridine (idU)/cldU ratio of wildtype, siSMUG1, Fancd2−/−, and Fancd2−/− siSMUG1 cells after 5hmdc treatment (n=200). (E) Mtt cell proliferation assay of wildtype, siSMUG1, Fancd2−/−, and Fancd2−/− siSMUG1 cells exposed to the indicated dose of 5hmdc for 3 days (n=4; means±Sd). Downloaded from https://www.science.org at Universidad de Sevilla on May 12, 2025 Peña-Gómez et al., Sci. Adv. 11, eads3227 (2025) 26 March 2025 Science AdvAnceS | ReSeARch ARticle 5 of 17 foci to untreated levels (Fig. 2B), suggesting that most replication fork–associated PARylation upon 5hmdC exposure depends on SMUG1. Consistent with this, SMUG1 knockdown also abolished 5hmdCinduced SSBs in Fancd2−/− cells (Fig. 2C), suggesting that SMUG1 is responsible for 5hmdCinduced ssDNA gap formation. Moreover, SMUG1 depletion largely restored replication fork progression measured by DNA fiber assay and cell viability in 5hmdCtreated Fancd2−/− cells (Fig. 2, D and E). These data demonstrate that SMUG1 loss suppresses ssDNA gap accumulation, replication fork impairment, and lethality observed in 5hmdCtreated HRD cells, most likely throughout the abolishment of AP site formation. Mammalian cells contain two class II APEXs, APEX1 and APEX2 (43,44). While APEX1 has strong APEX activity, APEX2 exhibits strong in vitro 3′- phosphodiesterase and 3′- 5′ exonuclease activities (44). We therefore examined the contribution of APEX1 or APEX2 to PARylation and ssDNA gap formation during 5hmdU elimination. Similar to SMUG1 loss, APEX1 knockdown had hardly any effect on nuclear PARylation in untreated cells (Fig. 3A). However, 5hmdCdependent nuclear PARylation in total or EdU+ Fancd2−/− cells significantly decreased to near–wildtype levels (Fig. 3A and fig. S2C). APEX1 loss significantly suppressed spontaneous and 5hmdCinduced EdUPAR foci in Fancd2−/− cells to wildtype levels (Fig. 3B). Moreover, loss of APEX1 significantly reduced 5hmdCinduced SSBs observed in Fancd2−/− cells to almost untreated levels (Fig. 3C). These data suggest that APEX1 is the main endonuclease responsible for most 5hmdCinduced SSBs in Fancd2−/− and, to a lesser extent, in wildtype cells. APEX1 depletion also substantially suppressed 5hmdCmediated Fancd2−/− replication fork impairment and cell lethality (Fig. 3, D and E), indicating that APEX1dependent SSBs account for the replication fork defects and lethality observed in 5hmdCtreated Fancd2−/− cells. As APEX1 knockdown does not fully restore cell viability or fork dynamics, these results also suggest that APEX1 may play additional roles during stability or resumption of replication forks. Similar to APEX1 depletion, APEX2 knockdown also reduced 5hmdCmediated nuclear PARylation in total or Sphase (EdU+) Fancd2−/− cells populations (fig. S4, A and B), to a lesser extent, than APEX1 depletion. APEX1 and APEX2 codepleted Sphase Fancd2−/− cells showed 5hmdCinduced PAR levels similar to those observed in APEX1depleted 5hmdCtreated Fancd2−/− cells, indicating an epistatic relationship between APEX1 and APEX2. Moreover, APEX1 or APEX2 knockdown completely suppressed 5hmdCinduced EdUPAR foci and SSBs (Fig. 3F and fig. S4C). These data indicate that APEX1, in combination with APEX2, is responsible for the increased PARylation and SSBs associated with nascent strand during 5hmdU removal. Consistent with this notion, APEX1 and APEX2 showed an epistatic relationship on PAR levels and lethality observed in 5hmdCtreated Fancd2−/− cells (Fig. 3E). These data point out to a concerted function of APEX1 and APEX2 at processing 5hmdUderived AP sites, likely generating ssDNA gaps. 5hmdUderived ssDNA gap persistence at replication forks exacerbates genomic instability and lethality in HRD cells In unperturbed Sphase, ssDNA gap persistence due to inefficient Okazaki fragment ligation is signaled by PARP1dependent PARylation, followed by XRCC1mediated PARP1 eviction, to promote faithful gap repair (45). We reasoned that defective 5hmdCdependent ssDNA gap processing either by excessive PARP1 retention or by PARP1 or XRCC1 loss would exacerbate ssDNA gap persistence and cell lethality, thus mimicking persistent unligated Okazaki fragments. XRCC1 knockdown increased further nuclear PARylation in 5hmdCtreated total or EdU+ Fancd2−/− cell populations while remaining unaffected in wildtype cells (Fig. 4A and fig. S2D). Likewise, the SIRF assay revealed a significant increase in EdUPAR foci in 5hmdCtreated Fancd2−/− cells upon XRCC1 depletion (Fig. 4B). PARP1 or XRCC1 depletion further exacerbated 5hmdCmediated Fancd2−/− cell lethality (Fig. 4C) similar to PARP1 trapping by olaparib (25) as a result of inefficient repair of ssDNA gaps. However, XRCC1 depletion did not further affect 5hmdCmediated Fancd2−/− replication fork impairment (Fig. 4D). These data indicate that ssDNA gaps associated with replication forks in the absence of FANCD2 persist or accumulate to a larger extent upon XRCC1 loss. Replication forks stalled by bulky base adducts stimulate postreplicative SCEs in an ssDNA gap– and PRIMPOLdependent fashion (14). We therefore examined SCEs induced by 5hmdCmediated ssDNA gaps in nascent DNA by exposing cells to 5hmdC for less than one cell cycle length (≈12 hours). We found that 5hmdC triggered a significant increase in SCEs in Fancd2−/− cells compared to untreated cells (Fig. 4E). Moreover, SMUG1 or APEX1 depletion suppressed 5hmdCinduced SCEs, while PARP1 or XRCC1 knockdown exacerbated them (Fig. 4E). These data indicate that 5hmdCmediated ssDNA gaps in nascent strands occurring during the current cell cycle are a source of SCEs in HRD cells. Our findings suggest that defective repair of persistent replicative 5hmdCinduced ssDNA gaps accounts for increased PARylation, replication fork impairment, heightened SCEs, and lethality observed in Fancd2−/− cells. HMCESDPCs are responsible for 5hmdCmediated ssDNA gaps in HRD cells AP sites and other intermediate DNA ends during BER hamper replication fork progression, causing BRCs (28,29,36,46). AP sites are protected by HMCES to avoid cytotoxic DSB formation at the replication fork while promoting TLS bypass mechanisms (35,36,40). We therefore reasoned that HMCES could react to 5hmdUderived AP sites arising during 5hmdC treatment. Upon preextraction, we observed a significantly higher proportion of nuclei stained positive for chromatinbound Flagtagged HMCES in Fancd2−/− than in wildtype cells under untreated conditions (Fig. 5A). Fancd2−/− cells also displayed a brighter chromatinbound HMCES signal compared to its wildtype counterpart under spontaneous, methyl methanesulfonate (MMS) used as a positive control, or 5hmdC exposure (Fig. 5B). Moreover, chromatinbound HMCES levels directly correlated with nascent strand 5hmdC misincorporation in a dosedependent manner, reaching significantly higher levels in Fancd2−/− than in wildtype cells (Fig. 5B). We also confirmed these findings by cellular subfractionation and by rapid approach to DNA adduct recovery (RADAR) assays, showing a 5hmdC dosedependent HMCES chromatin retention (fig. S5, A and B). These data suggest that Fancd2−/− cells present higher levels of endogenous AP site–bound HMCES than wildtype cells, which increase upon 5hmdC treatment in a dosedependent manner. We next examined the chromatin retention activity of cell lines overexpressing either Cys2→Ala (C2A) or Arg212→Glu (R212E), two mutant versions of HMCES showing defective DPC formation and ssDNA binding activities, respectively. Consistent with their reported in vitro biochemical activities (35), HMCESC2A showed impaired chromatin recruitment upon 5hmdC or MMS exposure, while HMCESR212E completely Downloaded from https://www.science.org at Universidad de Sevilla on May 12, 2025 Peña-Gómez et al., Sci. Adv. 11, eads3227 (2025) 26 March 2025 Science AdvAnceS | ReSeARch ARticle 6 of 17 Fig. 3. APEX1 and APEX2 contribute to 5hmdCinduced ssDNA gap generation in HRD cells. (A) left: Representative PAR (red) immunofluorescence images of siA1depleted (siA1) wildtype or Fancd2−/− cells exposed to 5hmdc (10 μM) for 3hours. dAPi (blue) stains nuclear dnA and edU (green) stains Sphase cells. Right: Plot depicting PAR mean intensity signal per edU+ nucleus. (B) left: Representative images of edUPAR foci (red) by SiRF assay from wildtype, siA1 Fancd2−/−, or Fancd2−/− siA1 cells exposed to 5hmdc (10 μM) for 3hours. dAPi (blue) stains nuclear dnA and edU (green) stains Sphase cells. Right: Plot depicting edUPAR foci per nucleus. (C) left: Representative images of alkaline comet assay of APeX1 knockdown wildtype or Fancd2−/− cells exposed to 5hmdc (10 μM) for 3hours. Right: Plot depicting comet tail moment per cell. (D) top left: Scheme of dnA fiber assay. Bottom left: Representative images of dnA fibers from Fancd2−/− and Fancd2−/− siA1 cells untreated or exposed to 5hmdc (40 μM) for 30 min. Right: Box plot of the frequency of idU/cldU ratio of siA1depleted wildtype or Fancd2−/− cells upon 5hmdc treatment (n=200 of each of the two biological replicates). (E) top: Mtt cell proliferation assay of siA1depleted wildtype or Fancd2−/− cells exposed to the indicated dose of 5hmdc for 3 days (n=9; means±Sd). Bottom: Mtt cell proliferation assay of APeX1or siA2depleted wildtype or Fancd2−/− cells exposed to the indicated dose of 5hmdc for 3 days. (F) Plot depicting edUPAR foci per nucleus by SiRF assay from wildtype, Fancd2−/−, siA1, siA2, Fancd2−/− siA1, and Fancd2−/− siA2 cells exposed to 5hmdc (10 μM) for 3hours (n=2). Downloaded from https://www.science.org at Universidad de Sevilla on May 12, 2025 Peña-Gómez et al., Sci. Adv. 11, eads3227 (2025) 26 March 2025 Science AdvAnceS | ReSeARch ARticle 7 of 17 Fig. 4. XRCC1 depletion exacerbates 5hmdCinduced genomic instability and lethality of HRD cells. (A) left: Representative PAR (red) immunofluorescence images of XRcc1depleted wildtype or Fancd2−/− cells exposed to 5hmdc (10 μM) for 3hours. dAPi (blue) stains nuclear dnA and edU (green) stains Sphase cells. Right: Plot depicting PAR mean intensity signal per nucleus. (B) left: Representative images of edUPAR foci (red) by SiRF assay of XRcc1depleted wildtype or Fancd2−/− cells exposed to 5hmdc (10 μM) for 3hours. dAPi (blue) stains nuclear dnA and edU (green) stains Sphase cells. Right: Plot depicting edUPAR foci per nucleus. (C) top: Mtt cell proliferation assay of wildtype or Fancd2−/− cells lacking XRcc1 exposed to the indicated dose of 5hmdc for 3 days (n=9; means±Sd). Bottom: Mtt cell proliferation assay of wildtype, siPARP1, Fancd2−/−, and Fancd2−/− siPARP1 cells exposed to the indicated dose of 5hmdc for 3 days (n=5; means±Sd). (D) top left: Scheme of dnA fiber assay. Bottom left: Representative images of dnA fibers from Fancd2−/− and Fancd2−/− siXRcc1 cells untreated or exposed to 5hmdc (40 μM) for 30 min. Right: Box plot of the frequency of idU/cldU ratio of XRcc1depleted wildtype or Fancd2−/− cells upon 5hmdc treatment (n=200 of each of the two biological replicates). (E) top right: Scheme of the Sce assay. left: Bar plot of Sce/chromosome per metaphase from SMUG1- , APeX1- , XRcc1- , or PARP1depleted Fancd2−/− cells exposed to 5hmdc (10 μM) for 12hours (n=50 of each of the two biological replicates; bar represents means±Sd). Downloaded from https://www.science.org at Universidad de Sevilla on May 12, 2025 Peña-Gómez et al., Sci. Adv. 11, eads3227 (2025) 26 March 2025 Science AdvAnceS | ReSeARch ARticle 8 of 17 Fig. 5. HMCES accumulates in nuclei and is responsible for 5hmdCmediated genomic instability observed in HRD cells. (A) left: Representative immunofluorescence images of Flagtagged hMceS (green) positive nuclei from wildtype or Fancd2−/− cells. Right: Percentage of hMceSFlAG+ wildtype and Fancd2−/− cells. (B) left: Representative immunofluorescence images of hMceS positive nuclei from wildtype or Fancd2−/− cells upon combined 5hmdc (160 μM) and Ncarbobenzyloxylleucyllleucyllleucinal (MG132; 10 μM) treatments for 3 hours. Right: hMceSFlAG mean intensity signal per nucleus from wildtype and Fancd2−/− cells upon 5hmdc (3 hours) or MMS (0.5 mM, 30 min). treatments were combined with MG132 (10 μM). (C) left: images of FlAGtagged hMceS positive nuclei from Fancd2−/− cells, expressing MmhMceSWtFlAG, MmhMceSc2AFlAG, or MmhMceSR212eFlAG upon 5hmdc exposure (160 μM, 3hours). Right: Plot depicting hMceSFlAG mean intensity signal per nucleus from wildtype or Fancd2−/− cells expressing MmhMceSWtFlAG, MmhMceSc2AFlAG, or MmhMceSR212eFlAG upon 5hmdc (indicated dose for 3hours) or MMS (0.5 mM). treatments were combined with MG132 (10 μM). ns, not significant. (D) left: Representative images of edUPAR foci by SiRF assay from hMceSdepleted wildtype or Fancd2−/− cells exposed to 5hmdc (10 μM) for 3hours. dAPi (blue) stains nuclear dnA and edU (green) stains Sphase cells. Right: Plot depicting edUPAR foci per nucleus. (E) left: Representative images of alkaline comet assay from hMceSdepleted wildtype or Fancd2−/− cells following 5hmdc treatment (10 μM) for 3hours. Right: Plot depicting comet tail moment per cell. Downloaded from https://www.science.org at Universidad de Sevilla on May 12, 2025 Peña-Gómez et al., Sci. Adv. 11, eads3227 (2025) 26 March 2025 Science AdvAnceS | ReSeARch ARticle 9 of 17 abolished it (Fig. 5C and fig. S6). These data suggest that 5hmdUderived AP sites on nascent DNA strand are recognized by HMCES, forming HMCESDPCs. We predicted that HMCES loss would increase cytotoxic ssDNA gaps on nascent DNA strand by APEX1mediated incision of AP sites right at or behind the fork, resulting in a synthetic lethal phenotype in HRD cells. To our surprise, HMCES knockdown markedly decreased 5hmdCinduced nuclear PARylation in total or EdU+ Fancd2−/− cell populations (fig. S7, A and B), EdUPAR foci in Sphase cells (Fig. 5D), and SSBs observed in Fancd2−/− cells (Fig. 5E), suggesting that HMCESDPCs on nascent DNA strand are responsible for a large subset of 5hmdCinduced ssDNA gaps observed in Fancd2−/− cells. Consistent with the protective role of HMCES during replication (35), HMCES depletion increased nuclear γH2AX, which was further elevated in the absence of FANCD2 (fig. S7C). Nevertheless, γH2AX levels remained relatively unchanged during 3hours of 5hmdC treatment, suggesting that ssDNA gaps in nascent DNA arising from AP sites unprotection due to HMCES loss do not result in cumulative DSBs (fig. S7C). HMCES depletion also restored Fancd2−/− replication fork progression and symmetry to wildtype levels (Fig. 6, A and B). 5hmdCinduced heightened chromosomal aberrations were markedly reduced upon HMCES depletion (Fig. 6C), specifically those of radial chromosomes, which arose at the expense of chromosomal gaps (fig. S8). As a consequence, 5hmdCinduced HRD cell viability was also notably improved upon HMCES depletion (Fig. 6D), indicating that HMCESDPCs formed at BER intermediates are responsible for replication fork impairment, chromosomal instability, and cell lethality in HRD cells. To rule out that the phenotypic suppression by HMCES knockdown was a consequence of pooled small interfering RNA (siRNA) offtargets, we validated our results using four independent siRNAs against HMCES (from number 1 to number 4) compared to pooled siRNAs, obtaining similar results to pooled siRNAs (fig. S9). We also validated our results using human BRCA2–deficient DLD1 cells, which were previously reported to be sensitive to 5hmdU (25). HMCES knockdown also suppressed 5hmdCmediated BRCA2−/− cell lethality (Fig. 6E), thus broadening these findings to different human genetic backgrounds. To directly examine the effects of HMCES depletion on DNA replication intermediates, we use DNA electron microscopy (EM) to monitor the frequency of broken replication fork intermediates, possibly resulting from ssDNA gap breakage (47). In agreement with our genetic data, compared to untreated condition, 5hmdCtreated Fancd2−/− cells displayed significant increase in DNA replication intermediates with asymmetric branches, indicating broken forks, possibly resulting from the processing of ssDNA gaps present in Fancd2−/− cells (Fig. 6F and fig. S10) (47). HMCES depletion largely suppressed 5hmdCmediated fork instability and ssDNA gap accumulation (Fig. 6F). We also attempted to validate these findings by S1 nuclease DNA fiber assay but failed to obtain reproducible results in MEFs. To overcome this, we knocked out FANCD2, HMCES, or both genes in human RPE1 cells by CRISPRCas9 approaches (fig. S11, A and B). Consistent with published data, HMCES loss did not affect unperturbed replication fork progression (40) or the amount of ssDNA gaps, suggesting that unprotection of spontaneous AP sites upon HMCES loss does not affect overall levels of ssDNA gaps (fig. S11C). Despite S1 insensitivity in untreated FANCD2−/− RPE1 cells, small ssDNA gaps were visible in all Fancd2−/− MEFs samples by EM (Fig. 6F and fig. S10), compatibly with AP site accumulation on template strands. Moreover, exposure of FANCD2−/− RPE1 cells to 5hmdC (10 μM) significantly affected 5′- iododeoxyuridine (IdU) track length, suggesting that FANCD2 is required to avoid ssDNA gap formation. Notably, HMCES depletion suppressed 5hmdCinduced ssDNA gap formation in FANCD2−/− cells detected by S1 nuclease (fig. S11C), confirming that HMCESDPCs are responsible for 5hmdUderived ssDNA gap accumulation in HRD cells. We also found that low expression levels of HMCES correlated with lower overall survival in two independent cohorts of patients with breast adenocarcinoma, one from TCGA (1089 patients) and another from a set of curated breast cancer cohorts from GEO (2976 patients) (Fig. 6G and fig. S11D). These data suggest that HMCES loss might play a role in breast cancer aggressiveness or patient outcome, although these findings need further validation. HMCESDPCs on template DNA strand showed an epistatic relationship over APEX1 incision to avoid toxic DSB formation (34,35). We therefore examined nuclear PARylation and viability phenotypes upon depletion of HMCES, APEX1, APEX2, or their combinations. Nuclear PARylation in HMCES APEX1 doubleknockdown EdU+ cells resembled the one seen in APEX1 knockdown Fancd2−/− cells (Fig. 7A). However, PARylation in HMCES APEX2 doubleknockdown Fancd2−/− cells resembled the one observed in Fancd2−/− HMCES singleknockdown EdU+ cells (Fig. 7B). Moreover, knockdown of HMCES, APEX1, or APEX2 also showed an epistatic suppressive phenotype on Fancd2−/− cell survival (Fig. 7, C and D). These data suggest that HMCES, APEX1, and APEX2 function in the same genetic pathway during the processing of 5hmdU from nascent DNA. PRIMPOL mediates a subset of 5hmdCmediated ssDNA gaps Replication fork stalling by bulky base adducts generates postreplicative ssDNA gaps throughout the primase and polymerase activities of PRIMPOL (14,19). We therefore examined the contribution of PRIMPOL to 5hmdCinduced ssDNA gap formation in our cell lines. PRIMPOL depletion had little effect on PARylation levels in untreated EdU+ Fancd2−/− cells. Nevertheless, PRIMPOL knockdown largely suppressed 5hmdCmediated PARylation, not reaching wildtype levels (fig. S12). In addition, PRIMPOL depletion partially suppressed 5hmdCinduced EdUPAR SIRF foci in Fancd2−/− cells (Fig. 8A). Moreover, PRIMPOL depletion significantly reduced 5hmdCinduced ssDNA gaps in Fancd2−/− cells, but not reaching levels seen in the untreated condition (Fig. 8B). These data suggest that a proportion of 5hmdCinduced ssDNA gaps depends on PRIMPOL. This situation contrasted to APEX1depleted cells, whereby APEX1 depletion completely abolished 5hmdCinduced ssDNA gaps in both backgrounds. These data indicate that, in addition to PRIMPOLmediated ssDNA gaps, 5hmdC induces other types of ssDNA gaps, probably arising from unrepaired APEX1mediated SSBs. Consistent with this, PRIMPOL partially accounted for the 5hmdCinduced lethality observed in Fancd2−/− cells (Fig. 8C). APEX1 PRIMPOL or HMCES PRIMPOL doubleknockdown cells displayed the same viability as single PRIMPOL–depleted Fancd2−/− cells (Fig. 8C). These data suggest that a subset of ssDNA gaps arising from cytotoxic HMCESDPCs is PRIMPOL dependent, whereas others are independently formed. We also examined the contribution of HMCESDPCs on nascent DNA to 5hmdCdependent SCEs. As previously reported for bulky adducts on template DNA strands (14), PRIMPOL depletion efficiently suppressed heightened 5hmdCdependent SCEs observed in Fancd2−/− cells. 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We are in debt to J. duxin (novo nordisk Foundation center for Protein Research, U. copenhagen, dK), G. P. crossan (Astra Zeneca, UK), and l. B. Pontel (instituto Josep carreras, Spain) for critical reading of the manuscript and M. Giannattasio of the iFOM dnA eM facility for assistance. Funding: this publication is part of the project Pid2021128988OBi00, funded by MiciU/Aei/10.13039/501100011033 and by eRdF/eU. this research work was also supported by cnS2022136055 financiado por Mcin/Aei/10.13039/501100011033/ UnióneuropeanextGenerationeU/PRtR. J.Y.M. is a tier i canada Research chair in dnA Repair and cancer therapeutics and was supported by a canadian institutes of health Research Foundation Grant (Fdn388879). v.c. received funding from AiRc under iG 2023id 28725 project. M.J.P.- G. was supported by consejería de transformación económica, industria, conocimiento y Universidades (PRedOc_00505). Y.R.- M. was supported by Pid2021128988OBi00/Mcin/Aei/10.13039/501100011033/FedeR, Ue. M.d.R.O. was supported by cnS2022136055/Mcin/Aei/10.13039/501100011033/Unión europea nextGenerationeU/ PRtR. Y.W.h. was supported by the 26596 AiRc fellowship for italy. S.B. is supported by an FRQS postdoctoral fellowship. Author contributions: M.J.P.- G.: investigation and methodology, data curation, validation, formal analysis, and visualization. Y.R.- M.: investigation, validation, formal analysis, and visualization. M.d.R.O.: investigation, methodology, validation, formal analysis, and visualization. Y.W.h.: investigation, methodology, data curation, validation, formal analysis, and visualization. S.B.: investigation and writing— review and editing. R.F.: Writing—review and editing and resources. J.Y.M.: Writing—review and editing, funding acquisition, and supervision. J.c.R.: investigation. v.c.: conceptualization, investigation, writing—review and editing, resources, funding acquisition, and supervision. i.v.R.: Writing—original draft, conceptualization, investigation, writing—review and editing, methodology, resources, funding acquisition, data curation, validation, supervision, formal analysis, project administration, and visualization. Competing interests: the authors declare that they have no competing interests. Data and materials availability: All data needed to evaluate the conclusions in the paper are present in the paper and/or the Supplementary Materials. Raw data are available at Zenodo (https://zenodo.org/records/14738090). Submitted 7 August 2024 Accepted 21 February 2025 Published 26 March 2025 10.1126/sciadv.ads3227 Downloaded from https://www.science.org at Universidad de Sevilla on May 12, 2025