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

Plessier, Alexandre; Chansard, Audrey; Petit, Eliane; Novion Ducassou, Julia; Couté, Yohann; Polo, Sophie E.

doi:10.1038/s41467-026-68781-x

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Published: 28 January 2026

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

Alexandre Plessier¹^na1,
Audrey Chansard¹^na1,
Eliane Petit¹^na1,
Julia Novion Ducassou²,
Yohann Couté ORCID: orcid.org/0000-0003-3896-6196² &
…
Sophie E. Polo ORCID: orcid.org/0000-0003-4821-9354¹

Nature Communications volume 17, Article number: 2127 (2026) Cite this article

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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.

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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 modifications^2,3 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 marks^2,7.

Epigenome stability is critical for the maintenance of cell function and homeostasis, as underlined by epigenome alterations in disease^8,9. Nevertheless, maintaining epigenome stability is a challenge as all DNA metabolic activities entail some level of chromatin reorganization^10,11. 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 histones^14,15 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 synthesis^15,18,19. 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.

**Fig. 1: IPOND-R, a proteomics method for isolating proteins on nascent DNA at repair sites.**

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 euchromatin^24,25. 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 lines^28,29. 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).

**Fig. 2: Identification of factors enriched at sites of UV damage repair by IPOND-R.**

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 forks^30,31 and the heat-shock protein DNAJC9³² was recently characterized as an evolutionarily conserved H3-H4 histone chaperone^33,34,35. 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 UV^26,27. 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.

**Fig. 3: DNAJC9 and MCM2 histone chaperones are recruited to sites of UV damage repair.**

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 cells^33,34, 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 chaperone^33,34. 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.

**Fig. 4: DNAJC9 stimulates new H3 deposition at sites of UV damage repair.**

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, 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.

**Fig. 5: DNAJC9 promotes old histone H3 recovery during chromatin repair.**

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 histone 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.

**Fig. 6: DNAJC9 and MCM2 cooperate to stimulate new histone deposition and parental histone recovery at DNA repair sites.**

Discussion

Preserving chromatin integrity together with genome stability in response to genotoxic stress is key for safeguarding cell identity and protecting cells from pathological development. Yet, our understanding of chromatin repair mechanisms is still very incomplete. Here, we bridge this knowledge gap and identify new players in these mechanisms through the proteomic profiling of repair patches following UV irradiation. Our methodology could be further applied to explore the proteome at other DNA lesions that require DNA synthesis for their repair.

While similar methods for labeling sites of DNA damage repair in non-replicating cells have been employed in recent works^26,38, our proteomic approach presents several notable improvements compared to previous proteomic studies in UV-damaged cells^26,27: (i) the present approach is focused on repair sites rather than a global analysis of the chromatin-bound proteome post damage infliction, (ii) our method is time-resolved with access to the whole repair process, from early to late repair steps, allowing us to discriminate proteins that dynamically bind to chromatin in response to damage from constitutively bound proteins, and (iii) the increased sensitivity of our methodology allows us to detect low abundant or weakly bound proteins that were not identified in previous datasets.

Among them, we focused our attention on the recently characterized histone chaperone DNAJC9^33,34. Thus, we uncover a function for DNAJC9 in chromatin assembly in vivo, which was previously characterized only in vitro³⁴. Our data support an upstream function of DNAJC9 in the histone supply chain, DNAJC9 providing new H3-H4 histones to CAF-1 and HIRA chaperones for deposition into chromatin. Whether DNAJC9 directly provides histones to CAF-1 and HIRA is still unclear, as the latter chaperones were not systematically identified as DNAJC9 interactors in basal conditions^34,39,40. By ensuring H3-H4 histone supply, DNAJC9 prevents mislocalization of the centromeric H3 variant CENP-A to chromosome arms⁴¹. It is thus tempting to speculate that DNAJC9 may similarly prevent ectopic CENP-A deposition at sites of UV damage repair.

In addition to the reported function of DNAJC9 in controlling histone protein folding in the cytoplasm^33,34, our work highlights a nuclear function of DNAJC9 in coordinating old and new H3-H4 histone dynamics during the repair process. Indeed, we show that DNAJC9 cooperates with MCM2 to stimulate both new histone deposition and old histone recovery during repair, which constitutes the first proof of a functional cooperation between these two chaperones. Our discoveries not only reveal a function of these histone chaperones in chromatin repair but also instruct us on their activity and central position in the histone supply chain, at the crossroads of new histone deposition and old histone recycling (Fig. 6f). The mechanism whereby DNAJC9 and MCM2 stimulate parental histone recovery during UV damage repair warrants further investigation.

MCM2 is enriched in the IPOND-R dataset together with the other MCM subunits (MCM3 to 7), showing that the whole MCM hexamer is present at sites of UV damage repair, like at replication forks. However, the observed recruitment of MCM2 to sites of UV damage repair does not require the passage of replication forks because it is detected outside of S phase, highlighting a function of MCM2 that is disconnected from DNA replication. In line with this, we do not retrieve other components of the replicative helicase complex at repair sites like Cdc45 and GINS subunits, which indicates that MCM2 likely operates independently of the replicative helicase complex at repair sites. Notably, an abundant pool of MCM proteins is not engaged with active replisomes in human cells⁴² and those could fulfill replication-independent roles.

Several pieces of evidence also suggest that DNAJC9 may coordinate new and parental histone dynamics in contexts other than DNA repair. Indeed, DNAJC9 is found enriched on nascent replicated chromatin²⁸ and we show that DNAJC9 stimulates new H3.1 deposition at undamaged replication forks. A potential role of DNAJC9 in parental histone recycling at replication forks, possibly in coordination with MCM2³⁰, deserves to be investigated.

How MCM2 and DNAJC9 are recruited to sites of UV damage repair is not yet elucidated. It is tempting to speculate that their recruitment may be mediated by direct or indirect interactions with UV damage repair factors or with other histone chaperones enriched at repair sites. In this respect, MCM2 is involved in histone-dependent interactions with the FACT histone chaperone³⁹ that is recruited to UV sites coupled to repair synthesis¹⁵. An additional candidate that could promote DNAJC9 recruitment to UV-damaged chromatin is RAD23A, which was identified in the DNAJC9 interactome in human cells³³ and binds the early UV damage repair factor XPC^43,44. UV-damage responsive post-translational modifications of these histone chaperones may also regulate their function in chromatin repair²⁷. Our proteomic approach does not distinguish between repair in euchromatin and heterochromatin. However, we envision that histone chaperones may play distinct roles in different chromatin compartments. The use of murine cellular models suited for the analysis of heterochromatin repair of UV lesions¹⁴ will be instrumental to address this issue.

We do not rule out the possibility that DNAJC9 and MCM2 may respond to other types of DNA lesions than UV photoproducts, and thus play a more general role in maintaining epigenome stability following genotoxic stress. Interestingly, a fungal ortholog of DNAJC9, Dnaj4, which also exhibits H3-H4 histone chaperone activity, was shown to safeguard genome integrity at least in part through the control of gene expression in response to DNA damage⁴⁵. Together with our findings, this hints towards an evolutionarily conserved function of DNAJC9 in genome and epigenome maintenance. DNAJC9 is a dual heat shock molecular and histone chaperone that recruits heat shock proteins of the HSP70 family via its J domain to promote histone protein folding^33,34. Whether DNAJC9 functions in chromatin repair depend on binding to HSP70 family proteins remains to be investigated, which could be tested using separation of function mutants in the J domain and the histone binding domain of DNAJC9³⁴. Notably, HSPA4/HSP74 is significantly enriched at sites of UV damage repair in the 1 h IPOND-R dataset and thus represents a potential candidate that could cooperate with DNAJC9 in controlling histone dynamics during repair.

While in this study we focused on the histone chaperones DNAJC9 and MCM2, our proteomic analysis uncovered a number of additional interesting candidates as potential new players in chromatin dynamics during DNA repair. Among them, several histone modifiers and readers highlight a fine-tuned regulation of histone modifications at repair sites, which deserves to be investigated in future studies. The histone modifiers and readers identified in our dataset and by others^26,27 will provide clues to the underlying mechanisms. Along the same lines, the DNA modifiers enriched at sites of UV damage repair will help characterize the machinery for repair-coupled DNA methylation maintenance, thus broadening our understanding of epigenome maintenance mechanisms in response to DNA damage. Intriguingly, several nuclear pore components and lamins were also enriched at sites of UV damage repair, as seen in a previous study²⁶. There is growing evidence for a role of the nuclear periphery in the repair of DNA double-strand breaks, with DSBs relocalizing to the nuclear periphery for repair in yeast and flies, while in human cells, the nuclear envelope forms intra-nuclear tubules that capture DSBs for repair, and nuclear pores and lamina-associated chromatin domains regulate DSB repair pathway choice^{46,47,48,49,50}. However, the connection of UV damage repair with the nuclear periphery is still elusive and will be interesting to explore to further dig into how nuclear architecture may govern genome maintenance and reciprocally. Overall, our proteomic dataset provides a molecular foundation for further exploration of the fundamental mechanisms underpinning chromatin plasticity and integrity following genotoxic stress.

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.

Table 1 Stable cell lines

Full size table

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.

Table 2 siRNA sequences

Full size table

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 Neon^TM Transfection System (Thermo Scientific) according to the manufacturer’s instructions and harvested 72 h later.

Table 3 Plasmids

Full size table

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 irradiation^51,52, 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 (RappOptoElectronics, 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 SysCon2 software (RappOpto). 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, Euromedex) 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-Naegel 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 1 X 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 (Bio Basic 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 nanoliquid 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, IonOpticks, 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 2.0^59,60 curated based on most recent literature. The proteins detected by IPOND-R coupled to mass spectrometry were visualized using Volcano plots generated with R studio 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 AlexaFluor 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 AlexaFluor 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 histones^36,61, 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 TMR star (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 TMR star (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 TMR star (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 epifluorescence 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 leads 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.

Reporting summary

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

Data availability

All data generated during this study are included in this article and its supplementary information files. Mass spectrometry data have been deposited to the ProteomeXchange Consortium via the PRIDE partner repository with the dataset identifier PXD055129. Source data have been deposited in the Figshare Repository (https://doi.org/10.6084/m9.figshare.30665291). Source data are provided with this paper.

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Acknowledgements

The authors thank Colin Hammond and Anja Groth for sharing plasmids, the EPI² Imaging platform and in particular Sandra Piquet—UMR7216 Epigenetics and Cell Fate Centre for access to instruments and technical advice. This work was supported by the European Research Council (ERC-2018-CoG-818625 “REMIND” to S.E.P.). The proteomic experiments were partially supported by Agence Nationale de la Recherche under projects ProFI (Proteomics French Infrastructure, ANR-10-INBS-08) and GRAL, a program from the Chemistry Biology Health (CBH) Graduate School of University Grenoble Alpes (ANR-17-EURE-0003) to Y.C.

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These authors contributed equally: Alexandre Plessier, Audrey Chansard, Eliane Petit.

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Laboratory of Epigenome Integrity, Epigenetics and Cell Fate Centre, UMR7216 CNRS, Université Paris Cité, Paris, France
Alexandre Plessier, Audrey Chansard, Eliane Petit & Sophie E. Polo
Grenoble Alpes University, CEA, INSERM, UA13 BGE, CNRS, CEA, UAR2048, Grenoble, France
Julia Novion Ducassou & Yohann Couté

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Alexandre Plessier
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Audrey Chansard
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Eliane Petit
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Julia Novion Ducassou
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Yohann Couté
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Sophie E. Polo
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A.P., A.C., E.P., and S.E.P. designed and performed experiments, analyzed the data, and wrote the manuscript. J.N.D. and Y.C. performed MS-based proteomic analyses. S.E.P. supervised the project.

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Correspondence to Sophie E. Polo.

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Plessier, A., Chansard, A., Petit, E. et al. Proteomic profiling of UV damage repair patches uncovers histone chaperones with central functions in chromatin repair. Nat Commun 17, 2127 (2026). https://doi.org/10.1038/s41467-026-68781-x

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Received: 04 November 2024
Accepted: 16 January 2026
Published: 28 January 2026
Version of record: 03 March 2026
DOI: https://doi.org/10.1038/s41467-026-68781-x

Subjects

Abstract

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Introduction

Results

The IPOND-R method captures proteins at sites of DNA damage repair

Identification of chromatin regulators enriched at sites of UV damage repair

The histone chaperone DNAJC9 is not involved in DNA repair following UV damage

DNAJC9 promotes the deposition of newly synthesized H3 variants at sites of UV damage repair

DNAJC9 promotes the recovery of parental H3 histones at sites of UV damage repair

DNAJC9 cooperates with MCM2 to coordinate old and new histone dynamics at sites of UV damage repair

Discussion

Methods

Cell culture

siRNA and plasmid transfections

UVC irradiation

IPOND-R (Isolation of Proteins On Nascent DNA at Repair sites)

EdU labeling of repair sites

Cell fixation and permeabilization

EdU biotinylation by Click-it chemistry

Cell lysis and sonication

Analysis of DNA shearing

Streptavidin capture of biotin-labeled nascent DNA and associated proteins

Mass spectrometry-based proteomic analyses

Data visualization and gene ontology analysis

EdU-labeling of replicating cells and repair sites

EU-labeling of nascent transcripts at UV damage sites

Immunofluorescence

SNAP-tag labeling of histones

Image acquisition and analysis

Cell extracts and Western blot

Flow cytometry

Cell survival assays

Statistical analyses

Reporting summary

Data availability

References

Acknowledgements

Author information

Authors and Affiliations

Contributions

Corresponding author

Ethics declarations

Competing interests

Peer review

Peer review information

Additional information

Supplementary information

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