Proteomic profiling of UV damage repair patches uncovers histone chaperones with central functions in chromatin repair

Domain: neuroscience
Published: 2026-01-28
Editorial outcome: major_revision

Abstract
DNA damage compromises not only genome stability but also chromatin integrity, which plays a central role in controlling cell identity. Chromatin repair entails the deposition of new histones at sites of DNA damage. How this is coordinated with parental histone dynamics for the maintenance of chromatin states is unknown. To bridge this knowledge gap, here we devise a novel proteomic strategy to characterize dynamic changes in the chromatin landscape during the repair of UV-induced DNA lesions in human cells, in a quantitative, unbiased and time-resolved manner. Thus, we identify the histone chaperones DNAJC9 and MCM2 as central players in chromatin repair. DNAJC9 provides new H3-H4 histones to CAF-1 and HIRA chaperones for deposition into chromatin and stimulates old H3-H4 histone recovery. MCM2 cooperates with DNAJC9 to coordinate old and new histone dynamics during UV damage repair. Together, our proteomic dataset provides a molecular framework for further dissecting epigenome maintenance mechanisms.

Introduction
Chromatin is made of DNA wrapped around histone proteins in the cell nucleus and provides epigenetic information that governs cell identity by modulating gene expression1. This pivotal information encompasses multiple layers of regulation, including DNA and histone modifications2,3 as well as histone variants4, which confer specific structural and functional features to the so-called epigenome. Chromatin is in a dynamic state owing to histone deposition and eviction by histone chaperones5, which can be assisted by chromatin remodelers6 in promoting histone dynamics. Histone post-translational modifications further contribute to chromatin plasticity via dedicated factors that deposit, erase, or read these marks2,7.

Epigenome stability is critical for the maintenance of cell function and homeostasis, as underlined by epigenome alterations in disease8,9. Nevertheless, maintaining epigenome stability is a challenge as all DNA metabolic activities entail some level of chromatin reorganization10,11. This challenge is particularly prominent during DNA damage repair because DNA damage can arise in an unscheduled manner at any time, anywhere in the genome12, which poses a major and unpredictable threat to both genome and epigenome stability. As such, chromatin plasticity is an integral part of the DNA damage response, DNA and chromatin repair being tightly coordinated13.

Histone dynamics during DNA repair have been extensively characterized in response to UV damage in mammalian cells and yet our knowledge of these fundamental mechanisms is still incomplete. Chromatin is transiently disrupted at UV damage sites with the eviction of H1 and H2A.Z histones14,15 while H3-H4 histones re-distribute away from damaged sites without dissociating from chromatin16. This local reorganization of chromatin is thought to increase chromatin accessibility, thus facilitating the recruitment of DNA repair factors. DNA repair is accompanied by a restoration of chromatin through the recovery of parental H3-H4 histones16 and the deposition of newly synthesized histones, as shown for H2A, H2A.X, H3.1, and H3.3 histone variants, all deposited de novo by specific histone chaperone complexes that act sequentially during the repair process. Deposition of new H3.3 histones by the histone chaperone complex HIRA occurs at early stages of UV damage repair, coupled to UV damage detection17, while the de novo deposition of H3.1 and H2A variants by the CAF-1 and FACT chaperone complexes, respectively, takes place during late repair steps, coupled to repair synthesis15,18,19. Repaired chromatin is thus a mixture of original and new information, brought about by parental histones that characterize the pre-damage chromatin, and by newly synthesized histones, respectively. Parental and new histone dynamics must be tightly coordinated to ensure the restoration of chromatin states but how this is achieved is a major unresolved issue.

Throughout the repair process, histone chaperones play a central role in controlling histone dynamics. These factors were identified through candidate approaches based on their cognate histone variants, which were shown to accumulate at sites of UV damage repair. We have now reached the limit of those approaches, and yet several important gaps remain in our knowledge of chromatin dynamics during DNA repair. In particular, (i) it is unknown which factors provide new histones to the chaperones identified so far for de novo deposition at repair sites, (ii) the chaperones regulating parental histone redistribution and recovery are uncharacterized, and therefore (iii) how parental and new histone dynamics are coordinated is still an open question. To fill these gaps, we have decided to adopt an unbiased strategy in order to identify new players in chromatin repair in an agnostic manner. We have devised a novel method for the analysis of proteins enriched at sites of UV damage repair in human cells with increased sensitivity and temporal resolution throughout the repair process. This method is based on the isolation of proteins on nascent DNA at repair sites (IPOND-R), combined with their characterization by mass spectrometry (MS)-based proteomics. With this method, we have identified histone chaperones that coordinate new H3-H4 deposition and parental H3-H4 recovery during chromatin repair.

Results
The IPOND-R method captures proteins at sites of DNA damage repair
To profile new chromatin interactors during the repair of DNA damage in human cells, we devised a novel approach based on the established IPOND methodology (Isolation of Proteins On Nascent DNA)20,21, allowing us to specifically identify proteins associated with DNA repair patches in an unbiased, quantitative and time-resolved manner. We called this method IPOND-R for Isolation of Proteins On Nascent DNA at Repair sites (Fig. 1a). In order to capture newly-synthesized DNA associated with DNA repair and not with DNA replication, IPOND-R includes a short-term treatment of cells with replication inhibitors, Hydroxyurea (HU) and Cytosine Arabinoside (Ara-C), for 2 h prior to DNA damage induction by global UVC irradiation. Cells are exposed to 10 J/m2 UVC, which generates UV lesions every 6 kb on average22 and broadly distributed through the genome. Cells are then pulsed with Ethynyl-deoxyUridine (EdU) to label repair patches, crosslinked at different time points after UV irradiation and EdU is biotinylated through a Click-it reaction. After sonication of chromatin, proteins associated with biotinylated nascent DNA are pulled down with streptavidin-coated beads for subsequent proteomic analyses (Fig. 1a). Since there are no major variations in UV damage repair during the cell cycle, this method can be run on asynchronous cell populations.

We verified that short term cell treatment with the replication inhibitor cocktail was sufficient to erase replicative DNA synthesis (Fig. 1b) while still allowing DNA repair synthesis at sites of UV damage, although with a decrease in efficacy (Fig. 1c). In line with this, we could detect a transient inhibition of gap filling at UV damage sites upon replication inhibition as shown by increased RPA levels 1 h after local UV irradiation returning to normal 2 h post UV (Supplementary Fig. 1a). Cells treated with replication inhibitors also showed signs of replicative stress as illustrated by a transient increase in γH2A.X and phospho-Chk1 signals and an altered cell cycle distribution with an accumulation of cells in G1 24 h post UV (Supplementary Fig. 1b–d). Nevertheless, replication inhibition did not impair the recruitment of the UV damage repair factor XPB (Xeroderma Pigmentosum, complementation group B) to damage sites (Supplementary Fig. 1e), and the kinetics of UV lesion removal (Supplementary Fig. 1f, g). Furthermore, we verified that cell treatment with replication inhibitors allowed efficient and timely recruitment of the histone chaperones CAF-1 and HIRA and subsequent deposition of newly synthesized H3.1 and H3.3 histone variants at UV damage sites (Supplementary Fig. 1h–k).

For EdU biotinylation, we used an enhanced version of biotin-azide conjugate called biotin picolyl azide, which improved in vivo biotinylation efficiency by 4.5-fold as verified by imaging with fluorescently-labeled streptavidin (Supplementary Fig. 2a). Note that DNA breakage arises during the biotinylation reaction due to copper-catalyzed hydrolysis of the DNA21 but as this occurs after cell fixation, it does not introduce any significant bias in the analysis. UV irradiation led to a detectable increase in biotinylated EdU (Supplementary Fig. 2b), thus supporting the sensitivity of the proposed methodology to monitor repair synthesis. Importantly, even though the UV damage repair machinery can excise EdU23, the UV-associated EdU signal was stable for up to 24 h post UVC irradiation in our experimental setting (Supplementary Fig. 2b), arguing against the death of UV-damaged, EdU-labeled cells between 1 h and 24 h post irradiation and allowing us to perform long-term analyses. Damage induction is quite homogeneous in a cell population (Supplementary Fig. 2c) so this does not introduce any bias in subsequent analyses. In addition, we verified that a 1 h EdU pulse immediately after UV irradiation allowed the capture of repair events in heterochromatin regions, known to repair UV lesions more slowly than euchromatin24,25. For this, we employed a validated murine cell line model, NIH/3T3 cells stably expressing the DNA damage sensor DNA damage binding protein 2 (DDB2)14, for studying UV damage repair in DAPI-dense pericentric heterochromatin domains. We observed that the relative levels of EdU in heterochromatin vs. euchromatin were comparable when we labeled early repair events (EdU pulse immediately after UV) or later repair events (EdU pulse 3 h after UV) (Supplementary Fig. 2d), indicating that the IPOND-R analysis is not biased towards repair in euchromatin regions.

As a proof-of-principle, we performed IPOND-R in human U2OS cells and analyzed by western-blot proteins associated with the repair patch 1 h post UV irradiation (Fig. 1d). Core histones, the damage-induced histone modification γH2A.X, PCNA and the histone chaperone CAF-1 were pulled down only in the presence of biotin and enriched in the capture samples from UV-treated cells compared to non-irradiated cells, demonstrating that IPOND-R efficiently and specifically captures chromatin repair patches. Altogether, these results indicate that the IPOND-R method is suited for studying chromatin dynamics specifically at sites of DNA damage repair, in different chromatin contexts and throughout the repair process, from early to late repair steps.

Identification of chromatin regulators enriched at sites of UV damage repair
In order to identify new players in chromatin repair, we combined IPOND-R with mass spectrometry-based quantitative proteomics. We detected 255 proteins that were significantly enriched at repair patches 1 h post UV irradiation in U2OS cells with a +/- UV fold change ≥ 2 and a limma p-value ≤ 0.01 (Fig. 2a, Supplementary Data 1). Among these proteins, we identified known UV damage repair factors (XPC, DDB1, CUL4A, PCNA) as expected, supporting the validity of the IPOND-R method in capturing UV damage repair patches. Much fewer proteins, 12, were retrieved as enriched at repair sites 24 h post UV irradiation and those were mostly core chromatin components (histones and high-mobility group proteins) (Fig. 2b, Supplementary Data 1). Comparing the 1 h and 24 h time points allows to eliminate factors constitutively bound to chromatin in order to focus on those that transiently bind to chromatin during repair (247 1 h-specific hits) (Fig. 2c, Supplementary Data 1). Gene ontology analysis of the 1 h-specific hits revealed an enrichment of biological processes associated with DNA and RNA metabolic activities, the nuclear periphery (nuclear pores, lamins), the immune system (isotype switching and somatic hypermutation), and chromatin organization (Fig. 2d, Supplementary Fig. 3a). We identified 194 proteins that were not found in previous proteomic studies post UV (Supplementary Fig. 3b, Supplementary Data 2)26,27. There was only a modest overlap with the dataset obtained by Stefos et al.26 in human skin fibroblasts 4 h post UV (41 proteins in common), which can be explained by the different cell types (U2OS osteosarcoma cells vs. 1BR3 skin fibroblasts) and different time points post UV (1 h vs. 4 h) as early repair responses cannot be captured 4 h post irradiation. We also compared the 1 h-specific hits identified by IPOND-R to the proteins identified on nascent replicated strands in undamaged conditions by IPOND in different human cell lines28,29. This analysis revealed several shared replication-associated factors including RFC1-5, PCNA, MCM2-7 but also highlighted important differences between the proteomes at replication forks and at sites of UV damage repair with replication-associated factors that were not retrieved at sites of UV damage repair, such as Cdc45, Claspin, MCM10, TONSL (Supplementary Fig. 3c, Supplementary Data 2). We confirmed by western blot that the replication-associated protein Cdc45 was not enriched in the IPOND-R eluate of the +UV condition (Supplementary Fig. 3d). To further study the chromatin regulators identified by IPOND-R, we crossed our dataset with the EpiFactors database (Fig. 2e). Histone modification activities (writers, readers, erasers) and histone chaperones were the most represented, followed by chromatin remodelers and DNA modifiers (Fig. 2e, f). We validated the presence of the chromatin remodeler catalytic subunit SMARCA5 and of the histone modifying enzyme HDAC1 in the +UV capture by IPOND-R coupled to western blot detection (Fig. 2g).

Among histone chaperones, we retrieved most of those known to be involved in chromatin repair following UV damage (Fig. 3a), with the three subunits of the CAF-1 complex (CHAF1A, CHAF1B, RBBP4), which deposits new H3.118, both subunits of the FACT complex (SUPT16H, SSRP1), which promotes new H2A.X deposition and ANP32E, which promotes H2A.Z eviction in UV-damaged chromatin15. In addition, we also detected additional histone chaperones with yet unknown functions in chromatin repair (Fig. 3a), namely Minichromosome Maintenance Complex Component 2 (MCM2) and DnaJ Heat Shock Protein Family Member C9 (DNAJC9). MCM2 was shown to chaperone H3-H4 histones at replication forks30,31 and the heat-shock protein DNAJC932 was recently characterized as an evolutionarily conserved H3-H4 histone chaperone33–35. Notably, our optimized IPOND-R methodology permits the unprecedented detection of DNAJC9 (+/- UV fold change = 2.36; limma p-value = 1.53E−3 at 1 h post UV) and MCM2 (+/- UV fold change = 3.07; limma p-value = 6.16E−4 at 1 h post UV) at sites of UV damage repair while these chaperones were not identified in previous proteomic analyses post UV26,27. All the histone chaperones that we identified on UV damaged chromatin were captured at 1 h and not 24 h post UV (Fig. 3a, b), highlighting their dynamic recruitment to chromatin during the repair process. We validated these findings by western blot on the capture material in U2OS cells (Fig. 3c) and in non-cancerous RPE-1 cells (Fig. 3d), which confirmed the transient recruitment of DNAJC9 to repair sites. Western blot analysis of the capture material also enabled the detection of the histone chaperone HIRA (Fig. 3c), which was not identified by mass spectrometry-based proteomics. We further validated DNAJC9 and MCM2 recruitment to sites of UV damage repair through an orthogonal approach that does not rely on using replication inhibitors. For this, we imaged these histone chaperones by immunofluorescence in cells exposed to local UVC irradiation. First, we assessed DNAJC9 enrichment at sites of UVC laser-induced damage in U2OS cells expressing a GFP-tagged form of the UV damage sensor protein DDB2. DNAJC9 showed a distinct and significant enrichment of about 2-fold at UV damage sites (Fig. 3e). We confirmed this result by irradiating cells with a UVC lamp through micropore filters and observed a similar albeit more modest enrichment of DNAJC9 and MCM2 at UV damage sites (1.2-fold, Fig. 3f, g, Supplementary Fig. 4a). The recruitment of DNAJC9 and MCM2 to UV damage sites was observed outside S phase (Supplementary Fig. 4b), pointing to a replication-independent function of these histone chaperones at UV damage sites. Collectively, these findings highlight the potential of IPOND-R in identifying chromatin-associated factors at sites of UV damage repair, including the histone chaperones DNAJC9 and MCM2.

The histone chaperone DNAJC9 is not involved in DNA repair following UV damage
To study the function of DNAJC9 and MCM2 in the DNA damage response, we first focused on DNAJC9 as it was the least characterized histone chaperone. We employed siRNA-mediated depletion because DNAJC9 knock-out was reported to be lethal in human cells33,34, and we reached an effective depletion of DNAJC9 of over 50% in U2OS cells (Supplementary Fig. 5a). Since DNAJC9 was recruited early during the DNA damage repair process, we addressed whether this histone chaperone may participate, directly or indirectly, in the repair of DNA damage. However, we did not observe any significant effect of DNAJC9 depletion on the recruitment of the UV-damage repair factor XPB, involved in the unwinding of the DNA double-helix prior to damage excision (Supplementary Fig. 5b), nor on DNA repair synthesis, monitored by EdU incorporation at UV damage sites (Supplementary Fig. 5c). Furthermore, the kinetics of UV photoproduct removal (CPD and 6,4-PP) were also unchanged upon DNAJC9 knock-down while depletion of the repair factor XPG prevented UV lesion removal (Supplementary Fig. 5d). In line with this, DNAJC9 depletion did not impair the survival of cells to UV irradiation (Supplementary Fig. 5e). Knocking down DNAJC9 also did not impair transcription recovery at sites of UV damage as monitored by ethynyl-uridine incorporation into nascent transcripts, whereas depleting the transcription-coupled repair factor CSB (Cockayne Syndrome group B) inhibited transcription recovery, as expected (Supplementary Fig. 5f). Taken together, these results rule out a function for DNAJC9 in the global genome and transcription-coupled repair of DNA damage and prompted us to explore a direct role of this chaperone in histone dynamics at sites of UV damage repair.

DNAJC9 promotes the deposition of newly synthesized H3 variants at sites of UV damage repair
Given the H3-H4 histone binding activity of DNAJC9, we investigated whether this histone chaperone may impact the deposition of newly synthesized H3 histone variants at UV damage sites. We employed the SNAP-tag technology to track new H3.1 and H3.3 histones in U2OS cells stably expressing SNAP-tagged H3 variants36 and monitored their accumulation at sites of local UVC irradiation upon depletion of DNAJC9 (Fig. 4a). Given that H3.1 is also de novo deposited at replication foci37, the incorporation of new H3.1 histones was analyzed outside S-phase to focus on repair-coupled deposition. We can easily discriminate between cells in and outside S phase because the nuclear levels of new H3.1 are significantly higher in S phase (Supplementary Fig. 6a). We observed a marked and reproducible decrease of new H3.1 and H3.3 levels at sites of UV damage upon DNAJC9 depletion (Fig. 4b, c), which we confirmed with a second siRNA targeting DNAJC9 (Supplementary Fig. 6b, c). Exogenous expression of siRNA-resistant DNAJC9 rescued new H3.3 deposition at UV damage sites in DNAJC9-knocked down cells (Supplementary Fig. 6d) which was not observed upon expression of a histone binding mutant form of DNAJC934 although both wild type and mutant DNAJC9 were similarly recruited to damage sites (Fig. 4d). These experiments establish that DNAJC9, through its histone binding domain, promotes the deposition of newly synthesized H3 variants at UV damage sites. We recapitulated these results in non-cancerous RPE-1 cells stably expressing SNAP-tagged H3 variants (Supplementary Fig. 6e, f). Contrary to what observed with H3 histone variants, DNAJC9 depletion did not affect the de novo deposition of the H2A.X variant at UV damage sites (Supplementary Fig. 6g), which is fully consistent with the H3-H4 specificity of this histone chaperone33,34. Importantly, DNAJC9 depletion did not affect the total protein levels of the histone chaperones CAF-1 and HIRA (Supplementary Fig. 6b, c), known to promote the de novo deposition of H3.1 and H3.3 in UV-damaged chromatin (Fig. 4e), and also did not impair the recruitment of these chaperones to UV damage sites (Fig. 4f, g). From these results, we conclude that DNAJC9 promotes new H3 variant deposition at sites of UV damage repair without affecting their cognate histone chaperones CAF-1 and HIRA.

In addition to a role at damage sites, we suspected that DNAJC9 may also promote new histone deposition in undamaged chromatin, during replication in particular, because this histone chaperone sustains cell cycle progression into S phase (Supplementary Fig. 6h), which is consistent with its reported function in cell proliferation33. Similar to what was observed at UV damage sites, DNAJC9 depletion impaired the incorporation of newly synthesized H3.1 histones at replication foci in S phase and also the replication-independent deposition of H3.3 in undamaged conditions (Supplementary Fig. 6a, i). These results highlight a functional role of DNAJC9 in chromatin assembly in vivo via the deposition of new H3.1 and H3.3 histones, both in basal conditions and in response to DNA damage.

We next sought to characterize whether DNAJC9 may function in the same histone deposition pathways as the CAF-1 and HIRA chaperones at repair sites. To do so, we compared single and double knock-downs of DNAJC9 and CAF-1 or HIRA (Supplementary Fig. 6j–m) for their impact on new H3 histone deposition at UV damage sites (Fig. 4h, i). Single depletions of DNAJC9 and CAF-1 similarly impaired the incorporation of new H3.1 histones at damage sites (Fig. 4h) and single depletions of DNAJC9 and HIRA had comparable effects on new H3.3 deposition at UV sites (Fig. 4i). The impact on new H3 variant deposition was partial in each condition, and we did not observe any additive effect upon double depletion of DNAJC9 and the other chaperones (Fig. 4h, i). These results establish that DNAJC9 functions in the same histone deposition pathways as CAF-1 and HIRA to promote the incorporation of new H3.1 and H3.3 histones at sites of UV damage repair. In contrast to CAF-1 and HIRA, DNAJC9 is loosely bound to UV-damaged chromatin, as it is mostly solubilized by detergents (Supplementary Fig. 6n). Together, these findings suggest an upstream function of DNAJC9 in the supply of newly synthesized histones to CAF-1 and HIRA chaperones.

DNAJC9 promotes the recovery of parental H3 histones at sites of UV damage repair
We next investigated whether, besides its activity in promoting new histone deposition at repair sites, DNAJC9 may also control the recovery of old H3-H4 histones that redistribute away from damaged chromatin regions16. The only factor known so far to control parental histone dynamics at UV sites is the UV damage sensor protein DDB2, whose binding to damaged chromatin and subsequent release promote the redistribution and recovery of parental histones, H3.3 respectively16. To test DNAJC9 contribution to these dynamics, we used the SNAP-tag technology to mark old H3.3 histones in U2OS cells stably expressing H3.3-SNAP, and quantified their levels at UV damage sites compared to the entire nucleus in cells fixed at different time points post irradiation as a proxy for measuring old histone redistribution and recovery (Fig. 5a). The depletion of old H3.3 histones at UV damage sites early after irradiation (10 min) and their subsequent enrichment at late time points (8 h) reflect parental histone redistribution and recovery, respectively (Fig. 5b, siLuc condition). The ratio of old histones at UV damage sites relative to the nucleus 8 h post UV in siLuc cells sets the level of recovery of old histones in control condition. Comparatively, although not completely abrogated, this recovery was impaired upon siRNA-mediated knock-down of the late repair factor XPG, as expected, because UV damage repair is compromised by reduced XPG levels (Fig. 5b, siXPG condition). Importantly, DNAJC9-knocked down cells showed a similar phenotype with an impaired recovery of old histones (Fig. 5b). This phenotype cannot be attributed to defective UV damage repair because these cells were fully proficient for the repair of UV lesions (Supplementary Fig. 5). These results thus demonstrate that DNAJC9 promotes parental H3.3 histone recovery at sites of UV damage repair. To test whether the defective recovery of parental histones may result from impaired DDB2 release from chromatin, we examined DDB2 dynamic binding to UV-damaged chromatin in the absence of DNAJC9 in U2OS cells stably expressing GFP-tagged DDB2. While XPG depletion prevented DDB2 release by inhibiting DNA repair progression, as expected, the depletion of DNAJC9 had no effect on DDB2 release from chromatin (Fig. 5c). Together, these findings demonstrate that the histone chaperone DNAJC9 promotes parental H3 histone recovery at repair sites independently of the UV damage sensor DDB2. Given that DNAJC9 promotes both new histone deposition and old histone recycling at UV-damaged sites, we reasoned that DNAJC9 may control the overall histone density at repair sites. We used the SNAP-tag technology to mark all H3.3-SNAP histones 8 h post irradiation (Fig. 5d) and observed a modest but significant decrease in the total levels of H3.3-SNAP histones at UV sites when DNAJC9 was depleted (Fig. 5e, f). Therefore, DNAJC9 maintains histone density at sites of UV damage repair through new histone deposition and old histone recycling.

DNAJC9 cooperates with MCM2 to coordinate old and new histone dynamics at sites of UV damage repair
We next investigated how DNAJC9 may promote old histone recovery at repair sites. We envisioned that DNAJC9 could cooperate with the MCM2 histone chaperone in this process because MCM2 is engaged in histone-dependent interactions with DNAJC934, and promotes the recycling of parental H3-H4 histones at replication forks30. We first tested by immunofluorescence in cells exposed to local UVC irradiation whether MCM2 was still enriched on damaged chromatin at the time of old histone recovery. MCM2 enrichment at UV damage sites was clearly detectable 8 h post irradiation, even more pronounced than at 1 h post irradiation (Fig. 6a, siLuc conditions), and we verified that this was not due to bleed-through from the CPD channel (Supplementary Fig. 7a). MCM2 nuclear levels and recruitment to sites of UV damage repair were unaffected by DNAJC9 knock-down and MCM2 depletion did not impair DNAJC9 nuclear levels and enrichment at repair sites (Supplementary Fig. 7b, Fig. 6a, b), arguing that both histone chaperones are independently recruited to UV damage sites. We next tested if MCM2 could function together with DNAJC9 in repair-coupled histone dynamics. Similar to DNAJC9 depletion, MCM2 depletion by siRNA impaired old H3.3 recovery at sites of UV damage repair (Fig. 6c) and the dual knock-down of both chaperones did not show any additive effect (Fig. 6d and Supplementary Fig. 7c). Thus, MCM2 cooperates with DNAJC9 to promote old H3 recovery at repair sites. We obtained comparable results when examining new H3.3 deposition at earlier times following UV irradiation, arguing that MCM2 also cooperates with DNAJC9 to stimulate new histone deposition at sites of UV damage repair (Fig. 6e and Supplementary Fig. 7d). This is consistent with the reported interaction of MCM2 with both new and old H3-H4 histones30. Similar to what was observed in cycling cells, DNAJC9 and MCM2 knock down in cells arrested in G1 by palbociclib treatment impaired new H3.3 deposition at sites of UV damage repair (Supplementary Fig. 7e), arguing that the function of these chaperones in new histone deposition coupled to repair does not require passage through S phase. We validated the contribution of MCM2 to new histone deposition at repair sites in functional rescue experiments, showing that exogenous expression of siRNA-resistant MCM2 restored at least partially new H3.3 deposition at UV damage sites in MCM2 knocked down cells (Supplementary Fig. 7f). This is an important finding because it demonstrates the role of MCM2 in new H3.3 deposition at sites of UV damage repair. We also ruled out the hypothesis that MCM2 and DNAJC9 contribution to old H3.3 recovery may be an indirect consequence of these chaperones stimulating new H3.3 deposition. Indeed, depletion of the H3.3 chaperone HIRA, known to deposit new H3.3 histones at sites of UV damage repair (Fig. 4i)17 did not impair old H3.3 recovery (Supplementary Fig. 7g). From these results, we conclude that the DNAJC9 and MCM2 histone chaperones play a central function in chromatin repair by coordinating old and new histone dynamics.

Methods
Cell culture
U2OS (ATCC HTB-96, human osteosarcoma, female), RPE-1 hTERT (ATCC CRL-4000, human retinal pigment epithelial cell, female), and NIH/3T3 cells (ATCC CRL-1658, mouse embryonic fibroblast, male) were grown at 37 °C and 5% CO2 in Dulbecco’s modified Eagle’s medium (DMEM, Invitrogen) supplemented with 10% fetal bovine serum (EUROBIO) and antibiotics (100 U/ml penicillin and 100 µg/ml streptomycin, Invitrogen) and the appropriate selection antibiotics for stable cell lines (Euromedex, Table 1). For seeding NIH/3T3 cells on coverslips, coverslips were first coated with 20 µg/ml Collagen Type I (MERCK Millipore) and 2 µg/ml fibronectin (Sigma-Aldrich) to increase cell adhesion.

In order to inhibit DNA replication, we use 10 mM Hydroxyurea (HU, Sigma-Aldrich) and 1 mM cytosine arabinoside (Ara-C, Sigma-Aldrich) in fresh growth medium for 2 h prior to DNA damage induction. To prevent S-phase entry, we use the cdk4/6 inhibitor palbociclib (Selleck Chemicals) at 150 nM final for 48 h. Palbociclib is applied to RPE-1 cells only because U2OS cells are resistant due to high cdk4 expression.

All cell lines generated in this study were authenticated by verifying the expression of the transgenes by immunofluorescence and western blot. They are available for reuse and can be provided upon request.

siRNA and plasmid transfections
siRNA purchased from Eurofins MWG Operon, Sigma Aldrich and Dharmacon (Table 2) were transfected into cells using Lipofectamine RNAiMAX (Invitrogen) following the manufacturer’s instructions. The final concentration of siRNA in the culture medium was 50 nM. Cells were harvested 48-72 h after transfection.

Cells were transfected with plasmid DNA (Table 3) using Lipofectamine 2000 (Invitrogen) or by nucleofection with the Neon™ Transfection System (Thermo Fisher Scientific) according to the manufacturer’s instructions. For stable cell line establishment (Table 1), plasmid DNA was transfected into cells at 1 µg/ml final, 48 h before antibiotic selection of clones. For rescue experiments, cells were co-transfected with plasmid DNA (3 µg/ml final) and siRNA (50 nM final), using the NeonTM Transfection System (Thermo Scientific) according to the manufacturer’s instructions and harvested 72 h later.

All plasmids generated in this study are available for reuse and can be provided upon request.

UVC irradiation
Cells grown on glass coverslips (12 mm diameter, thickness No. 1.5, Thorlabs) were irradiated with UVC (254 nm) using a low-pressure mercury lamp. Conditions were set using a VLX-3W dosimeter (Vilber Lourmat). For global UVC irradiation, cells in Phosphate Buffer Saline (PBS) were exposed to UVC doses ranging from 4 to 12 J/m2 for survival assays and to 10 J/m2 in other experiments. For local UVC irradiation51,52, cells were covered with a polycarbonate filter (5 μm pore size, Millipore) and irradiated with 150, 300 or 500 J/m2 UVC. Irradiated cells were allowed to recover in culture medium for the indicated times before fixation.

For UVC laser micro-irradiation53, cells were grown on quartz coverslips (25 mm diameter, thickness No. 1, SPI supplies) and nuclei were stained by adding Hoechst 33258 (10 μg/mL final, Sigma-Aldrich) to the culture medium 30 min before UVC irradiation. Quartz coverslips were transferred to a Chamlide magnetic chamber (Gataca-systems) on a custom stage insert (Live Cell Instrument) and cells were irradiated for 100 ms using a 2 mW pulsed diode-pumped solid-state laser emitting at 266 nm (Rapp OptoElectronics, Hamburg GmbH) directly connected to a Zeiss LSM900 confocal microscope adapted for UVC transmission with all-quartz optics. The laser was attenuated using a 0.1% neutral density filter and focused through a 40x/0.6 Ultrafluar glycerol objective with quartz lenses. The laser is controlled by a UGA Firefly module with SysCon 2 software (Rapp Opto). The laser impact has an average size of 1 μm in diameter and damages around 1-2% of the total nuclear volume. The corresponding UVC dose, not directly measurable but estimated by comparing the intensity of the CPD damage generated by the laser and the UVC lamp, is 800 J/m2.

IPOND-R (Isolation of Proteins On Nascent DNA at Repair sites)
EdU labeling of repair sites. Cells are seeded in 15-cm dishes to harvest at least 70 million cells per condition on the day of the experiment. In order to label only repair patches, replication is inhibited with 10 mM HU and 1 mM Ara-C (Sigma-Aldrich) in fresh growth medium for 2 h at 37 °C prior to global UVC irradiation (10 J/m2) in PBS supplemented with 10 mM HU and 1 mM Ara-C to maintain the inhibition of replication. A non-irradiated sample is used as negative control. After irradiation, cells are incubated in fresh medium containing the replication inhibitor cocktail and 10 µM Ethynyl-deoxyUridine (EdU, Eurodemex) for 1 h at 37 °C. At this step, a chase can be performed in fresh medium supplemented with 10 µM Thymidine (Sigma-Aldrich).

Cell fixation and permeabilization. Cells are fixed with 1% Formaldehyde (Sigma-Aldrich) for 15 minutes on a rocking platform. The fixation is stopped by adding 1.33 M Glycine (Sigma-Aldrich) for 5 minutes on the rocking platform. Cells are washed twice in cold PBS, scrapped in 1% BSA solution in PBS and pelleted for 5 min at 300 g at 4 °C. Cell pellets are permeabilized with 1% Triton X-100 (1 mL/ 10 million cells, Euromedex) for 30 min at room temperature and washed twice with cold 1% BSA solution.

EdU biotinylation by Click-it chemistry. Cell pellets are resuspended in a Click-it reaction cocktail containing 2 mM CuSO4 (Sigma-Aldrich), 1 mM Tris((1-hydroxy-propyl-1H-1,2,3-triazol-4-yl)methyl)amine (THPTA, Euromedex), 10 mM Sodium Ascorbate (Sigma-Aldrich), 5 mM Amino-guanidine hydrochloride (Sigma-Aldrich), and 10 μM Biotin Picolyl azide or Biotin azide (Sigma-Aldrich). Cell suspensions are left on a rotating wheel at room temperature for 1 h. Cells are then washed twice in cold 1% BSA solution containing 0.5X EDTA-free Protease inhibitor cocktail (PIC, Roche), and once in cold PBS. A negative control without biotin reagent was performed to check for the specificity of the Streptavidin pull-down.

Cell lysis and sonication. Cell pellets are resuspended in lysis buffer (200 µl per 15 million cells) containing 1% SDS, 50 mM Tris pH 7.5, and 1X PIC. The cell suspensions are transferred to 1.5 ml sonication tubes (Diagenode) and sonication is performed using a Bioruptor Pico (Diagenode) with 5-10 cycles of 30 sec ON and 30 sec OFF. The sonicated samples are transferred to new Eppendorf tubes and centrifuged 10 min at 16,000 g at 4 °C. The supernatants are transferred to Low Protein Binding tubes (Thermo Fisher Scientific).

Analysis of DNA shearing. A 20 µl aliquot of each cell lysate is incubated with 200 mM NaCl and 200 µg/ml RNAse A (Millipore) overnight at 65 °C. After addition of 100 µg/ml Proteinase K (Sigma-Aldrich), samples are incubated for a further 2 h at 45 °C prior to DNA purification using the PCR purification kit from Macherey-Nagel following the manufacturer’s instructions. DNA samples are analyzed with a TapeStation (Agilent) to check that the average fragment size is about 500 bp.

Streptavidin capture of biotin-labeled nascent DNA and associated proteins. The remaining solution is resuspended in an equal volume of cold PBS supplemented with 1X PIC to dilute the SDS present in the sonication buffer. At this stage, input samples are collected, diluted in Laemmli buffer (50 mM Tris-HCl pH 6.8, 1.6% Sodium Dodecyl Sulfate (SDS), 8% glycerol, 4% β-mercaptoethanol, 0.0025% bromophenol blue) and decrosslinked by heating for 5 min at 95 °C.

For streptavidin capture, Dynabeads MyOne Streptavidin-C1 beads (Invitrogen) are washed in lysis buffer and in PBS supplemented with 1X PIC and then are added to cell lysates (1 µl beads per 1 million cells) and incubated overnight at 4 °C on a rotating wheel. After capture, beads are washed once in lysis buffer (1 ml wash solution per 10 million cells), once in 1 M NaCl supplemented with 1X PIC, followed by 3 additional washes in lysis buffer.

For western blot analysis, the beads are resuspended in Laemmli buffer and decrosslinking is achieved by heating for 5 min at 95 °C. For mass spectrometry analysis, beads are transferred to new Low Protein binding tubes and washed in 50 mM Tris-HCl pH 8 before resuspension in Laemmli buffer and decrosslinking by heating for 5 min at 95 °C. Ten percent of the protein solution is kept for a Silver staining analysis.

Mass spectrometry-based proteomic analyses
Proteins from three and four IPOND-R biological replicates, purified 1 h and 24 h after UV irradiation, respectively (-UV, negative control) were solubilized in Laemmli buffer (50 mM Tris-HCl pH 6.8, 1.6% Sodium Dodecyl Sulfate (SDS), 8% glycerol, 4% β-mercaptoethanol, 0.0025% bromophenol blue) and separated by SDS-PAGE (Mini Protean TGX, 4-20%, Biorad), before staining with Coomassie blue R-250 (BioBasic Canada Inc.). The band corresponding to streptavidin was discarded and the remaining proteins were digested in-gel using modified trypsin (Promega, sequencing grade) as previously described54. The resulting peptides were analyzed by online nano liquid chromatography coupled to MS/MS (Ultimate 3000 RSLCnano and Q-Exactive HF, Thermo Fisher Scientific, for the 1 h dataset, and Ultimate 3000 RSLCnano and Orbitrap Exploris 480, Thermo Fisher Scientific, for the 24 h dataset) using a 120-min gradient. For this purpose, the peptides were sampled on a precolumn (300 μm × 5 mm PepMap C18, Thermo Scientific) and separated in a 75 μm × 250 mm C18 column (Reprosil-Pur 120 C18-AQ, 1.9 μm, Dr. Maisch, for the 1 h dataset, and Aurora Generation 2, 1.6 µm, Ion Opticks, for the 24 h dataset). The MS and MS/MS data were acquired using Xcalibur (Thermo Fisher Scientific).

Peptides and proteins were identified by Mascot (version 2.8.0, Matrix Science) through concomitant searches against the Uniprot database (20230719 download, 207’981 sequences) and a homemade database containing the sequences of classical contaminant proteins found in proteomic analyses (human keratins, trypsin…, 250 sequences). Trypsin/P was chosen as the enzyme and two missed cleavages were allowed. Precursor and fragment mass error tolerances were set at respectively at 10 and 20 ppm. Peptide modifications allowed during the search were: Carbamidomethyl (C, fixed), Acetyl (Protein N-term, variable) and Oxidation (M, variable). The Proline software55 version 2.2.0 was used for the compilation, grouping, and filtering of the results (conservation of rank 1 peptides, peptide length ≥ 6 amino acids, false discovery rate of peptide-spectrum-match identifications <1%56, and a minimum of one specific peptide per identified protein group). MS data have been deposited to the ProteomeXchange Consortium via the PRIDE partner repository57 with the dataset identifier PXD055129. Proline was then used to perform an MS1 label-free quantification of the identified protein groups based on razor and specific peptides.

Statistical analysis was performed using the ProStaR software58 based on the quantitative data obtained with the different biological replicates analyzed for each condition. Proteins identified in the contaminant database, proteins identified by MS/MS in less than two replicates of one condition, and proteins quantified in less than all replicates of one condition were discarded. After log2 transformation, abundance values were normalized using the variance stabilizing normalization (vsn) method applied condition-wise, before missing value imputation (SLSA algorithm for partially observed values in the condition and DetQuantile algorithm for totally absent values in the condition).

Statistical testing was conducted with limma, whereby differentially expressed proteins were selected using a log2(Fold Change) cut-off of 1 and a p-value cut-off of 0.01, allowing to reach a false discovery rate inferior to 3% according to the Benjamini-Hochberg estimator. Proteins found differentially abundant but identified by MS/MS in less than two replicates, and detected in less than all replicates, in the condition in which they were found to be more abundant were invalidated (p-value = 1).

Data visualization and gene ontology analysis
Biological process gene ontology terms were extracted from the PANTHER database (v.18 2023-08-01 release) and analyzed through the PANTHER Overrepresentation Test (released 2023-07-05). Significance was calculated using a Fisher test with a false-discovery rate correction. The list of EpiFactors was extracted from https://epifactors.autosome.ru/public_data version 2.059,60 curated based on most recent literature. The proteins detected by IPOND-R coupled to mass spectrometry were visualized using Volcano plots generated with Rstudio 2021.09.0 using ggplot2 version 3.3.5, ggrepel version 0.9.1, and dplyr version 1.1.2. Venn diagrams were generated with eulerr version 6.1. Statistical analyses and all other graphs were generated using GraphPad Prism version 8.4.3, and test details and p-values are referred to in the figure legends.

EdU-labeling of replicating cells and repair sites
To visualize replication foci, 10 µM Ethynyl-deoxyUridine (EdU) was incorporated into cells on glass coverslips during 15 min at 37 °C and revealed using the Click-It EdU Alexa Fluor 488 or 594 Imaging kit (Invitrogen) according to the manufacturer’s instructions. To localize the sites of UV damage repair, cells were incubated with 10 µM EdU for 1 h after local UVC irradiation and EdU was revealed using the Click-It EdU Alexa Fluor 488, 594 or 647 Imaging kit (Invitrogen).

EU-labeling of nascent transcripts at UV damage sites
Cells were exposed to local UVC irradiation (300 J/m2) 48 h post siRNA transfection and allowed to recover for 2 h (to monitor transcription inhibition at UV sites) or 24 h (to monitor transcription recovery). For nascent transcript labeling, cells were incubated in DMEM supplemented with 0.5 mM ethynyl-uridine (EU) for 45 min at 37 °C, and rinsed in PBS before fixation in 2% paraformaldehyde (Electron Microscopy Sciences). EU incorporation was revealed with Click-iT RNA Imaging kit (Thermo-Fisher Scientific) using Alexa Fluor 594 dye according to the manufacturer’s instructions. Cells were post-fixed in 2% paraformaldehyde before CPD immunostaining with AlexaFluor 488 dye as described in the immunofluorescence section below. Coverslips were mounted in Vectashield medium with DAPI (Vector laboratories). The mean EU fluorescence intensity was measured at UV damage sites (delineated based on CPD staining) relative to the entire nucleus (delineated based on DAPI staining).

Immunofluorescence
Cells grown on coverslips were either fixed directly with 2% paraformaldehyde (Electron Microscopy Sciences) for 10 min and permeabilised for 5 min with 0.5% Triton X-100 in PBS or cells were pre-extracted before fixation with 0.5% Triton X-100 in CSK buffer (Cytoskeletal buffer: 10 mM PIPES pH 7.0, 100 mM NaCl, 300 mM sucrose, 3 mM MgCl2) for 5 min at room temperature to remove soluble proteins. For the detection of UVC photoproducts, the DNA was denatured with 2 N HCl for 10 min at 37 °C (6,4-PP detection) or with 0.5 M NaOH for 5 min at room temperature (CPD detection). Since this denaturation quenches GFP fluorescence, when CPD detection was combined with the visualization of GFP-DDB2, immunofluorescence was performed with GFP immunodetection using a rat anti-GFP antibody (Supplementary Table 1) combined with CPD immunodetection. Samples were blocked for 10 min in 5% BSA (Bovine Serum Albumin, Sigma-Aldrich) in PBT (PBS 0.1% Tween-20), followed by 45 min incubation with primary antibodies and 30 min incubation with secondary antibodies coupled to Alexa Fluor dyes (Supplementary Tables 1 and 2) diluted in blocking buffer. Coverslips were mounted in Vectashield medium with DAPI (Vector laboratories).

SNAP-tag labeling of histones
For specific labeling of newly synthesized histones36,61, cells were grown on glass coverslips and pre-existing SNAP-tagged histones were first quenched by incubating cells with 10 μM of the non-fluorescent substrate SNAP-cell Block (New England Biolabs) for 30 min followed by a 30 min-wash in fresh medium and a 2 h (U2OS) or 4 h (RPE-1) chase. The new SNAP-tagged histones synthesized during the chase were fluorescently labeled with 2 μM of the red-fluorescent reagent SNAP-cell TMRstar (New England Biolabs) during a 15 min-pulse step followed by 30 min wash in fresh medium. Cells were subsequently permeabilised with Triton X-100, fixed and processed for immunostaining. Cells were irradiated with a UVC lamp before the pulse step.

For specific labeling old histones, cells were grown on glass coverslips and pre-existing SNAP-tagged histones were fluorescently labeled with 2 μM of the red-fluorescent reagent SNAP-cell TMRstar (New England Biolabs) during a 30 min-pulse step followed by 30 min wash in fresh medium. Cells were irradiated with a UVC lamp 48 h after the pulse step. Cells were subsequently permeabilised with Triton X-100, fixed and processed for immunostaining.

For labeling all histones, cells were grown on glass coverslips and SNAP-tagged histones were fluorescently labeled with 2 μM of the red-fluorescent reagent SNAP-cell TMRstar (New England Biolabs) during a 30 min-pulse step. Cells were subsequently permeabilised with Triton X-100, fixed and processed for immunostaining. Cells were irradiated with a UVC lamp before the pulse step.

Image acquisition and analysis
Fluorescence imaging was performed with a Leica DMI6000 epi-fluorescence microscope using a Plan-Apochromat 40x/1.3 oil objective. Images were captured using a CCD camera (Photometrics) and Metamorph software (version 7.10.2.240). Images were assembled with Adobe Photoshop. Live cell imaging coupled to UVC laser micro-irradiation was performed using a 40x/0.6 Ultrafluar Glycerol objective on a Zeiss LSM900 confocal microscope. Images were captured using Zen blue software version 3.5, and analyzed with ImageJ v.2.0.0-rc-69/1.52.p (U.S. National Institutes of Health, Bethesda, Maryland, USA, http://imagej.nih.gov/ij/). The subtract background function was applied to all images prior to quantification, and images were smoothed using the median filter function. Nuclei and heterochromatin domains were segmented based on DAPI or Hoechst staining, and UVC-damaged regions based on GFP-DDB2 fluorescence or immunostaining for UV damage or repair factors using the default threshold function with manual adjustment. Mean intensities were measured in each region of interest using the measure and set measurement functions. When calculating the ratios of histone signals at UV sites relative to the nucleus, a correction factor of 1.19 was applied to nuclear histone signals to take into account lower histone content in nucleoli as measured in independent experiments with histone and nucleolin co-staining.

Cell extracts and Western blot
Total extracts were obtained by scraping cells on plates or resuspending cell pellets in Laemmli buffer (50 mM Tris-HCl pH 6.8, 1.6% Sodium Dodecyl Sulfate (SDS), 8% glycerol, 4% β-mercaptoethanol, 0.0025% bromophenol blue) followed by 5 min denaturation at 95 °C.

For western blot analysis, extracts were run on 4%–20% Mini-PROTEAN TGX gels (Bio-Rad) in running buffer (200 mM glycine, 25 mM Tris, 0.1% SDS). Proteins were transferred onto nitrocellulose membranes (0.2 μm, Amersham Protran) for 30 min at 15 V with a Trans-Blot SD semidry transfer cell (Bio-Rad) in transfer buffer (50 mM Tris, 40 mM glycine, 0.04% SDS, 15% ethanol). Total proteins were revealed by Reversible Stain (Pierce). Proteins of interest were probed using the appropriate primary and Horse Radish Peroxidase (HRP)-conjugated secondary antibodies (Supplementary Tables 1 and 2), detected using SuperSignal West Pico or Femto chemiluminescence substrates (Pierce) on hyperfilms MP (Amersham) or with an Odyssey Fc-imager (LI-COR Biosciences). When fluorescence detection was used instead of chemi-luminescence, secondary antibodies were conjugated to IRDye 680RD or 800CW (Supplementary Table 2), membranes were scanned with the Odyssey Fc-imager (LI-COR Biosciences) and analyzed with Image Studio Lite software (version 5.2.5) using total protein stain for normalization.

Flow cytometry
For cell cycle analysis, cells were fixed in ice-cold 70% ethanol before DNA staining with 50 ug/ml propidium iodide (Sigma-Aldrich) in PBS containing 0.05% Tween and 0.5 mg/ml RNase A (USB/Affymetrix). DNA content was analyzed by flow cytometry using a BD FACScalibur flow cytometer equipped with CellQuest Pro version 6.0 (BD Biosciences) and FlowJo software version 9.7.5 (TreeStar).

Cell survival assays
Cells were replated 48 h after siRNA transfection and exposed to global UVC irradiation (4, 8, and 12 J/m2) the following day. Colonies were stained 12 days later with 0.5% crystal violet/20% ethanol and counted. Results were normalized to plating efficiencies.

Statistical analyses
Percentages of positively stained cells were obtained by scoring at least 150 cells in each experiment. Statistical tests were performed using GraphPad Prism (versions 8 to 10). We run the statistical analyses on the means of the biological replicates in conditions of global UV irradiation, which lead to homogeneous damage induction in the cell population, while analyses are performed on all data points and not on the means of the biological replicates in conditions of local UV irradiation because in this case each nucleus receives a different UV dose, which depends on the pore distribution in the micropore filters used for local UVC irradiation, so each nucleus should be considered independently. P-values for mean comparisons between two groups were calculated with a Student’s t test with Welch’s correction when necessary. Multiple comparisons were performed by one-way or two-way ANOVA. Comparisons of clonogenic survival were based on non-linear regression with a polynomial quadratic model. ns: non-significant, *: p<0.05, **: p<0.01, ***: p<0.001.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "The current manuscript presents a development of the IPOND technique aimed at capturing the proteome at UVC damage repair sites (IPOND-Repair). This is an interesting new technique that expands the arsenal of methods to study DNA repair in the context of chromatin, and could be further applied to explore the proteome at sites of diverse DNA lesions that require DNA synthesis for their resolution. Using IPOND-R, the authors identify ~250 proteins enriched at UVC damage sites (one-hour post-irradiation). These include several histone chaperones (DNAJC9, MCM2, CAF1, FACT…), nucleosome remodelling enzymes and histone modifiers. The work then focuses on the function of DNAJC9 and MCM2 at UVC damage sites. Chromatin is dynamically remodelled at UVC damage sites, including the transient displacement and recovery of ‘old’ histones, and the deposition of ‘new’ histones. A full picture of which histone chaperones are involved in this remodelling of chromatin is missing, and this work represents an important advance. Here, the authors show that DNAJC9 is required for incorporation of new histones H3 (H3.1 and H3.3 variants) at UVC damage sites (as well as their genome-wide incorporation); and that DNAJC9 and MCM2 similarly function in the recovery of old histone H3.3 levels at UVC repair sites. Together, the work implicates DNAJC9 and MCM2 in the maintenance of chromatin structure at UVC damage sites, without affecting the actual repair of UV lesions.\n\nThe main point that would greatly strengthen the manuscript is a deeper mechanistic understanding of how DNAJC9 and MCM2 function at sites of DNA damage. First, the establishment of a functional rescue system will be key to unequivocally assign the function of DNAJC9 and MCM2 during chromatin remodelling at UVC damage sites. Second, as also pointed out by the authors, based on the current literature a number of mechanisms can be invoked to explain how DNAJC9 and MCM2 accumulate at sites of DNA damage to promote histone incorporation. These could be direct binding to histone H3-H4 (histones evicted from chromatin could become ‘visible’ to these chaperones?); interaction of MCM2 with other CMG components; regulation of ‘damaged’ chromatin binding by the J domain of DNAJC9. Importantly, point mutants that disrupt these interactions have all been described. The contribution of these mechanisms could be tested in a rescue system, and would transform our understanding of how DNAJC9 and MCM2 function at DNA repair sites.\n\nOther points:\nLine 116: Figure S1E only shows XPB recruitment, not ‘UV damage dependent factors’, so the text should be amended accordingly.\nFig. 2D is not very informative, particularly the ‘reference number’ for the gene ontology terms. Can this panel be reshaped?\nHow do the authors explain the difference in proteins retrieved by aniFOUND as compared to IPOND-R (Fig. 3A)? adding a short explanation in the text would be helpful for readers to compare the two techniques. And how is the overlap between IPOND-R and IPOND (untreated conditions)?\nIn Figure 4, the pattern of H3.1 incorporation into chromatin is used to define whether cells are in S phase or not. However, given that loss of DNAJC9 decreases new H3.1-SNAP incorporation and the percentage of S-phase cells (Fig. S5G-I), how reliable is the observed H3.1 pattern? Can the authors test whether DNAJC9-depleted cells still consistently show the expected ‘S-phase’ (early, mid, late) patterns for H3.1-SNAP, p150 and/or EdU? This would validate the use of H3.1-SNAP patterns as a marker of S-phase in Fig. 4.\nFig. 4H: Given that DNACJ9 and HIRA depletion leads to lower H3.3-SNAP incorporation, the H3.3-SNAP images seem not very representative. Has the contrast been changed differently between samples to highlight the decrease in H3.3-SNAP accumulation specifically at UVC sites? if so, this should be stated in the legend.", "reviewer_2": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts.", "reviewer_3": "The paper by Plessier et al, describes a new proteomic method to isolate factors accumulating on newly synthesized DNA after UV DNA damage, named IPOND-R. After validation of the method, the authors focus on analyzing the effect of depletion of DNAJC9 on the dynamics of new and old histones at DNA damage sites. The authors find that DNAJC9 controls the dynamics of both histone populations, but does not affect DNA damage repair per se (i.e. recruitment of repair proteins, or survival after DNA damage). The authors conclude by suggesting an interplay between MCM2 and DNAJC9 to control chromatin dynamics at UV damage sites.\n\nThe paper contains high quality analyses of the effect of DNAJC9 depletion, and also provides a well validated new approach to study the proteome of DNA damage repair sites. In general, the findings are interesting and may attract attention from several research communities involved in DNA repair and chromatin biology. However, there are few concerns that need to be addressed before publication.\n\nThe most critical issue is that it is not clear that the effects deescribed are direct and related to DNA damage repair, rather than indirect effects from the cells going through S-phase without critical histone chaperones. DNAJC9, MCM2 as well as CAF-1 and HIRA are critical histone chaperones and they will alter chromatin dynamics independently of DNA damage, as previously shown. Because the authors do not see an effect of DNAJC9 on DNA repair or survival after damage, it could be that the authors are simply measuring changes in chromatin dynamics that are not dependent on DNA damage, but related to global chromatin alterations. This is due in part to the experimental setup of the assay: the use of siRNA and the conditions used to inhibit DNA replication. Controls should be added to address this concern.\n\n1) The authors run many important controls to validate the IPOND-R approach to minimize the contribution of DNA replication in the dataset. However, treatment of cells with HU and Ara-C to inhibit replicative synthesis also stresses these cells significantly. I wonder if the authors could use synchronization of the cells in G1/S by Palbociclib (which is known to not induce stress) to conclusively demonstrate the power of the method. Running IPOND-R and the follow up (key) experiments in Palbociclib arrested cells will provide important confirmation that DNA replication and S-phase mechanisms are not at play in the chromatin dynamics presented here.\n2) As the authors deplete their proteins with siRNA, the cells are cycling in the absence of the target proteins for at least one full cell cycle. As the authors are targeting critical histone chaperones (DNAJC9, MCM2, CAF-1 and HIRA), it is expected that chromatin is already affected at the time of DNA damage induction. The authors should find ways to circumvent this experimental limitation (i.e. use rapid degron-based degradation or test their key findings in G1 arrested cells).\n3) In line 228-229 it is stated that the quantification of Figure 4B focuses on non S-phase cells. It is not clear to this reviewer how one can reliably distinguish S and non S-phase cells from new H3.1 distribution (I can only imagine a difference between early and late S-phase). I expect a difference could be in the total amount of new H3.1 (should be significantly higher in S-phase cells), but not clear how the distribution can alone explain S vs non-S phase cells. At any rate, the authors should include a quantification for S-phase cells to test if there is a difference. Could the authors use EdU to distinguish various populations otherwise? Or synchronize with Palbociclib as suggested above?\n4) The involvement of MCM2 is also unusual, without evoking the passage of DNA replication forks. The authors do not really mention this, but this needs to be addressed/discussed. How can they envision MCM2 working at these sites, or together with DNAJC9, outside of DNA replication? Hypotheses need to be tested to clarify their data, which is very preliminary in the current form.\n5) In Figure 1D, the authors should probe for DNA replication proteins to show that they are not pulled down (e.g. Cdc45, GINS, or fork protection complex components).\n6) The authors should also show the overlap between IPOND-R and previously reported IPOND hits (in Figure 2), to show how many of the fork proteins are present in the data.\nThere is an instance of “data not shown” in the discussion (lines 339-340). These data should be shown, or the statement needs to be removed.", "reviewer_4": "The study investigates chromatin dynamics during UV damage repair, focusing on identifying proteins that regulate histone deposition at damaged sites. The authors modified the IPOND proteomic method, naming it IPOND-R, to analyze protein dynamics specific to repair sites. Through this approach, they identified DNAJC9 and MCM2, which had not previously been linked to UV-induced damage. DNAJC9 is suggested to facilitate the deposition of new histones and recovery of parental histones, while MCM2 is proposed to collaborate in these processes to maintain chromatin integrity. This finding adds to the understanding of the dynamic regulation of chromatin following DNA damage. However, several issues undermine the study's conclusions. Many results lack appropriate controls, and the experimental descriptions are insufficiently detailed, making the findings difficult to interpret and raising concerns about their reliability. Furthermore, the logical connections between key results are unclear. These shortcomings lead to ambiguous conclusions, and some interpretations appear to extend beyond what the data can support.\n\nMajor:\n1. Based on the proposed model of chromatin repair (Figure 6F), it appears that DNA breaks are repaired first, followed by histone deposition to restore chromatin structure. However, the authors did not assess when DNA repair is complete. Notably, the persistence of γH2AX after 24 hours suggests that repair may not be fully resolved, and DNAJC9 is no longer present at the damage sites at this time point. Furthermore, while DNAJC9 is transiently present at UVC damage sites, the authors report no evidence of its involvement in DNA repair. These findings raise important questions about the proposed model and the role of DNAJC9 at DNA damage sites.\nTo clarify these points and strengthen the model, the authors should provide additional details, including:\n• Evidence indicating when DNA repair is complete.\n• Whether the loss of DNAJC9 affects transcription recovery near break sites following UVC damage.\nAddressing these points will help establish a clearer connection between DNA repair, histone deposition, and the function of DNAJC9 in the context of DNA damage.\n2. The authors treated cells with 10 mM HU and 1 mM AraC to inhibit DNA replication, raising several concerns:\n• While AraC can inhibit DNA replication, it is also incorporated into repair sites, potentially causing repair defects. This issue is not addressed in the manuscript, undermining the validity of the experimental approach.\n• The use of 1 mM AraC is an extremely high concentration for dividing cells, especially since concentrations as low as 40 µM can be toxic to some post-mitotic cells. The observation that cells accumulate in the G1 phase even 24 hours post-treatment with DNA synthesis inhibitors suggests that the inhibitors themselves may have induced significant cellular stress.\nGiven these concerns, we recommend replacing HU+AraC with other inhibitors, such as G1 arrest inhibitors (eg. Pablociclib), which would allow the cells to arrest in G1 phase and prevent EdU incorporation during DNA replication. This adjustment would enhance the reliability of the approach and reduce potential confounding effects.\n3. In Figure 5B, siXPG should prevent UV lesion removal, thereby halting DNA repair. However, the data still show a recovery of old H3.3, which is confusing. If DNA repair is blocked, it would be expected to also impact histone deposition. The authors should clarify how histone deposition occurs independently of DNA repair in this context and provide additional insights into DNAJC9's function.\nMinor:\n1. To identify proteins that transiently bind to chromatin during repair, the authors compared IPOND-R results at 1-hour and 24-hour post UV time points. However, this approach does not rule out the possibility that cells with the most severe damage at 1 hour post-UV may not survive until the 24-hour time point, thus create bias. To address these concerns, the authors should perform EdU pull-down sequencing to confirm that the EdU-incorporated sites at 24 hours are comparable in abundance and distribution to those at 1 hour post-UV treatment.\n2. The analysis in Figures 4 and 5 presents data at 20 minutes, 2 hours, and 8 hours post-UVC exposure without providing any explanation for this shift, despite earlier data focusing on 1 hour post-UVC. This raises questions about whether DNAJC9 remains present at UV damage sites at these time points in the control. To address these issues, the authors should provide a clear rationale for the time-point selection.\n3. The IF results for new H3.3-SNAP in siDNAJC9#3 in Figures 4C and 4H display inconsistent deposition patterns (completely absent in one instance and still present in another), which raises concerns about data reliability. The authors should use figures displaying multiple cells instead of single cells to ensure greater consistency and reliability in the presentation of the data.\n4. There is no description of how the survival assay was performed. It is unclear whether the UVC dose of 10 J/m² causes 85% cell death within 24 hours, 48 hours, or over the course of a week, which creates confusion and raises questions about the experimental conditions.\nIn light of these concerns, the paper's conclusions about DNAJC9's role in chromatin repair appear premature and not fully supported by the data presented. A substantial revision addressing these issues would be necessary to strengthen the study's findings and interpretations.", "reviewer_5": "Plessier et al presents an innovative proteomic approach to capture chromatin in the process of UVC irradiation repair based on the IPOND protocol (IPOND-R). This protocol relies on the complete inhibition of DNA replication such that EdU is only incorporated during UV-damage coupled DNA repair processes. As can be seen in Figure 1B, the inhibition of DNA replication with HU and Ara-C combined is highly effective. Furthermore, the post UVC repair EdU signal is similar in non S-phase cells when not treated with these inhibitors (Figure 1C). These are important controls since small amounts of DNA replication could significantly skew the interpretation of the resulting proteomic analyses from these experiments. Whilst the proteomic datasets have merit, there should be a more detailed comparison and benchmarking of them compared to classical iPOND experiments. For example, by comparing them to the datasets in Sirbu et al JBC VOL. 288, NO. 44, pp. 31458 –3146 (2013). This would help to highlight the differences to factors identified at the replication fork to those enriched at sites of UV damage repair.\nBeyond this, I was not sufficiently convinced by the follow-up studies whereby DNAJC9 was selected as the key factor in this process despite being close to the cut-off for statistical significance in the proteomic datasets. The most convincing aspect of this was that depletion of dNAJC9 causes a loss of new SNAP tagged H3.3/H3.1. In the UV irradiated experiments, the signal of DNAJC9 and MCM2 at irradiated sites was not convincing, no easily observable foci are formed. Furthermore, since this could also be explained by bleed through from the GFP channel into the red channel, and I was not able to find evidence that this has been correctly controlled for, this calls into doubt the validity of conclusions made based on this approach – thus at this stage it is unclear of the significance of many findings. Furthermore, the statistical handling of data is questionable. Whilst microscopy experiments have been performed with three biological replicates, paired t-tests appear to use all data points from these experiments rather than the three means of the biological replicates. This is a very confusing way to present the data and makes its difficult to assess how robust the conclusions made are. For example, are the changes in Figure 5 really strong enough to conclude that DNAJC9 has a role in old histone recovery/histone recycling?\nTogether, whilst I believe this is a promising story, it is not yet sufficiently developed for publication in Nature Comms."}, "author_response": "Point-by-point response to reviewer comments\nWe thank all five referees for insightful and constructive suggestions that proved\nvery useful to improve our manuscript. As main additions to the manuscript:\n we provide additional comparisons of IPOND-R data to IPOND and aniFOUND\n we consolidate the recruitment of DNAJC9 and MCM2 to UV-damaged\nchromatin and assess their function in the UV damage response through\nfunctional rescue experiments\n we demonstrate that the effects observed are replication-independent by using\nalternative experimental approaches including G1-arrest with Palbociclib.\nThe current manuscript presents a development of the IPOND technique aimed at\ncapturing the proteome at UVC damage repair sites (IPOND-Repair). This is an\ninteresting new technique that expands the arsenal of methods to study DNA repair in\nthe context of chromatin, and could be further applied to explore the proteome at sites\nof diverse DNA lesions that require DNA synthesis for their resolution. Using IPOND-\nR, the authors identify ~250 proteins enriched at UVC damage sites (one-hour post-\nirradiation). These include several histone chaperones (DNAJC9, MCM2, CAF1,\nFACT…), nucleosome remodelling enzymes and histone modifiers. The work then\nfocuses on the function of DNAJC9 and MCM2 at UVC damage sites. Chromatin is\ndynamically remodelled at UVC damage sites, including the transient displacement\nand recovery of ‘old’ histones, and the deposition of ‘new’ histones. A full picture of\nwhich histone chaperones are involved in this remodelling of chromatin is missing, and\nthis work represents an important advance. Here, the authors show that DNAJC9 is\nrequired for incorporation of new histones H3 (H3.1 and H3.3 variants) at UVC damage\nsites (as well as their genome-wide incorporation); and that DNAJC9 and MCM2\nsimilarly function in the recovery of old histone H3.3 levels at UVC repair sites.\nTogether, the work implicates DNAJC9 and MCM2 in the maintenance of chromatin\nstructure at UVC damage sites, without affecting the actual repair of UV lesions.\nThe main point that would greatly strengthen the manuscript is a deeper mechanistic\nunderstanding of how DNAJC9 and MCM2 function at sites of DNA damage. First, the\nestablishment of a functional rescue system will be key to unequivocally assign the\nfunction of DNAJC9 and MCM2 during chromatin remodelling at UVC damage sites.\nWe had established a functional rescue system for DNAJC9 (Supplementary Fig. 6d\nnow with 3 experiments instead of 2) and we now provide a rescue experiment also for\nMCM2 showing that re-expression of wild-type MCM2 restores at least partially new\nH3.3 deposition at UV sites (Suppl Fig. 7f).\nSecond, as also pointed out by the authors, based on the current literature a number\nof mechanisms can be invoked to explain how DNAJC9 and MCM2 accumulate at sites\nof DNA damage to promote histone incorporation. These could be direct binding to\nhistone H3-H4 (histones evicted from chromatin could become ‘visible’ to these\nchaperones?); interaction of MCM2 with other CMG components; regulation of\n‘damaged’ chromatin binding by the J domain of DNAJC9. Importantly, point mutants\nthat disrupt these interactions have all been described. The contribution of these\nmechanisms could be tested in a rescue system, and would transform our\nunderstanding of how DNAJC9 and MCM2 function at DNA repair sites.\nThrough functional rescue experiments, we have tested the impact of DNAJC9 histone\nbinding mutant on the recruitment of this chaperone to sites of UV damage repair and\non the resulting repair-coupled histone deposition. We initially planned to test both a\nhistone binding mutant of DNAJC9 and a mutant in the J domain, which abolishes the\ninteraction with Hsp70 family members (PMID: 33857403) but we could not achieve\nstable expression of the J domain mutant. We observed that expression of wild type\nDNAJC9 rescued, at least in part, new H3.3 deposition at UV damage sites in DNAJC9-\nknocked down cells, which was not observed upon expression of a histone binding\nmutant form of DNAJC9 although both wild type and mutant DNAJC9 were similarly\nrecruited to UV damage sites (new Fig. 4d). These experiments establish that\nsynthesized histones at UV damage sites.\nAs for MCM2, the H3-H4 histone binding mutant (Y81A, Y90A) is not a perfect\nseparation of function mutant because all MCM2 histone binding mutants also affect\nthe assembly of the CMG replicative helicase complex to some extent (PMID:\n26167883). Moreover, stable expression of the MCM2 histone binding mutant was\nhighly cytotoxic, preventing functional rescue experiments with this construct. Note that\nwhile the whole MCM hexamer is captured at sites of UV damage repair, other CMG\ncomponents like Cdc45 and GINS are not (Supplementary Fig. 3c), which indicates\nthat MCM2 likely operates independently of the CMG complex at repair sites (now\ndiscussed on p. 17).\nTo further characterize MCM2 and DNAJC9 function in histone dynamics at repair\nsites, we ruled out the hypothesis that MCM2 and DNAJC9 contribution to old H3.3\nrecovery may be an indirect consequence of these chaperones stimulating new H3.3\ndeposition. Indeed, depletion of the H3.3 chaperone HIRA, known to deposit new H3.3\nhistones at sites of UV damage repair (PMID: 24074863), did not impair old H3.3\nrecovery (Supplementary Fig. 7g).\nOther points:\nLine 116: Figure S1E only shows XPB recruitment, not ‘UV damage dependent\nfactors’, so the text should be amended accordingly.\nWe have rephrased this part (p. 6)\nFig. 2D is not very informative, particularly the ‘reference number’ for the gene ontology\nterms. Can this panel be reshaped?\nWe have removed the GO term reference numbers from the representation of the gene\nontology analysis in Fig. 2d and included instead additional broad terms for the most\nenriched GO terms.\nHow do the authors explain the difference in proteins retrieved by aniFOUND as\ncompared to IPOND-R (Fig. 3A)? adding a short explanation in the text would be\nhelpful for readers to compare the two techniques. And how is the overlap between\nIPOND-R and IPOND (untreated conditions)?\nThe modest overlap with the aniFOUND dataset can be explained by the different cell\ntypes (U2OS osteosarcoma cells for IPOND-R vs. 1BR3 skin fibroblasts for\naniFOUND) and most likely by the different time points post UV (1 h for IPOND-R vs.\n4 h for aniFOUND) as early repair responses are not captured 4 h post irradiation (now\nexplained on p. 8).\nWe have also compared the IPOND-R mass spectrometry dataset (1 h-specific hits in\nU2OS cells) to the proteins identified on nascent replicated strands in undamaged\nconditions by IPOND combined with mass spectrometry (593 proteins across 4 human\ncell lines HEK293T, HCT-116, HeLa and RPE-1 (PMID: 31553917) and 522 proteins\nin human TIG-3 cells (PMID: 36680776). This analysis revealed several shared\nreplication-associated factors including RFC1-5, PCNA, MCM2-7 but also highlighted\nimportant differences between the proteomes at replication forks and at sites of UV\ndamage repair with replication-associated factors that are not retrieved at sites of UV\ndamage repair such as Cdc45, Claspin, MCM10, TONSL (Supplementary Fig.3c).\nIn Figure 4, the pattern of H3.1 incorporation into chromatin is used to define whether\n3\ncells are in S phase or not. However, given that loss of DNAJC9 decreases new H3.1-\nSNAP incorporation and the percentage of S-phase cells (Fig. S5G-I), how reliable is\nthe observed H3.1 pattern? Can the authors test whether DNAJC9-depleted cells still\nconsistently show the expected ‘S-phase’ (early, mid, late) patterns for H3.1-SNAP,\np150 and/or EdU? This would validate the use of H3.1-SNAP patterns as a marker of\nS-phase in Fig. 4.\nWe can easily discriminate between cells in and outside S phase based on new H3.1\nnuclear levels even in DNAJC9-knocked down cells because the levels of new H3.1\nare always significantly higher in S phase cells (Supplementary Fig. 6a, S phase cells\nwere identified based on CAF-1 p60, which is unchanged upon DNAJC9 knockdown).\nFig. 4H: Given that DNACJ9 and HIRA depletion leads to lower H3.3-SNAP\nincorporation, the H3.3-SNAP images seem not very representative. Has the contrast\nbeen changed differently between samples to highlight the decrease in H3.3-SNAP\naccumulation specifically at UVC sites? if so, this should be stated in the legend.\nThe image settings are kept the same between samples. We now show more\nrepresentative pictures that better reflect the results of the quantification presented on\nthe graph (Fig. 4i).\nReviewer #2:\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.\nThe paper by Plessier et al, describes a new proteomic method to isolate factors\naccumulating on newly synthesized DNA after UV DNA damage, named IPOND-R.\nAfter validation of the method, the authors focus on analyzing the effect of depletion of\nDNAJC9 on the dynamics of new and old histones at DNA damage sites. The authors\nfind that DNAJC9 controls the dynamics of both histone populations, but does not affect\nDNA damage repair per se (i.e. recruitment of repair proteins, or survival after DNA\ndamage). The authors conclude by suggesting an interplay between MCM2 and\nDNAJC9 to control chromatin dynamics at UV damage sites. The paper contains high\nquality analyses of the effect of DNAJC9 depletion, and also provides a well validated\nnew approach to study the proteome of DNA damage repair sites. In general, the\nfindings are interesting and may attract attention from several research communities\ninvolved in DNA repair and chromatin biology. However, there are few concerns that\nneed to be addressed before publication.\nThe most critical issue is that it is not clear that the effects described are direct and\nrelated to DNA damage repair, rather than indirect effects from the cells going through\nS-phase without critical histone chaperones. DNAJC9, MCM2 as well as CAF-1 and\nHIRA are critical histone chaperones and they will alter chromatin dynamics\nindependently of DNA damage, as previously shown. Because the authors do not see\nan effect of DNAJC9 on DNA repair or survival after damage, it could be that the\nauthors are simply measuring changes in chromatin dynamics that are not dependent\non DNA damage, but related to global chromatin alterations. This is due in part to the\nexperimental setup of the assay: the use of siRNA and the conditions used to inhibit\nDNA replication. Controls should be added to address this concern.\n1) The authors run many important controls to validate the IPOND-R approach to\nminimize the contribution of DNA replication in the dataset. However, treatment of cells\nwith HU and Ara-C to inhibit replicative synthesis also stresses these cells significantly.\nI wonder if the authors could use synchronization of the cells in G1/S by Palbociclib\n(which is known to not induce stress) to conclusively demonstrate the power of the\nmethod. Running IPOND-R and the follow up (key) experiments in Palbociclib arrested\ncells will provide important confirmation that DNA replication and S-phase mechanisms\nare not at play in the chromatin dynamics presented here.\nThe combination of HU and ara-C indeed triggers genotoxic stress. To address this\nvalid concern, we used instead the cdk4/6 inhibitor Palbociclib to arrest cells in G1.\nThis treatment was applied to RPE-1 cells because U2OS cells are resistant due to\nhigh cdk4 expression. We confirmed by EdU staining the complete depletion of\nreplicating cells upon Palbociclib treatment while repair synthesis remained unaffected.\nHowever, we noticed that Palbociclib treatment hindered the UV-induced stress\nresponse as evidenced by a lack of increase of H2A.X signal between non-irradiated\nand UV-irradiated cells, preventing us from running successful IPOND-R experiments\nin Palbociclib-treated cells (no enrichment of material in the +UV capture, see Figure\nfor Reviewerbelow). Nevertheless, we could monitor repair-couped histone dynamics,\nsuch as new H3.3 deposition, in cells arrested in G1 by Palbociclib treatment. Similar\nto what observed in cycling cells, DNAJC9 and MCM2 knock-down in G1-arrested cells\nimpaired new H3.3 deposition at sites of UV damage repair (Supplementary Fig. 7e),\narguing that the function of these chaperones in histone deposition coupled to repair\ndoes not require passage through S phase. Furthermore, DNAJC9 and MCM2\nrecruitment to UV damaged chromatin was observed outside of S phase\n(Supplementary Fig. 4b), indicating that they are replication-independent.\nFigure for Reviewer: Set up of IPOND-R in cells arrested in G1 by Palbociclib treatment\n(a) Inhibition of replication-coupled DNA synthesis measured by EdU incorporation (pulse of 10 min)\nin RPE-1 cells treated for 24 h with 500 nM Palbociclib.\n(b) Effect of Palbociclib pre-treatment on repair-coupled DNA synthesis measured by EdU\nincorporation (pulse of 1 h) at sites of local UV irradiation (marked by CPD immunostaining) in RPE-\n1 cells. S phase cells present in the non-treated population were identified by pan-nuclear EdU staining\nand excluded from the analysis.\n(c) H2A.X levels analyzed by western blot in total extracts from RPE-1 cells treated for 24 h with 500\nnM Palbociclib and exposed or not to 10 J/m2 global UV irradiation. Cells were harvested 1 h post\nirradiation.\n(d) Western blot analysis of proteins captured by IPOND-R 1 h post UV irradiation in RPE-1 cells\narrested in G1 by 24 h treatment with 500 nM Palbociclib. -UV is used as negative control.\nDot plots: mean ± SD. Statistical significance is calculated with a two-sided Student’s t-test with\nWelch’s correction.\n2) As the authors deplete their proteins with siRNA, the cells are cycling in the absence\nof the target proteins for at least one full cell cycle. As the authors are targeting critical\nhistone chaperones (DNAJC9, MCM2, CAF-1 and HIRA), it is expected that chromatin\nis already affected at the time of DNA damage induction. The authors should find ways\nto circumvent this experimental limitation (i.e. use rapid degron-based degradation or\ntest their key findings in G1 arrested cells).\nAs explained above, we validated our findings in G1-arrested RPE-1 cells by\npalbociclib treatment and thus confirmed the contribution of DNAJC9 and MCM2 to\nrepair-coupled histone deposition, independently of cell passage through S phase\n(Supplementary Fig. 7e).\n3) In line 228-229 it is stated that the quantification of Figure 4B focuses on non S-\nphase cells. It is not clear to this reviewer how one can reliably distinguish S and non\nS-phase cells from new H3.1 distribution (I can only imagine a difference between early\nand late S-phase). I expect a difference could be in the total amount of new H3.1\n(should be significantly higher in S-phase cells), but not clear how the distribution can\nalone explain S vs non-S phase cells. At any rate, the authors should include a\nquantification for S-phase cells to test if there is a difference. Could the authors use\nEdU to distinguish various populations otherwise? Or synchronize with Palbociclib as\nsuggested above?\nnuclear levels even in DNAJC9-knocked down cells because the levels of new H3.1\nare always significantly higher in S phase cells (Supplementary Fig. 6a, S phase cells\nwere identified based on CAF-1 p60, which is unchanged upon DNAJC9 knockdown).\n4) The involvement of MCM2 is also unusual, without evoking the passage of DNA\nreplication forks. The authors do not really mention this, but this needs to be\naddressed/discussed. How can they envision MCM2 working at these sites, or\ntogether with DNAJC9, outside of DNA replication? Hypotheses need to be tested to clarify their\ndata, which is very preliminary in the current form.\nMCM2 is enriched in the IPOND-R dataset together with the other MCM subunits\n(MCM3 to 7) showing that the whole MCM hexamer is present at sites of UV damage\nrepair like at replication forks. However, the observed recruitment of MCM2 to sites of\nUV damage repair does not require the passage of replication forks because it is\ndetected outside of S phase (new Supplementary Fig. 4b), highlighting a function of\nMCM2 that is disconnected from DNA replication. In line with this, we do not retrieve\nother components of the CMG replicative helicase complex at repair sites like Cdc45\nand GINS subunits (Supplementary Fig. 3c, Supplementary Table 1), which indicates\nthat MCM2 likely operates independently of the replicative helicase complex at repair\nsites. Notably, an abundant pool of MCM proteins is not engaged with active\nreplisomes in human cells (PMID: 36241664) and those could fulfill replication-\nindependent roles (now discussed on p.17-18).\n5) In Figure 1D, the authors should probe for DNA replication proteins to show that\nthey are not pulled down (e.g. Cdc45, GINS, or fork protection complex components).\nThe comparison between IPOND-R and IPOND data sets (see point 6 below) reveals\nimportant differences between the proteomes at replication forks and at sites of UV\ndamage repair. Some replication-associated proteins are not captured at repair sites\nincluding Claspin, Cdc45, MCM10, TONSL (Supplementary Fig. 3c). We confirmed by\nwestern blot that the replication-associated protein Cdc45 was not enriched in the +UV\ncapture (Supplementary Fig. 3d).\n6) The authors should also show the overlap between IPOND-R and previously\nreported IPOND hits (in Figure 2), to show how many of the fork proteins are present\nin the data.\nWe have compared the IPOND-R mass spectrometry dataset (1 h-specific hits in\nU2OS cells) to the proteins identified on nascent replicated strands in undamaged\nconditions by IPOND-mass spectrometry (593 proteins across 4 human cell lines\nHEK293T, HCT-116, HeLa and RPE-1 (PMID: 31553917) and 522 proteins in human\nTIG-3 cells (PMID: 36680776). This analysis revealed several shared replication-\nassociated factors including RFC1-5, PCNA, MCM2-7 but also highlighted important\ndifferences between the proteomes at replication forks and at sites of UV damage\nrepair with replication-associated factors that are not retrieved at sites of UV damage\nrepair such as Claspin, Cdc45, MCM10, TONSL (Supplementary Fig.3c)\nThere is an instance of “data not shown” in the discussion (lines 339-340). These data\nshould be shown, or the statement needs to be removed.\nWe now show that DNAJC9 is loosely bound to UV-damaged chromatin, as it is mostly\nsolubilized by detergents, in contrast to CAF-1 and HIRA (Supplementary Fig. 6n).\nThe study investigates chromatin dynamics during UV damage repair, focusing on\nidentifying proteins that regulate histone deposition at damaged sites. The authors\nmodified the IPOND proteomic method, naming it IPOND-R, to analyze protein\ndynamics specific to repair sites. Through this approach, they identified DNAJC9 and\nMCM2, which had not previously been linked to UV-induced damage. DNAJC9 is\nsuggested to facilitate the deposition of new histones and recovery of parental\nhistones, while MCM2 is proposed to collaborate in these processes to maintain\nchromatin integrity. This finding adds to the understanding of the dynamic regulation\nof chromatin following DNA damage. However, several issues undermine the study's\nconclusions. Many results lack appropriate controls, and the experimental descriptions\nare insufficiently detailed, making the findings difficult to interpret and raising concerns\nabout their reliability. Furthermore, the logical connections between key results are\nunclear. These shortcomings lead to ambiguous conclusions, and some interpretations\nappear to extend beyond what the data can support.\nMajor:\n1. Based on the proposed model of chromatin repair (Figure 6F), it appears that DNA\nbreaks are repaired first, followed by histone deposition to restore chromatin structure.\nHowever, the authors did not assess when DNA repair is complete. Notably, the\npersistence of γH2AX after 24 hours suggests that repair may not be fully resolved,\nand DNAJC9 is no longer present at the damage sites at this time point. Furthermore,\nwhile DNAJC9 is transiently present at UVC damage sites, the authors report no\nevidence of its involvement in DNA repair. These findings raise important questions\nabout the proposed model and the role of DNAJC9 at DNA damage sites.\nTo clarify these points and strengthen the model, the authors should provide additional\ndetails, including:\n• Evidence indicating when DNA repair is complete.\n• Whether the loss of DNAJC9 affects transcription recovery near break sites following\nUVC damage.\nAddressing these points will help establish a clearer connection between DNA repair,\nhistone deposition, and the function of DNAJC9 in the context of DNA damage.\nSome histone deposition arises before DNA repair completion as shown for new H3.3\ndeposition, which is coupled to early repair steps, namely UV damage detection (PMID:\n24074863). To better illustrate this point, we now indicate on the model in Fig. 6f “New\nhistone deposition coupled to DNA repair”. Therefore, the presence of a histone\nchaperone on UV-damaged chromatin before repair completion does not preclude an\ninvolvement of this chaperone in chromatin restoration coupled to DNA repair.\nEvidence for the completion of UV damage repair around 24 h post UV is two-fold: (1)\nThe kinetics of UV lesion removal (Supplementary Fig. 1f-g) shows that 6,4-\nphotoproducts are repaired fast (within 2 h) and the CPDs are removed in 24-30 h. (2)\nUV damage repair factors are no longer captured by IPOND-R 24 h post UV indicating\nthat most repair is complete by that time. The persistence of some H2A.X signal 24 h\npost UV (Fig. 3d) is not a direct readout of the presence of DNA damage. It shows that\nDNA damage signaling is not yet turned off, which occurs after repair completion.\nTo further characterize the function of DNAJC9 in UV-damaged chromatin, we\ninvestigated the contribution of this histone chaperone to transcription recovery\nfollowing UVC damage by labeling nascent transcripts with ethynyl-uridine. Knocking\n8\ndown DNAJC9 did not impair transcription recovery at UV damage sites in contrast to\nknocking down the transcription-coupled repair factor CSB (Supplementary Fig. 5f).\nThese results are consistent with a previous multiomic analysis of the UV damage\nresponse (PMID: 27184836) where DNAJC9 did not stand out as a factor regulating\nnascent transcription post UV, and rule out a function of DNAJC9 in transcription-\ncoupled repair.\n2. The authors treated cells with 10 mM HU and 1 mM AraC to inhibit DNA replication,\nraising several concerns:\n• While AraC can inhibit DNA replication, it is also incorporated into repair sites,\npotentially causing repair defects. This issue is not addressed in the manuscript,\nundermining the validity of the experimental approach.\n• The use of 1 mM AraC is an extremely high concentration for dividing cells, especially\nsince concentrations as low as 40 µM can be toxic to some post-mitotic cells. The\nobservation that cells accumulate in the G1 phase even 24 hours post-treatment with\nDNA synthesis inhibitors suggests that the inhibitors themselves may have induced\nsignificant cellular stress.\nGiven these concerns, we recommend replacing HU+AraC with other inhibitors, such\nas G1 arrest inhibitors (eg. Pablociclib), which would allow the cells to arrest in G1\nphase and prevent EdU incorporation during DNA replication. This adjustment would\nenhance the reliability of the approach and reduce potential confounding effects.\nAlthough we verified that the combination HU+Ara-C did not cause UV damage repair\ndefects (Supplementary Fig. 1e-g), we acknowledge that such treatment can trigger\ngenotoxic stress (Supplementary Fig. 1a-c). To address this valid concern, also raised\nby Reviewer #3, we used the cdk4/6 inhibitor Palbociclib to arrest cells in G1. This\ntreatment was applied to RPE-1 cells because U2OS cells are resistant due to high\ncdk4 expression. We confirmed by EdU staining the complete depletion of replicating\ncells upon Palbociclib treatment while repair synthesis remained unaffected. However,\nwe noticed that Palbociclib treatment hindered the UV-induced stress response, as\nevidenced a lack of increase of H2A.X signal between non-irradiated and UV-\nirradiated cells, preventing us from running successful IPOND-R experiments in\nPalbociclib-treated cells (no enrichment of material in the +UV capture, see Figure for\nReviewer below). Nevertheless, we could monitor repair-couped histone dynamics,\nsuch as new H3.3 deposition, in cells arrested in G1 by Palbociclib treatment. Similar\nto what observed in cycling cells, DNAJC9 and MCM2 knock-down in G1-arrested cells\nimpaired new H3.3 deposition at sites of UV damage repair (Supplementary Fig. 7e),\narguing that the function of these chaperones in histone deposition coupled to repair\ndoes not require passage through S phase.\n(a) Inhibition of replication-coupled DNA synthesis measured by EdU incorporation (pulse of 10 min)\nin RPE-1 cells treated for 24 h with 500 nM Palbociclib.\n(b) Effect of Palbociclib pre-treatment on repair-coupled DNA synthesis measured by EdU\nincorporation (pulse of 1 h) at sites of local UV irradiation (marked by CPD immunostaining) in RPE-\n1 cells. S phase cells present in the non-treated population were identified by pan-nuclear EdU staining\nand excluded from the analysis.\n(c) H2A.X levels analyzed by western blot in total extracts from RPE-1 cells treated for 24 h with 500\nnM Palbociclib and exposed or not to 10 J/m2 global UV irradiation. Cells were harvested 1 h post\nirradiation.\n(d) Western blot analysis of proteins captured by IPOND-R 1 h post UV irradiation in RPE-1 cells\narrested in G1 by 24 h treatment with 500 nM Palbociclib. -UV is used as negative control.\nDot plots: mean ± SD. Statistical significance is calculated with a two-sided Student’s t-test with\nWelch’s correction.\n3. In Figure 5B, siXPG should prevent UV lesion removal, thereby halting DNA repair.\nHowever, the data still show a recovery of old H3.3, which is confusing. If DNA repair\nis blocked, it would be expected to also impact histone deposition. The authors should\nclarify how histone deposition occurs independently of DNA repair in this context and\nprovide additional insights into DNAJC9's function.\nIn Fig. 5b, siXPG is indeed used as a positive control because XPG depletion prevents\nUV lesion removal thus impacting old histone recovery. XPG knock-down still allows\nsome old histone recovery because the siRNA-mediated knock-down does not fully\nremove the protein thus repair is not 100% abrogated (as now stated on p. 13-14) but\nold histone recovery is significantly impaired in siXPG compared to the siLuc control\ncondition (p<0.0001).\nMinor:\n1. To identify proteins that transiently bind to chromatin during repair, the authors\ncompared IPOND-R results at 1-hour and 24-hour post UV time points. However, this\napproach does not rule out the possibility that cells with the most severe damage at 1\nhour post-UV may not survive until the 24-hour time point, thus create bias. To address\nthese concerns, the authors should perform EdU pull-down sequencing to confirm that\nthe EdU-incorporated sites at 24 hours are comparable in abundance and distribution\nto those at 1 hour post-UV treatment.\nWe show that the levels of incorporated EdU in cell nuclei are not lower at 24 h\ncompared to 1 h post UV (Supplementary Fig. 2b), arguing against the death of UV-\ndamaged, EdU-labeled cells between 1 h and 24 h post irradiation (now stated on p.6).\nshown by CPD staining immediately after global UVC irradiation (new Supplementary\nFig. 2c) so there is not a large proportion of cells with more severe damage and it does\nnot introduce any bias in the analysis.\n2. The analysis in Figures 4 and 5 presents data at 20 minutes, 2 hours, and 8 hours\npost-UVC exposure without providing any explanation for this shift, despite earlier data\nfocusing on 1 hour post-UVC. This raises questions about whether DNAJC9 remains\npresent at UV damage sites at these time points in the control. To address these\nissues, the authors should provide a clear rationale for the time-point selection.\nTwenty minutes post UV was previously determined as the best time point to visualize\nHIRA accumulation at UV sites, 2 h is the best time point to visualize CAF-1 and new\nH3 deposition at repair sites (PMID: 24074863) and 8 h is suited to visualize old histone\nrecovery (PMID: 27642047). In addition to the 1 h and 8 h time points (Fig. 3e-f and\n6a), we now also show DNAJC9 and MCM2 enrichment at sites of UV damage repair\n20 min and 2 h following local UVC irradiation (Supplementary Fig. 4a).\n3. The IF results for new H3.3-SNAP in siDNAJC9#3 in Figures 4C and 4H display\ninconsistent deposition patterns (completely absent in one instance and still present in\nanother), which raises concerns about data reliability. The authors should use figures\ndisplaying multiple cells instead of single cells to ensure greater consistency and\nreliability in the presentation of the data.\nWe now show more representative pictures in Fig. 4i that better reflect the results of\nthe quantification presented on the graph. We believe that quantifying the signal in at\nleast 150 cells scored in 3 independent experiments is the most reliable way of\npresenting the data.\n4. There is no description of how the survival assay was performed. It is unclear\nwhether the UVC dose of 10 J/m² causes 85% cell death within 24 hours, 48 hours, or\nover the course of a week, which creates confusion and raises questions about the\nexperimental conditions.\nThe description of the survival assay was present in the method section under the\ncaption “Colony forming assays”, which we have now rephrased as “Cell survival\nassays” for clarity. In these assays, cell colonies are counted after 12 days of growth\nfollowing UV irradiation. As previously stated, there is hardly any detectable cell death\nwithin 24 h post UV.\nIn light of these concerns, the paper's conclusions about DNAJC9's role in chromatin\nrepair appear premature and not fully supported by the data presented. A substantial\nrevision addressing these issues would be necessary to strengthen the study's findings\nand interpretations.\nPlessier et al presents an innovative proteomic approach to capture chromatin in the\nprocess of UVC irradiation repair based on the IPOND protocol (IPOND-R). This\nprotocol relies on the complete inhibition of DNA replication such that EdU is only\nincorporated during UV-damage coupled DNA repair processes. As can be seen in\nFigure 1B, the inhibition of DNA replication with HU and Ara-C combined is highly\neffective. Furthermore, the post UVC repair EdU signal is similar in non S-phase cells\nwhen not treated with these inhibitors (Figure 1C). These are important controls since\nsmall amounts of DNA replication could significantly skew the interpretation of the\nresulting proteomic analyses from these experiments. Whilst the proteomic datasets\nhave merit, there should be a more detailed comparison and benchmarking of them\ncompared to classical iPOND experiments. For example, by comparing them to the\ndatasets in Sirbu et al JBC VOL. 288, NO. 44, pp. 31458 –3146 (2013). This would\nhelp to highlight the differences to factors identified at the replication fork to those\nenriched at sites of UV damage repair.\nAs suggested, we have compared the IPOND-R mass spectrometry dataset (1 h-\nspecific hits in U2OS cells) to the proteins identified on nascent replicated strands in\nundamaged conditions by IPOND-mass spectrometry (593 proteins across 4 human\ncell lines HEK293T, HCT-116, HeLa and RPE-1 (PMID: 31553917) and 522 proteins\nin human TIG-3 cells (PMID: 36680776). This analysis revealed several shared\nreplication-associated factors including RFC1-5, PCNA, MCM2-7) but also highlighted\nimportant differences between the proteomes at replication forks and at sites of UV\ndamage repair with replication-associated factors that are not retrieved at sites of UV\ndamage repair such as Cdc45, Claspin, MCM10, TONSL (Supplementary Fig.3c).\nBeyond this, I was not sufficiently convinced by the follow-up studies whereby DNAJC9\nwas selected as the key factor in this process despite being close to the cut-off for\nstatistical significance in the proteomic datasets. The most convincing aspect of this\nwas that depletion of DNAJC9 causes a loss of new SNAP tagged H3.3/H3.1. In the\nUV irradiated experiments, the signal of DNAJC9 and MCM2 at irradiated sites was\nnot convincing, no easily observable foci are formed. Furthermore, since this could\nalso be explained by bleed through from the GFP channel into the red channel, and I\nwas not able to find evidence that this has been correctly controlled for, this calls into\ndoubt the validity of conclusions made based on this approach – thus at this stage it is\nunclear of the significance of many findings.\nDNAJC9 was chosen with MCM2 because those were the only two histone chaperones\nin the IPOND-R capture that had not previously been involved in the UV damage\nresponse. Even if close to the cut-off for statistical significance, DNAJC9 recruitment\nwas confirmed at UV sites by imaging approaches (Fig. 3e-f). We acknowledge that\nthe observed enrichment of DNAJC9 at UV sites is rather mild and this is likely due to\nthe fact that DNAJC9 is loosely bound to chromatin contrary to CAF-1 and HIRA\nchaperones (Supplementary Fig. 6n). MCM2 recruitment is more readily detectable at\nlate time points post irradiation (Fig. 6a) and we verify that the observed histone\nchaperone signal enrichment at UV damage sites does not result from a bleed-through\nfrom the UV damage channel (new Supplementary Fig. 7a). To consolidate our\nfindings, we examine more thoroughly the recruitment of DNAJC9 and MCM2 to sites\nof UV damage repair at different time points following damage infliction (Fig. 3e-f,\nSupplementary Fig. 4a, Fig. 6a-b) and we demonstrate that their recruitment to UV\nsites occurs outside of S phase (Supplementary Fig. 4b). DNAJC9 recruitment to UV\n12\ndamage sites 8h post irradiation was not very convincing and we thus decided to\nremove the data from the manuscript.\nFunctionally, our follow-up studies demonstrate that DNAJC9 affects histone H3\ndynamics at repair sites, which we confirmed and further characterized through\nfunctional rescue experiments (Fig. 4d). These experiments establish that DNAJC9,\nthrough its histone binding domain, promotes the deposition of newly synthesized H3\nvariants at UV damage sites. In addition, we have run additional control experiments\nshowing that the function of DNAJC9 in new histone deposition at repair sites does not\nrequire the passage of cells through S phase (Supplementary Fig. 7e).\nFurthermore, the statistical handling of data is questionable. Whilst microscopy\nexperiments have been performed with three biological replicates, paired t-tests\nappear to use all data points from these experiments rather than the three means of\nthe biological replicates. This is a very confusing way to present the data and makes it\ndifficult to assess how robust the conclusions made are. For example, are the changes\nin Figure 5 really strong enough to conclude that DNAJC9 has a role in old histone\nrecovery/histone recycling?\nTogether, whilst I believe this is a promising story, it is not yet sufficiently developed\nfor publication in Nature Comms.\nWe run the statistical analyses on all data points and not on the means of the biological\nreplicates in conditions of local UV irradiation because in this case each nucleus\nreceives a different UV dose, which depends on the pore distribution in the micropore\nfilters used for local UVC irradiation (mostly 1 or 2 and up to 5 UV spots per nucleus,\nPMID: 39820273), so each nucleus should be considered independently. This is now\nstated in the method section (p. 35).\nEven if modest, the defects observed upon knockdown of DNAJC9 are comparable to\nthe defects observed upon depletion of the HIRA and CAF-1 chaperones for new\nhistone deposition (Fig. 4h-i) and comparable to the defects observed upon depletion\nof the late repair factor XPG for old histone recovery (Fig. 5b). Furthermore, the\nimpaired recovery of old histones observed upon DNAJC9 depletion cannot be\nattributed to defective UV damage repair because DNAJC9-knocked down cells are\nfully proficient for the repair of UV lesions (Supplementary Fig. 5). This is now\nexplained on p. 14 of our revised manuscript.\nTo further strengthen the role of DNAJC9 and MCM2 in old histone recovery, we ruled\nout the hypothesis that MCM2 and DNAJC9 contribution to old H3.3 recovery may be\nan indirect consequence of these chaperones stimulating new H3.3 deposition. Indeed,\ndepletion of the H3.3 chaperone HIRA, known to deposit new H3.3 histones at sites of\nUV damage repair (PMID: 24074863), does not impair old H3.3 recovery\n(Supplementary Fig. 7g).\n13"}, {"round": 2, "reviewer_comments": {"reviewer_1": "The authors have answered most of the questions we raised. A couple of minor points need to be clarified in the text.\nFig. 4D: in the H3.3-SNAP panel, what are the authors measuring? Is it the total levels of H3.3-SNAP expression? In siLuc or siDNAJC9 cells (and if it is in the latter, how does that fit with DNAJC9 leading to overall decreased histone incorporation?)\nOn page 13, the authors state that ‘DNAJC9 likely provides new H3-H4 histones to CAF-1 and HIRA chaperones because DNAJC9 is loosely bound to UV-damaged chromatin, as it is mostly solubilized by detergents’. While I agree that DNAJC9 likely acts upstream of HIRA and CAF1, I think that this cannot be inferred by a detergent extraction method. There is literature suggesting this, and I think that this sentence should be rephrased accordingly.", "reviewer_2": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts.", "reviewer_3": "The authors have done a good job at addressing my concerns, it's unfortunate that they could not perform IPOND-R after treatment with Palbociclib due to technical reasons. Nonetheless, the manuscript has significantly improved and now provides important contributions to the field, that should be published.", "reviewer_4": "The authors have adequately addressed most comments from the reviewers. I'm still somewhat concerned about whether the histone chaperones at sites of DNA repair at UV lesions is related to stalled replication forks. However, the authors tried to address this concern and the results are consistent with their hypothesis. Thus, I do recommend publication.", "reviewer_5": "Reviewer #5:\n• Please see comments below in red. Some are major concerns which are easily addressed, yet authors have not attempted to address them. Specifically, my main concerns are surrounding the significance of the data presented in Figure 5. I have detailed steps to address my concerns below, and these should be addressed prior to acceptance for publication.\n• In addition to the steps below, I noted Sandra Piquet is missing from the author list in the manuscript file.\nPlessier et al presents an innovative proteomic approach to capture chromatin in the process of UVC irradiation repair based on the IPOND protocol (IPOND-R). This protocol relies on the complete inhibition of DNA replication such that EdU is only incorporated during UV-damage coupled DNA repair processes. As can be seen in Figure 1B, the inhibition of DNA replication with HU and Ara-C combined is highly effective. Furthermore, the post UVC repair EdU signal is similar in non S-phase cells when not treated with these inhibitors (Figure 1C). These are important controls since small amounts of DNA replication could significantly skew the interpretation of the resulting proteomic analyses from these experiments. Whilst the proteomic datasets have merit, there should be a more detailed comparison and benchmarking of them compared to classical iPOND experiments. For example, by comparing them to the datasets in Sirbu et al JBC VOL. 288, NO. 44, pp. 31458 –3146 (2013). This would help to highlight the differences to factors identified at the replication fork to those enriched at sites of UV damage repair.\nAs suggested, we have compared the IPOND-R mass spectrometry dataset (1 h-specific hits in U2OS cells) to the proteins identified on nascent replicated strands in undamaged conditions by IPOND-mass spectrometry (593 proteins across 4 human cell lines HEK293T, HCT-116, HeLa and RPE-1 (PMID: 31553917) and 522 proteins in human TIG-3 cells (PMID: 36680776). This analysis revealed several shared replication-associated factors including RFC1-5, PCNA, MCM2-7) but also highlighted important differences between the proteomes at replication forks and at sites of UV damage repair with replication-associated factors that are not retrieved at sites of UV damage repair such as Cdc45, Claspin, MCM10, TONSL (Supplementary Fig.3c).\n• Thank you for including this analysis in the revised manuscript. Please include the source data for the Venn Diagram in the supplemental excel file since it would be useful for the field to have the lists of proteins identified in each section of the Venn Diagrams depicted in Supplementary Figure 3b-c.\nBeyond this, I was not sufficiently convinced by the follow-up studies whereby DNAJC9 was selected as the key factor in this process despite being close to the cut-off for statistical significance in the proteomic datasets. The most convincing aspect of this was that depletion of DNAJC9 causes a loss of new SNAP tagged H3.3/H3.1. In the UV irradiated experiments, the signal of DNAJC9 and MCM2 at irradiated sites was not convincing, no easily observable foci are formed. Furthermore, since this could also be explained by bleed through from the GFP channel into the red channel, and I was not able to find evidence that this has been correctly controlled for, this calls into doubt the validity of conclusions made based on this approach – thus at this stage it is unclear of the significance of many findings.\nDNAJC9 was chosen with MCM2 because those were the only two histone chaperones in the IPOND-R capture that had not previously been involved in the UV damage response. Even if close to the cut-off for statistical significance, DNAJC9 recruitment was confirmed at UV sites by imaging approaches (Fig. 3e-f). We acknowledge that the observed enrichment of DNAJC9 at UV sites is rather mild and this is likely due to the fact that DNAJC9 is loosely bound to chromatin contrary to CAF-1 and HIRA chaperones (Supplementary Fig. 6n). MCM2 recruitment is more readily detectable at late time points post irradiation (Fig. 6a) and we verify that the observed histone chaperone signal enrichment at UV damage sites does not result from a bleed-through from the UV damage channel (new Supplementary Fig. 7a). To consolidate our findings, we examine more thoroughly the recruitment of DNAJC9 and MCM2 to sites of UV damage repair at different time points following damage infliction (Fig. 3e-f, Supplementary Fig. 4a, Fig. 6a-b) and we demonstrate that their recruitment to UV sites occurs outside of S phase (Supplementary Fig. 4b). DNAJC9 recruitment to UV damage sites 8h post irradiation was not very convincing and we thus decided to remove the data from the manuscript.\nFunctionally, our follow-up studies demonstrate that DNAJC9 affects histone H3 dynamics at repair sites, which we confirmed and further characterized through functional rescue experiments (Fig. 4d). These experiments establish that DNAJC9, through its histone binding domain, promotes the deposition of newly synthesized H3 variants at UV damage sites. In addition, we have run additional control experiments showing that the function of DNAJC9 in new histone deposition at repair sites does not require the passage of cells through S phase (Supplementary Fig. 7e).\n• The data in figure 4d shows quite clearly that exogenous DNAJC9 expression does rescue new H3.3 levels at UVC DNA repair sites and this is not observed in the 4A histone binding mutant. This is an excellent addition to the manscript.\nFurthermore, the statistical handling of data is questionable. Whilst microscopy experiments have been performed with three biological replicates, paired t-tests appear to use all data points from these experiments rather than the three means of the biological replicates. This is a very confusing way to present the data and makes it difficult to assess how robust the conclusions made are. For example, are the changes in Figure 5 really strong enough to conclude that DNAJC9 has a role in old histone recovery/histone recycling?\nTogether, whilst I believe this is a promising story, it is not yet sufficiently developed for publication in Nature Comms.\nWe run the statistical analyses on all data points and not on the means of the biological replicates in conditions of local UV irradiation because in this case each nucleus receives a different UV dose, which depends on the pore distribution in the micropore filters used for local UVC irradiation (mostly 1 or 2 and up to 5 UV spots per nucleus, PMID: 39820273), so each nucleus should be considered independently. This is now stated in the method section (p. 35).\n• On page 6 the authors state that “Damage induction is quite homogenous in a cell population (Supplementary Fig. 2c) so this does not introduce any bias in subsequent analyses.” This is at odds with the statistical handling of the data in Figure 5 and 6, where all data points across three biological replicates are used instead of the three biological replicate means during statistical testing. In addition, since the above critique would have been an easy point to rebut by simply recalculating P values based on the three biological replicate means, does this mean that maintaining the current statistical representation is the only way to maintain the significance of this observation?\n• Figure 4d should have indication of the UVC treatment in the figure (time post local UVC), similar to 4b.\nEven if modest, the defects observed upon knockdown of DNAJC9 are comparable to the defects observed upon depletion of the HIRA and CAF-1 chaperones for new histone deposition (Fig. 4h-i) and comparable to the defects observed upon depletion of the late repair factor XPG for old histone recovery (Fig. 5b). Furthermore, the impaired recovery of old histones observed upon DNAJC9 depletion cannot be attributed to defective UV damage repair because DNAJC9-knocked down cells are fully proficient for the repair of UV lesions (Supplementary Fig. 5). This is now explained on p. 14 of our revised manuscript.\n• In addition, why does siLuc in Figure 5b promote over-recovery of the TMR mean intensity at UVC sites/nucleus (the mean is >1)? In contrast with siDNAJC9 and siXPG the TMR mean intensity at UVC sites/nucleus is approx. =1, wouldn’t this be consistent with full signal recovery? Therefore, wouldn’t this data argue that DNAJC9 and XPG have no role in old histone recovery after UV damage. I therefore question the interpretation of the data here, and question whether this analysis is actually worthy of inclusion in the publication. I would consider the analysis in Supplementary Figure 7f as more pertinent to the story, this panel demonstrates the role of MCM2 in new H3.3 deposition at UVC sites, a novel discovery that should be given more prominence.\nTo further strengthen the role of DNAJC9 and MCM2 in old histone recovery, we ruled out the hypothesis that MCM2 and DNAJC9 contribution to old H3.3 recovery may be an indirect consequence of these chaperones stimulating new H3.3 deposition. Indeed, depletion of the H3.3 chaperone HIRA, known to deposit new H3.3 histones at sites of UV damage repair (PMID: 24074863), does not impair old H3.3 recovery (Supplementary Fig. 7g).\n• This is an independent experiment, as such an experiment that included siRNA depletion of DNAJC9 and MCM2 would be more convincing. E.g. in the same experiment comparing.\n1) siControl\n2) siDNAJC9\n3) siMCM2\n4) siHIRA\n5) siDNAJC9+siMCM2\n6) siDNAJC9+siHIRA\n7) siMCM2+siHIRA\n• One final point is that the conclusion that siDNAJC9 affects the proportion of cells in S phase (Supplementary figure 6h) is based on the proportion of cells that are positive for the histone chaperone Caf1. Can this result be repeated with another more classical readout? E.g. FAC analysis, or using microscopy read-outs e.g. BrdU or EdU?"}, "author_response": "Point-by-point response to reviewer comments\nWe thank all referees for a careful analysis of our manuscript and for their additional\nsuggestions, which we fully address below.\nReviewer #1:\nThe authors have answered most of the questions we raised. A couple of minor points\nneed to be clarified in the text.\nFig. 4D: in the H3.3-SNAP panel, what are the authors measuring? Is it the total levels\nof H3.3-SNAP expression? In siLuc or siDNAJC9 cells (and if it is in the latter, how\ndoes that fit with DNAJC9 leading to overall decreased histone incorporation?)\nWe are measuring the total levels of H3.3-SNAP in nuclei in the absence of DNAJC9\nknockdown. This is now indicated in the corresponding figure legend (p. 45).\nOn page 13, the authors state that ‘DNAJC9 likely provides new H3-H4 histones to\nCAF-1 and HIRA chaperones because DNAJC9 is loosely bound to UV-damaged\nchromatin, as it is mostly solubilized by detergents’. While I agree that DNAJC9 likely\nacts upstream of HIRA and CAF1, I think that this cannot be inferred by a detergent\nextraction method. There is literature suggesting this, and I think that this sentence\nshould be rephrased accordingly.\nWe have rephrased this sentence not to infer DNAJC9 upstream function in the histone\nsupply chain solely based on detergent extraction (p.13).\nReviewer #2:\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.\nReviewer #3:\nThe authors have done a good job at addressing my concerns, it's unfortunate that\nthey could not perform IPOND-R after treatment with Palbociclib due to technical\nreasons. Nonetheless, the manuscript has significantly improved and now provides\nimportant contributions to the field, that should be published.\nReviewer #4:\nThe authors have adequately addressed most comments from the reviewers. I'm still\nsomewhat concerned about whether the histone chaperones at sites of DNA repair at\nUV lesions is related to stalled replication forks. However, the authors tried to address\nthis concern and the results are consistent with their hypothesis. Thus, I do recommend\npublication.\n• Please see comments below in red. Some are major concerns which are easily\naddressed, yet authors have not attempted to address them. Specifically, my main\nconcerns are surrounding the significance of the data presented in Figure 5. I have\ndetailed steps to address my concerns below, and these should be addressed prior\nto acceptance for publication.\n• In addition to the steps below, I noted Sandra Piquet is missing from the author list in\nthe manuscript file.\nSandra Piquet is not listed as an author because she did not perform experiments in\nthis study. She is leading the microscopy facility and is therefore listed in the\nacknowledgement section.\nAs suggested, we have compared the IPOND-R mass spectrometry dataset (1 h-\nspecific hits in U2OS cells) to the proteins identified on nascent replicated strands in\nundamaged conditions by IPOND-mass spectrometry (593 proteins across 4 human\ncell lines HEK293T, HCT-116, HeLa and RPE-1 (PMID: 31553917) and 522 proteins\nin human TIG-3 cells (PMID: 36680776). This analysis revealed several shared\nreplication-associated factors including RFC1-5, PCNA, MCM2-7) but also highlighted\nimportant differences between the proteomes at replication forks and at sites of UV\ndamage repair with replication-associated factors that are not retrieved at sites of UV\ndamage repair such as Cdc45, Claspin, MCM10, TONSL (Supplementary Fig.3c).\n• Thank you for including this analysis in the revised manuscript. Please include the\nsource data for the Venn Diagram in the supplemental excel file since it would be useful\nfor the field to have the lists of proteins identified in each section of the Venn Diagrams\ndepicted in Supplementary Figure 3b-c.\nWe have included this data in Supplementary Table 2 (p.8).\n• The data in figure 4d shows quite clearly that exogenous DNAJC9 expression does\nrescue new H3.3 levels at UVC DNA repair sites and this is not observed in the 4A\nhistone binding mutant. This is an excellent addition to the manuscript.\nWe thank the reviewer for underlining this.\n• On page 6 the authors state that “Damage induction is quite homogenous in a cell\npopulation (Supplementary Fig. 2c) so this does not introduce any bias in subsequent\nanalyses.” This is at odds with the statistical handling of the data in Figure 5 and 6,\nwhere all data points across three biological replicates are used instead of the three\nbiological replicate means during statistical testing. In addition, since the above critique\nwould have been an easy point to rebut by simply recalculating P values based on the\nthree biological replicate means, does this mean that maintaining the current statistical\nrepresentation is the only way to maintain the significance of this observation?\nA distinction needs to be made between global UV irradiation, which leads to\nhomogeneous damage induction in the cell population (Supplementary Fig. 2c), and\nlocal UV irradiation (performed in Figures 5 and 6), where each nucleus receives a\ndifferent UV dose, which depends on the pore distribution in the micropore filters.\nThe statistical handling of the data is thus different between global and local UV irradiation\n2\nexperiments. We now more clearly explain in the method section how we run the\nstatistical analyses of the data in each case (p.35).\n• Figure 4d should have indication of the UVC treatment in the figure (time post local\nUVC), similar to 4b.\nThis has been added (Fig. 4d).\n• In addition, why does siLuc in Figure 5b promote over-recovery of the TMR mean\nintensity at UVC sites/nucleus (the mean is >1)? In contrast with siDNAJC9 and siXPG\nthe TMR mean intensity at UVC sites/nucleus is approx. =1, wouldn’t this be consistent\nwith full signal recovery? Therefore, wouldn’t this data argue that DNAJC9 and XPG\nhave no role in old histone recovery after UV damage. I therefore question the\ninterpretation of the data here, and question whether this analysis is actually worthy of\ninclusion in the publication. I would consider the analysis in Supplementary Figure 7f\nas more pertinent to the story, this panel demonstrates the role of MCM2 in new H3.3\ndeposition at UVC sites, a novel discovery that should be given more prominence.\nOld histone recovery ideally is measured through live cell imaging post UVC laser\ndamage by comparing old histone levels in the UV-damaged region at a late time point\npost irradiation to the condition before UV irradiation (PMID: 27642047). In this context,\nfull recovery would give a value of 1 but this live imaging method is not feasible with\nmultiple siRNAs. To measure old histone recovery in fixed cells instead, we used as a\nproxy the ratio of old histones at UV damage sites relative to the nucleus at an early\n(10 min) and a late time point (8 h) post UV irradiation. The ratio of old histones at UV\ndamage sites relative to the nucleus 8 h post UV in siLuc cells sets the level of recovery\nof old histones in control condition (even if slightly above 1). Comparatively, old histone\nrecovery is diminished in siXPG and siDNAJC9 conditions. This is now better explained\nin our revised manuscript (p.13).\nWe have also emphasized the results shown in Supplementary Fig. 7f in the main text\n(p.16).\nTo further strengthen the role of DNAJC9 and MCM2 in old histone recovery, we ruled\nout the hypothesis that MCM2 and DNAJC9 contribution to old H3.3 recovery may be\nan indirect consequence of these chaperones stimulating new H3.3 deposition. Indeed,\ndepletion of the H3.3 chaperone HIRA, known to deposit new H3.3 histones at sites of\nUV damage repair (PMID: 24074863), does not impair old H3.3 recovery\n(Supplementary Fig. 7g).\n• This is an independent experiment, as such an experiment that included siRNA\ndepletion of DNAJC9 and MCM2 would be more convincing. E.g. in the same\nexperiment comparing.\n1) siControl\n2) siDNAJC9\n3) siMCM2\n4) siHIRA\n5) siDNAJC9+siMCM2\n6) siDNAJC9+siHIRA\n7) siMCM2+siHIRA\nAs shown below (Fig. R1), we had performed siDNAJC9 as a control in the same\nexperiment as siHIRA and observed that while siDNAJC9 reduced old H3.3 recovery,\n3\nconsistent with previous experiments, siHIRA did not diminish old H3.3 recovery. We\ndecided to focus on siHIRA and did not include the siDNAJC9 data in Suppl. Fig. 7g\nbecause it already appeared in other figure panels (Fig. 6c-d).\nFigure R1: Quantification of old H3.3-SNAP histone\nenrichment at sites of UVC irradiation (marked by\nCPD) relative to the nucleus in U2OS cells stably\nexpressing SNAP-tagged H3.3 and treated with the\nindicated siRNAs (siLuc, negative control). Mean ± SD\nfrom cells scored in 3 independent experiments (the\nmean of each experiment is represented by a grey\ntriangle). Statistical significance is calculated by one-\nway ANOVA with multiple comparison.\n• One final point is that the conclusion that siDNAJC9 affects the proportion of cells in S\nphase (Supplementary figure 6h) is based on the proportion of cells that are positive\nfor the histone chaperone Caf1. Can this result be repeated with another more classical\nreadout? E.g. FAC analysis, or using microscopy read-outs e.g. BrdU or EdU?\nWe scored S-phase cells based on CAF-1 staining in Suppl. Fig. 6h because we\nverified that siDNAJC9 did not affect CAF-1 levels (western blot in Suppl. Fig. 6b) and\nwe obtained comparable results by scoring S phase cells based on EdU staining (Fig.\nR2). We now state this in the corresponding figure legend.\nFigure R2: Percentage of cells in S-\nphase determined by CAF-1 p60\nimmunostaining of replication foci (left) or\nby EdU nuclear staining (right) in U2OS\ncells treated with the indicated siRNAs\n(siLuc, control). Mean ± SD from cells\nscored in 3 independent experiments.\nStatistical significance is calculated with a\ntwo-sided Student’s t-test with Welch’s\ncorrection.\n4"}, {"round": 3, "reviewer_comments": {"reviewer_5": "I am satisfied that the authors have now addressed my concerns."}, "author_response": ""}]

Decision letter
[Editorial Note: See attachment]