XPC–PARP complexes engage the chromatin remodeler ALC1 to catalyze global genome DNA damage repair

ARTICLE XPC–PARP complexes engage the chromatin remodeler ALC1 to catalyze global genome DNA damage repair Charlotte Blessing 1,2,13, Katja Apelt3,13, Diana van den Heuvel3, Claudia Gonzalez-Leal1,2, Magdalena B. Rother3, Melanie van der Woude4, Román González-Prieto 5,6,7, Adi Yifrach8, Avital Parnas8, Rashmi G. Shah9, Tia Tyrsett Kuo 1,2, Daphne E. C. Boer3, Jin Cai1,2, Angela Kragten3, Hyun-Suk Kim10, Orlando D. Schärer10,11, Alfred C. O. Vertegaal5, Girish M. Shah 9, Sheera Adar8, Hannes Lans 4, Haico van Attikum3, Andreas G. Ladurner 1,2,12,14✉& Martijn S. Luijsterburg 3,14✉ Cells employ global genome nucleotide excision repair (GGR) to eliminate a broad spectrum of DNA lesions, including those induced by UV light. The lesion-recognition factor XPC initiates repair of helix-destabilizing DNA lesions, but binds poorly to lesions such as CPDs that do not destabilize DNA. How difficult-to-repair lesions are detected in chromatin is unknown. Here, we identify the poly-(ADP-ribose) polymerases PARP1 and PARP2 as constitutive interactors of XPC. Their interaction results in the XPC-stimulated synthesis of poly- (ADP-ribose) (PAR) by PARP1 at UV lesions, which in turn enables the recruitment and activation of the PAR-regulated chromatin remodeler ALC1. PARP2, on the other hand, modulates the retention of ALC1 at DNA damage sites. Notably, ALC1 mediates chromatin expansion at UV-induced DNA lesions, leading to the timely clearing of CPD lesions. Thus, we reveal how chromatin containing difficult-to-repair DNA lesions is primed for repair, providing insight into mechanisms of chromatin plasticity during GGR. https://doi.org/10.1038/s41467-022-31820-4 OPEN 1Biomedical Center (BMC), Physiological Chemistry, Faculty of Medicine, LMU Munich, Planegg-Martinsried, Germany. 2International Max Planck Research School (IMPRS) for Molecular Life Sciences, Planegg-Martinsried, Germany. 3Department of Human Genetics, Leiden University Medical Center (LUMC), Leiden, The Netherlands. 4Department of Molecular Genetics, Erasmus MC Cancer Institute, Erasmus University Medical Center, Rotterdam, The Netherlands. 5Department of Cell and Chemical Biology, Leiden University Medical Center (LUMC), Leiden, The Netherlands. 6Genome Proteomics Laboratory, Andalusian Center For Molecular Biology and Regenerative Medicine (CABIMER), University of Seville, Seville, Spain. 7Department of Cell Biology, University of Seville, Seville, Spain. 8Department of Microbiology and Molecular Genetics, The Institute for Medical Research Israel-Canada, The Faculty of Medicine, The Hebrew University of Jerusalem, Jerusalem, Israel. 9Laboratory for Skin Cancer Research, CHU-Q: Laval University Hospital Research Centre of Quebec (CHUL site), Quebec City, Canada. 10 Center for Genomic Integrity, Institute for Basic Science, Ulsan, Republic of Korea. 11 Department of Biological Sciences, School of Life Sciences, Ulsan National Institute of Science and Technology, Ulsan, Republic of Korea. 12 Eisbach Bio GmbH, Planegg-Martinsried, Germany. 13 These authors contributed equally: Charlotte Blessing, Katja Apelt. 14 These authors jointly supervised this work: Andreas G. Ladurner, Martijn S. Luijsterburg. ✉email: [email protected];[email protected] NATURE COMMUNICATIONS | (2022) 13:4762 | https://doi.org/10.1038/s41467-022-31820-4 | www.nature.com /naturecommunications 1 1234567890():,; The integrity of the human genome is constantly threatened by endogenous and exogenous sources, which cause up to 105DNA lesions per cell per day1. Cells thus critically depend on the accuracy of dedicated DNA repair mechanisms to recognize and remove genomic DNA lesions and maintain genome integrity2. One principal source of DNA damage is UV light, which results in the crosslinking of neighboring bases on the same DNA strand, forming so-called 6-4 photoproducts (6-4PPs) and cyclobutane pyrimidine dimers (CPDs). Nucleotide excision repair (NER) can repair these bulky lesions3. The initiation of this repair pathway depends on the position of the lesion in the genome. While RNA polymerase II stalling at lesions in transcribed strands initiates transcription-coupled repair (TCR or TC-NER)4,5, lesions in transcriptionally inactive genome regions are recognized by specialized damage sensors that initiate global genome repair (GGR or GG-NER)6,7. Recognition through both sub-pathways ultimately leads to a common pathway of verification, excision, and re-synthesis of the damaged DNA. In principle, all core NER factors have been identified and the fundamental DNA repair process can be reconstituted in vitro8,9. However, the cellular repair mechanism is not well understood in the chromatin context, as knowledge about chromatin factors that allow and promote the efficient action of the core repair factors is limited. In mammalian cells, GGR is initiated by XPC, which forms a complex with RAD23B and CEN26,10–12. Rather than binding to lesions directly, XPC binds the accessible, non-damaged DNA that is opposite the DNA injury. This allows the recognition of a broad spectrum of lesions that are structurally unrelated13,14. XPC binding results in the slight opening of the DNA surrounding a lesion (~6 nucleotides)15, which facilitates the binding of subsequent factors. The recruitment of these downstream NER proteins, including TFIIH, RPA, XPA, XPG, and ERCC1-XPF, to sites of UV-induced DNA lesions is abolished in XPC-deficient cells. This demonstrates that the GGR repair pathway is strictly dependent on XPC10. Although XPC has a high affinity for 6-4PPs, it's binding to CPDs is rather inefficient due to minimal thermodynamic helix destabilization caused by the latter lesion. The recognition and repair of CPDs, therefore, requires the additional action of the damaged DNA-binding protein 216, which does not seem to influence in vitro reconstituted NER8,17. DDB2 further utilizes slide-assisted site exposure to detect inaccessible lesions occluded in nucleosomes18, and creates a local chromatin environment around lesions that facilitates the assembly of repair complexes19–22. DDB2 is thus often considered the factor that prepares chromatin for GGR, and its dissociation from DNA lesions is subsequently required for the progression of repair23. Whether this is an exclusive feature of DDB2 or whether and how XPC also contributes to local chromatin changes is unknown. In addition to XPC and DDB2, PARP1 is also known to associate with UV-induced DNA lesions resulting in poly-(ADPribos)ylation (PARylation) at sites of DNA damage24–26. In vitro approaches showed that PARP1 and DDB2 can simultaneously bind to a UV-induced CPD25. In agreement, in situ fractionation showed that endogenous PARP1 is recruited to sites of local UV damage24,25,27. However, the precise interplay between the three lesion-recognition proteins XPC, DDB2 and PARP1 during GGR is poorly understood. Initially, DDB2 was found to associate with and potentially stimulate the catalytic activity of PARP1, resulting in the PARylation of DDB2. This was suggested to counteract DDB2 auto-ubiquitylation and its subsequent degradation28. Consistently, inhibition of PARP activity was found to accelerate DDB2 degradation28 and to reduce XPC recruitment to UV lesions under conditions of low damage load20,29. PARP1 was also shown to stimulate XPC recruitment to DNA damage sites in a DDB2-independent manner and to regulate XPC release24. Thus, it appears that both DDB2 and PARP1 may stimulate XPC recruitment to initiate GGR. However, the relevance of the poly- (ADP-ribose) response for GGR and how it is related to XPC activity remains to be established. In addition to PARP1, the nuclear and DNA damagedependent PARP2 enzyme seems to have independent functions in the DNA damage response, which are not well understood. In general, PARP1 is thought to provide ~90% of the PAR signal at DNA lesions, while PARP2 contributes a minor part30,31. Instead, PARP2 is suggested to increase the branching of PAR chains32. The double knockout of both PARP1 and PARP2 is embryonic lethal in mice33 and renders cells highly sensitive to DNA-damaging agents, such as the alkylating agent methyl methanesulfonate34. This suggests that both proteins cooperatively act in the DNA damage response. However, whether PARP2 contributes to the poly-(ADP-ribose) response and has roles in GGR remains to be established. Here, we sought to elucidate the role of PARylation, PARP2, and active chromatin remodeling in GGR. Several chromatin remodelers have been implicated in DNA repair, notably ALC1, which acts downstream of PARP1/2 activation, due to its strict PARylation-dependent nucleosome remodeling activity35–38. Using proteomics, live-cell imaging, and UV-induced DNA damage, our data revealed a new XPC-PARP axis that links ATPdependent and PARylation-activated ALC1-mediated chromatin remodeling to GGR. We identify the poly-(ADP-ribose) polymerases PARP1 and PARP2 as constitutive interactors of the damage-recognition protein XPC. The close interaction between these proteins results in an XPC-dependent stimulation of the poly-(ADP-ribose) response, which facilitates the recruitment of the poly-(ADP-ribose)-dependent chromatin remodeler ALC1. We thus identify a new XPC-dependent mechanism that impacts the chromatin environment and promotes chromatin remodeling at UV lesions. Results XPC interacts with PARP1 and PARP2. To identify potential new factors involved in GGR, we analyzed the interactome of the damage-recognition factor XPC by mass spectrometry. We generated a knockout (KO) of XPC in U2OS (FRT) cells containing an FRT Flp-In integration site. Having confirmed the successful knockout of XPC by sequencing and western blot analysis (Supplementary Fig. 1a), we exploited the site-specific transgene integration of the Flp-In system to re-express XPC-GFP in these cells under a doxycycline-inducible promoter. To demonstrate the functionality of our newly generated cell system, we measured the ability to repair UV lesions in unscheduled DNA synthesis (UDS) assays. While the XPC-KO showed a severe repair defect, the re-expression XPC-GFP restored the capacity of the cells to repair UV-induced DNA damage (Supplementary Fig. 1b, c). Label-free proteomics after pull-down of XPC-GFP revealed several known XPC-binding proteins, including RAD23A/B and CEN2/3, as the top interactors (Fig. 1a). After UV irradiation, XPC additionally became tightly bound to several GGR factors, such as the DDB2 complex (containing DDB1 and CUL4A/B), the TFIIH subunits GTF2H1-4 (p62, p44, p34, p52), XPB (p89/ ERCC3) and XPD (p80/ERCC2) (Fig. 1b). This confirms that our label-free proteomics approach is suitable to detect interactions within an active GGR process. Interestingly, our analysis further identified the poly-(ADPribose) polymerases PARP1 and PARP2 as strong XPC-associated proteins (Fig. 1a). In contrast to the main GGR factors, the interaction between XPC and PARP1/2 was not significantly ARTICLE NATURE COMMUNICATIONS | https://doi.org/10.1038/s41467-022-31820-4 2NATURE COMMUNICATIONS | (2022) 13:4762 | https://doi.org/10.1038/s41467-022-31820-4 | www.nature.com/naturecommunications affected by UV irradiation (Fig. 1b). Immunoprecipitation experiments confirmed that the XPC-PARP1/2 interaction was constitutive and independent of PARylation, as treatment with the PARP inhibitor olaparib did not affect the interactions (Fig. 1c). Our findings indicate that XPC forms a constitutive interaction with both PARP1 and PARP2, which is not affected by DNA damage or PARylation. XPC and ALC1 interact more strongly with PARP2 than PARP1. To obtain first insights into the role of PARP1 and PARP2 in GGR and the relevance of their interaction with XPC, we conducted an orthogonal experiment to identify the interactome of the two PARP enzymes. We stably expressed PARP1GFP or GFP-PARP2 in U2OS (FRT) cells (Supplementary Fig. 1d) and performed label-free proteomics after GFP-pulldown of the tagged proteins. PARP1 most abundantly interacted with XRCC1-LIG3 and POLB, and further showed robust interactions with PARP2, histones, as well as the known poly-(ADPribose)-binding proteins ALC1 and macroH2A (Fig. 2a). In contrast, XPC was not significantly enriched in the PARP1 interactome (Fig. 2a, b), which is likely caused by the high abundance of PARP1 and a potentially low stoichiometric interaction with XPC. It should be noted that earlier studies did report an interaction between immunoprecipitated endogenous PARP1 and XPC24. Interestingly, the interactome of PARP2 revealed the poly- (ADP-ribose)-dependent chromatin remodeler ALC1 as its most abundant interactor. PARP2 further interacted with XRCC1LIG3, PARP1, histones, and macroH2A (Fig. 2c). PARP2 also clearly interacted with XPC in a manner that was not affected by UV irradiation (Fig. 2c, d). Intensity-based absolute quantification (iBAQ) of protein amounts indicated that ~15% of the isolated PARP2 molecules were associated with ALC1, while only 0.07% of PARP1 molecules interacted with the remodeler. Additionally, the fraction of PARP2 molecules associated with XPC was ten-fold higher than for PARP1 (Fig. 2e). The stoichiometry among PARP enzymes was very low with 1.76% of the PARP2 molecules interacting with PARP1, ruling out indirect interactions through PARP heterodimerization. Immunoprecipitation experiments confirmed that PARP2 robustly interacted with both ALC1 and XPC, while these interactions were not or only weakly detected after pull-down of PARP1 (Fig. 2f). This demonstrates that XPC and ALC1 both preferentially associate with PARP2 over PARP1 under our experimental conditions. PARP1/2 recruitment to UV lesions is independent of XPC. The DNA damage-recognition proteins XPC and DDB2 can both bind to UV-induced lesions. DDB2 thereby stimulates XPC recruitment at difficult-to-detect lesions, such as CPDs16,20,39, and facilitates lesion-recognition by XPC in a chromatin context. PARP1 was also reported to bind to UVinduced DNA lesions together with XPC24,25.Consideringthe interaction between XPC and PARP1/2, we asked whether the PARP enzymes are recruited to UV lesions and if this is stimulatedbyXPC.Tothisend,weemployedaUV-C(266nm) laser live-cell imaging set-up, in which all optics have been replaced by quartz glass. Local irradiation with UV-C led to the a UV XPC-GFP XPC-GFP Input GFP NLS -+-+ -+ PARP2 PARP1 Olaparib - - - - + + GFP-NLS IP: GFP XPC-GFP PARP2 PARP1 GFP-NLS b c UV XPC-GFP GFP NLS -+-+ -+ Olaparib - - - - + + - 100 - 25 - 75 - 150 kDa - 100 - 25 - 75 - 150 kDa 0 1 2 3 4 5 -4 -2 0 2 4 6 XPC (+UV) vs. XPC (-UV) Significance(-log 10 ) Difference (log 2 ) XPD CUL4A NEDD8 GTF2H4 XPB CUL4B GTF2H5 GTF2H3 GTF2H1 DDB2 DDB1 PARP1 PARP2 XPC XPC RAD23B RAD23A CEN2 CEN3 XPB GTF2H4 GTF2H1 PARP2 GTF2H2 PARP1 0 2 4 6 8 -8 -4 0 4 8 12 16 Significance(-log 10 ) Difference (log 2 ) XPC-GFP vs. GFP-NLS Fig. 1 XPC interacts with PARP1 and PARP2 in a UV-independent manner. a Volcano plot displaying the interactome of XPC-GFP over GFP-NLS after GFP-pull-down from U2OS (FRT) XPC-KO cells and analysis by label-free proteomics. bDifferential interactome of XPC-GFP comparing UV-C-irradiated (20 J/m2, 1 h) vs. unirradiated U2OS (FRT) XPC-KO cells. a,bThe dashed lines indicate a twofold enrichment on the xaxis (log 2 of 1) and a significance of 0.05 (−log 10 Pvalue of 1.3; two-sided ttest) on the yaxis. cCo-Immunoprecipitation (Co-IP) of GFP-NLS and XPC-GFP in the presence and absence of UVC (20 J/m2, 1 h) and the PARP inhibitor olaparib (10 µM). Three independent replicates of each IP experiment were performed obtaining similar results. NATURE COMMUNICATIONS | https://doi.org/10.1038/s41467-022-31820-4 ARTICLE NATURE COMMUNICATIONS | (2022) 13:4762 | https://doi.org/10.1038/s41467-022-31820-4 | www.nature.com /naturecommunications 3 rapid recruitment of PARP1-GFP to sites of DNA damage within seconds, after which steady-state bound levels decreased in the first several minutes (Fig. 3a, b). GFP-PARP2 was also recruited to UV-C laser damage. Steady-state bound levels remained high for ~10 min without an apparent decrease (Fig. 3c, d). The enrichment of PARP1-GFP shortly after UV-C micro-irradiation was more pronounced (1.4-fold) compared to the more modest recruitment of PARP2-GFP (1.2-fold). Interestingly, the recruitment kinetics of PARP1 and PARP2 were identical in XPC-KO cells (Fig. 3b, d). This indicates that Difference (log 2 ) a cd b PARP1-GFP PARP1 GFP NLS ALC1 GFP-PARP2 XPC PARP2 GFP-NLS Input IP: GFP e UV -+-+ -+ PARP1-GFP PARP1GFP NLS ALC1 GFP-PARP2 XPC PARP2 GFP-NLS UV -+-+ -+ f PARP1 PARP2 0.00 0.05 0.10 0.15 iBAQ ratio (%) XPC PARP1 PARP2 0.0 0.1 5 10 15 20 iBAQ ratio (%) ALC1 PARP1 PARP2 0 4 20 40 60 80 100 iBAQ ratio (%) PARP1 PARP1 PARP2 0.0 0.1 20 40 60 80 100 iBAQ ratio (%) PARP2 - 100 - 150 kDa - 100 - 100 - 25 - 100 - 150 kDa - 100 - 100 - 25 PARP1 LIG3 XRCC1 APTX POLB PARP2 CHFR ALC1 SPT16 H1 H2A APLF H3 macro H2A H2B XPC 0 2 4 6 8 10 12 14 -5 0 5 10 15 PARP1-GFP vs. GFP-NLS Significance(-log 10 ) ) SETDB1 YBX3 CTCF YBX1 TCF20 WDR18 H3 PARP1 0 1 2 3 4 5 6 7 8 -3 -1 1 3 5 PARP1 (+UV) vs. PARP1 (-UV) Difference (log 2 ) Significance(-log 10 ) PARP2 ALC1 LIG3 XRCC1 APTX macro H2A H2A PARP1 H1x H3 XPC 0 2 4 6 8 10 -3036912 GFP-PARP2 vs. GFP-NLS Significance(-log 10 ) Difference (log 2 ) macro H2A.2 YBX3 YBX1 H3 C1ORF174 H2B RSF1 H2AZ PARP2 XPC TRIM22 0 1 2 3 4 5 6 -2-1012345 PARP2 (+UV) vs. PARP2 (-UV) Significance(-log 10 ) Difference (log 2 ) XPC ARTICLE NATURE COMMUNICATIONS | https://doi.org/10.1038/s41467-022-31820-4 4NATURE COMMUNICATIONS | (2022) 13:4762 | https://doi.org/10.1038/s41467-022-31820-4 | www.nature.com/naturecommunications PARP enzymes accumulate independently of XPC at sites of UV-induced DNA damage, which is in line with earlier findings showing that PARP1 recruitment is similar between WT and XPC-deficient cells at 10 min after UV irradiation24. Deletion of either PARP1 or PARP2 had a minor impact on the recruitment of XPC to UV lesions, as measured by immunofluorescence after local UV irradiation through micropore filters (Fig. 3e, f). The impact of PARP enzymes was milder than KO of DDB2, which led to considerably reduced XPC recruitment, as reported previously16,20,39. Our data thus suggest that XPC and PARP enzymes are recruited to DNA lesions largely independently, although PARP enzymes may stimulate XPC recruitment under certain conditions, such as low doses of DNA damage20,24. Fig. 2 XPC and ALC1 interact more strongly with PARP2 than PARP1. a,cVolcano plots displaying the interactomes of aPARP1-GFP and cGFP-PARP2 after GFP-pull-down from U2OS (FRT) WT cells and analysis by label-free proteomics. b,dDifferential interactomes of bPARP1-GFP and dGFP-PARP2 from UV-C-irradiated (20 J/m2, 1 h) vs. unirradiated U2OS (FRT) WT cells. a–dThe dashed lines indicate a twofold enrichment on the xaxis (log 2 of 1) and a significance of 0.05 (−log 10 Pvalue of 1.3; two-sided ttest) on the yaxis. eEnrichment of XPC, ALC1, PARP1, and PARP2 in the co-IPs of PARP1 (a) and PARP2 (b), calculated by intensity-based quantification (IBAQ). Each data point represents a biological replicate (n=4). fCo-IP of GFP-NLS, PARP1-GFP, and GFPPARP2 in the presence and absence of UV-C (20 J/m2, 1 h). Three independent replicates of each IP experiment were performed obtaining similar results. a 0 100 200 300 400 500 600 1.0 1.1 1.2 1.3 1.4 1.5 Time after irradiation (s) WT XPC-KO Relative recruitment 1.5 0 100 200 300 400 500 600 1.0 1.1 1.2 1.3 1.4 Time after irradiation (s) WT XPC-KO XPC-KO WT pre-damage 30 s 10 min PARP1-GFP GFP-PARP2 pre-damage 30 s 10 min b cd e WT DDB2 KO WT PARP1 KO PARP2 KO HoechstXPCCPD XPC f Relative recruitment PARP1-GFP GFP-PARP2 XPC-KO WT WT PARP1-KO PARP2-KO WT DDB2-KO 0 1 2 3 Relative XPC enrichment CPD 0 1 2 3 Relative CPD enrichment WT PARP1-KO PARP2-KO WT DDB2-KO Fig. 3 PARP1/2 recruitment to UV lesions is independent of XPC. a,cRepresentative images of aPARP1-GFP and cGFP-PARP2 association with sites of local UV-C laser irradiation in U2OS (FRT) WT and XPC-KO cells at 30 sec and 10 min post-irradiation. b,dKinetics of the recruitment of bPARP1-GFP and dGFP-PARP2 to and dissociation from UV-C lesions measured over 10 min in U2OS (FRT) WT and XPC-KO cells; 70–91 nuclei were analyzed in three independent biological replicates. The data are shown as mean +SEM normalized to pre-damage GFP intensity at micro-irradiation sites. eRepresentative images and fquantification of XPC colocalization with local UV-C irradiation sites (100 J/m2) marked by CPD, measured 10 min post-irradiation. In all, 58–230 cells were analyzed per condition. All cells are depicted as individual data points (gray). The medians of four biological replicates are depictedas colored points, while the bar represents the median of all data points. The scale bar in a,c,eis 5 µm. NATURE COMMUNICATIONS | https://doi.org/10.1038/s41467-022-31820-4 ARTICLE NATURE COMMUNICATIONS | (2022) 13:4762 | https://doi.org/10.1038/s41467-022-31820-4 | www.nature.com /naturecommunications 5 XPC stimulates the poly-(ADP-ribose) response to UV lesions. The PARP response involves the rapid and robust DNA damageinduced PARylation mainly of PARP1 itself, but also of other chromatin substrates40. To better understand how the PARP response modulates GGR, we measured nuclear PAR levels at sites of local UV-induced DNA damage by immunofluorescence in PARP1-KO and PARP2-KO cells (Fig. 4a, b). PARylation strongly increased at sites of local UV irradiation, marked by DDB2 recruitment, in WT cells and PARP2-KO cells, but not in PARP1-KO cells (Figs. 4a, b; Supplementary Fig. 2a). Similarly, nuclear PAR levels also increased after global UV irradiation in a manner that was dependent on PARP1, but not on PARP2 UV -+-+-+ WT PARP1KO PARP2KO 0 1 2 3 4 5 6 7 Relativenuclear PAR levels a b e f cWT PARP1-KO PARP2-KO HoechstPAR 5 min post UV d h g 1.8 PAR signal 1.6 1.4 1.2 1.0 0.8 XPC PARP1 1 1 1 0.25 0.501 1 In vitro PARylation 0 1 2 3 4 5 6 7 8 WT XPCKO DDB2KO UV-+-+-+ Relative nuclear PARlevels WT XPC-KO DDB2-KO 5 min post UV Hoechst PAR XPC PARP1 XPC PARP1 - + + -1 1 1 1 0.25 0.50 1In vitro PARylation PAR 116 - kDa 116 - 200 - i j kl WT PARP1-KO PARP2-KO DDB2PAR 10 min post UV WT PARP1 KO 0 1 2 3 4 5 Relative local PAR levels PARP2 KO 0 5 4 3 2 1 relative local PAR signal WT XPC KO 0 5 4 3 2 1 relative local PAR signal WT DDB2 KO WT XPC-KO DDB2PAR 5 min post UV WT DDB2-KO XPCPAR 5 min post UV ARTICLE NATURE COMMUNICATIONS | https://doi.org/10.1038/s41467-022-31820-4 6NATURE COMMUNICATIONS | (2022) 13:4762 | https://doi.org/10.1038/s41467-022-31820-4 | www.nature.com/naturecommunications (Fig. 4c, d; Supplementary Fig. 2b). This demonstrates that the PAR response at UV lesions is largely dependent on PARP1, as demonstrated for other DNA-damaging agents29,41–43. Having established that XPC interacts with PARP1 and PARP2 (Figs. 1and 2), we sought to investigate whether XPC impacts the PAR response. Strikingly, XPC-KO cells showed an attenuated PAR response at sites of local UV damage, while DDB2-KO cells established PAR levels similar to wild-type U2OS cells (Fig. 4e–h; Supplementary Fig. 2c, d). Identical results were obtained when PAR levels were monitored after global UV irradiation (Figs. 4i, j; Supplementary Fig. 2e). Knockdown of XPA in XPC-deficient cells did not further reduce DNA damage-induced PARylation at sites of local UV-induced DNA damage (Supplementary Fig. 2f±i), suggesting that this phenomenon is dependent on GGR and not on transcription-coupled repair. To understand whether XPC may directly stimulate the activity of PARP enzymes through their interaction, we performed in vitro PARylation assays using recombinant PARP1 in the presence of UV-irradiated DNA and increasing amounts of recombinant XPC-RAD23B complex (Fig. 4k, l). In this minimal in vitro system without additional components, we observed that XPC directly stimulated the catalytic activity of PARP1 by ~1.5 fold, which was already observed at a 4:1 ratio of PARP1 over XPC-RAD23B complex and did not increase when more XPC was added to the reaction (Fig. 4k, l). These findings show that XPC stimulates the initial and rapid PAR response at UV lesions by enhancing the protein activity of PARP1. PARP1 and ALC1 are UV-induced substrates of PARylation. Based on the close interaction of XPC and PARP1/2, we next asked which of these proteins become PARylated upon UV damage. To this end, we performed pull-down experiments under high-salt conditions to disrupt protein-protein interactions and capture the PARylation status of the immunoprecipitated proteins. Pull-down of PARP1-GFP revealed robust PARylation in response to UV irradiation (Fig. 5a). By contrast, GFP-PARP2 and XPC-GFP were already PARylated in control cells at lower levels with no further increase following UV irradiation (Fig. 5b, c). The poly-(ADP-ribose)-dependent chromatin remodeler ALC1 was also strongly PARylated in response to UV irradiation (Fig. 5d). To better capture the dynamics of protein PARylation in response to UV irradiation, we performed an adapted LacO-based colocalization assay, in which we fused the PAR-binding macrodomain of macroH2A1.1 to LacR and tethered this PARbinding module to a LacO array in U2OS 263 cells44,45. We then induced the PARylation response by local irradiation with a 266 nm UV-C laser. Micro-irradiation triggered the recruitment of GFP-tagged versions of XPC, ALC1, PARP1, and PARP2 to sites of UV-C-induced laser damage (Figs. 5e, f and Supplementary Fig. 3a). Within one minute following irradiation, we also detected the capture of XPC, ALC1 and PARP1 at the LacO array bound by the LacR-fused macrodomain PAR-binding module. This suggests that protein complexes containing XPC, ALC1 and PARP1 become PARylated at laser micro-irradiation sites and dissociate from these sites in a modified, PARylated state, which in turn allows their interaction with the immobilized macrodomain at the LacO array. Treatment with the PARP inhibitor olaparib prevented the accumulation of XPC at the LacO site, demonstrating that its capture at the LacO array is fully dependent on PARylation (Fig. 5e, f). Interestingly, PARP2 was recruited to UV sites, but could not be detected at the LacO site following micro-irradiation, suggesting that its UV-induced PARylation is not sufficiently high or that the protein is not sufficiently mobile to enable capture of PARP2 by the immobilized PAR-binding module at the LacO array. The capture of either PARP1 or ALC1 at the immobilized PAR-binding module is consistent with their UV-induced PARylation detected in pull-down experiments (Fig. 5a, d). In contrast, XPC is captured by the immobilized PAR-binding module, but we could not detect UV-induced PARylation of XPC following immunoprecipitation (Fig. 5c). This means that either the capture of XPC at the immobilized PAR-binding module is mediated indirectly through its interaction with PARP1, which is heavily PARylated after UV (Fig. 5a). Alternatively, XPC may undergo a conformational change at sites of DNA damage that results in the exposure of its PARylated residues and its UVinduced capture at the immobilized PAR-binding module. Either way, our data indicate that the PARP1-XPC complex and chromatin remodeler ALC1 become robustly PARylated at sites of UV-induced DNA damage. PARP1 and PARP2 protect against UV-induced DNA damage. To establish the relevance of PARP1 and PARP2 in UV damage repair, we first assayed PARP1-KO and PARP2-KO cells for UV sensitivity. Western blot analysis confirmed the knockout of PARP1 or PARP2 using specific antibodies (Fig. 6a). Clonogenic survival assays showed that KO of either PARP1 or PARP2 conferred sensitivity to UV irradiation in human cells (Fig. 6b). The siRNA-mediated knockdown of XPC, which was confirmed by western blot analysis, caused similar sensitivities in wild-type and PARP-deficient cells, suggesting that the PARP enzymes cause UV sensitivity mainly through the GGR pathway (Supplementary Fig. 3b, c). To further validate these findings, we Fig. 4 XPC stimulates the poly-(ADP-ribose) response at UV lesions. a Representative images and bquantification of poly-(ADP-ribose) (PAR) levels 10 minutes after local UV-C irradiation (30 J/m2) by immunofluorescence (Trevigen, 4335-MC-100) in the indicated cells. Quantification of DDB2 levels is shown in Fig. S2a. >100 cells were analyzed per condition from three independent experiments. cRepresentative images and dquantification of poly-(ADPribose) (PAR) levels 5 minutes after UV-C irradiation (20 J/m2) by immunofluorescence (Millipore; MABE1031) in the indicated cells. >75 cells were analyzed per condition from three independent experiments. Additional representative images are found in Fig. S2b. eRepresentative images and fquantification of poly-(ADP-ribose) (PAR) levels 10 minutes after local UV-C irradiation (30 J/m2) by immunofluorescence (Trevigen, 4335-MC-100) in in the indicated cells. Quantification of DDB2 levels is shown in Fig. S2c. >100 cells were analyzed per condition from 3 independent experiments. gRepresentative images and hquantification of poly-(ADP-ribose) (PAR) levels 10 minutes after local UV-C irradiation (30 J/m2) by immunofluorescence (Trevigen, 4335-MC-100) in the indicated cells. Quantification of XPC levels is shown in Fig. S2d. >100 cells were analyzed per condition from 3 independent experiments. iRepresentative images and jquantification of poly-(ADP-ribose) (PAR) levels 5 minutes after UV-C irradiation (20 J/m2)by immunofluorescence (Millipore; MABE1031) in the indicated cells. >65 cells were analyzed per condition from four independent experiments. b,d,f,h,jAll cells are depicted as individual data points (gray). The median of each biological replicate is depicted as a colored point, while the bar represents the median of all data points. kRepresentative images and lquantification of the PARylation assay in which recombinant PARP1 (1 pmol) together with NAD was incubated with recombinant XPC-RAD23B (0, 0.25, 0.5, and 1 pmol) for 5 min after which UV-irradiated plasmid was added to the mixture for 30 min. The reaction was stopped and PARylation of PARP1 was monitored. The colored points represent the individual quantification from three independent experiments. The bar represents the median of all data points. The scale bar in a,c,e,g,iis 5 µm. NATURE COMMUNICATIONS | https://doi.org/10.1038/s41467-022-31820-4 ARTICLE NATURE COMMUNICATIONS | (2022) 13:4762 | https://doi.org/10.1038/s41467-022-31820-4 | www.nature.com /naturecommunications 7 ef a UV Ctrl + PARPi XPC ALC1 PARP1PARP2 -+-+-+-+-+ -+ LacR-macro 0 1 2 3 Accumulation at LacR No UV 1 min PARPi LacR macro LacR control XPC GFP XPC GFP LacR macro XPC GFP - 100 - 150 kDa - 250 PARP1GFP PAR Input UV - + - + - 100 - 150 - 250 IP: GFP WT kDa GFPPARP2 PAR Input UV-+ -+ IP: GFP - 100 - 150 - 250 - 100 - 150 - 250 WT dWT kDa GFPALC1 PAR Input UV -+ -+ IP: GFP - 100 - 150 - 250 - 100 - 150 - 250 c kDa XPCGFP PAR Input UV -+ -+ - 100 - 150 - 250 IP: GFP - 100 - 150 - 250 WT b Fig. 5 PARP1 and ALC1 are PARylated in response to UV. a–dImmunoprecipitation of aPARP1-GFP, bGFP-PARP2, cXPC-GFP, dGFP-ALC1 under highsalt conditions in the presence and absence of UV-C (20 J/m2, 15 min) stained for PAR (Millipore; MABE1016) or GFP. Three independent replicates of each IP experiment were performed obtaining similar results. eRepresentative images and fquantification of GFP-tagged XPC, ALC1, PARP1, or PARP2 recruitment to the LacO array upon tethering to the indicated mCherry-LacR-macrodomain. Pictures were taken before and 1 min after UV-C microirradiation. 24–45 nuclei were analyzed in three independent biological replicates (n=3). Additional representative images are found in Fig. S3a. The scale bar in eis 5 µm. ARTICLE NATURE COMMUNICATIONS | https://doi.org/10.1038/s41467-022-31820-4 8NATURE COMMUNICATIONS | (2022) 13:4762 | https://doi.org/10.1038/s41467-022-31820-4 | www.nature.com/naturecommunications performed CPD dot blot assays in PARP1-KO and PARP2-KO cells. To this end, genomic DNA was isolated from cells at varying time points after UV-C irradiation to determine the amount of remaining UV-induced photoproduct. This approach revealed that both PARP1-KO and PARP2-KO cells displayed delayed CPD repair (Supplementary Fig. 3d, e). To extend the biological relevance of these findings, we asked whether the role of PARP1 and PARP2 in protecting against UV irradiation is evolutionarily conserved in an animal model. We obtained PARP1-deficient (ok988) and PARP2-deficient (ok344) C. elegans (Fig. 6c) and performed germ cell and embryo survival assays after UV irradiation, which specifically monitor GGR46. Deletion of either PARP1 or PARP2 strongly sensitized nematodes to UV-B light compared to WT animals (Fig. 6d). Interestingly, animals containing a double knockout for XPC (tm3886) and either PARP1 (ok988) or PARP2 (ok344), were as sensitive to UV-B irradiation as single XPC-deficient nematodes (Fig. 6e). These findings indicate evolutionary conservation of the involvement of both PARP enzymes in GGR. ALC1 is recruited to UV lesions by XPC. Our proteomics analyses revealed that XPC binds more robustly to PARP2 than PARP1, and that ALC1 is an abundant interactor of PARP2 (Fig. 2). Moreover, ALC1 was robustly PARylated in response to UV irradiation (Fig. 5d). To better understand the links between XPC, PARP enzymes, poly-(ADP-ribose)-responses and ALC1 in GGR, we profiled the ALC1 interactome, dissected the role of its PAR-binding macrodomain and ATPase activity, as well as measured the UV-C-dependent recruitment dynamics of ALC1. We, therefore, generated ALC1-KO cells in U2OS (FRT) and stably re-expressed GFP-tagged versions of ALC1 (WT, ATPasedead; E175Q, and PAR-binding-deficient; Δmacrodomain; Supplementary Fig. 4a). Label-free proteomics after GFP-ALC1 pulldown confirmed a strong interaction of wild-type ALC1 with PARP2, PARP1, the FACT subunit SPT16 and core histones (Fig. 7a). In contrast, the ATPase-dead version of ALC1 interacted less with PARP2, and core histones, suggesting that the ATPase activity of ALC1 impacts the association of the enzyme with PARP2 (Fig. 7b). Consistently, immunoprecipitation experiments confirmed that ALC1 robustly bound PARP2 and to a lesser extent PARP1, and that these interactions were decreased with ALC1 E175Q (Fig. 7c). Moreover, an ALC1 Δmacrodomain mutant showed a completely disrupted interaction with PARP1/2. Next, we tested the recruitment of ALC1 to UV-C DNA damage sites. Local UV-C laser irradiation experiments showed a PARP2-KO 0246 10 100 UV-C (J/m2) Surviving colonies (%) WT PARP1-KO b 0 40 80 120 160 1 10 100 UV-B (J/m ) 2 Survival C. elegans (%) c WT PARP1 -KO PARP2 -KO PARP1 PARP2 Tubulin - 55 - 70 - 130 kDa ok988 2000 bp parp-1 ok344 parp-2 200 bp d 0 60 120 1 10 100 UV-B (J/m ) 2 Survival C. elegans (%) WT parp-2 (ok344); xpc-1 (tm3886) parp-1 (ok988); xpc-1 (tm3886) xpc-1 (tm3886) e WT parp-2 (ok344) parp-1 (ok988) xpc-1 (tm3886) 20 40 80 100 Fig. 6 PARP1 and PARP2 impact the repair of CPD lesions. a Western blot of U2OS WT, PARP1-KO and PARP2-KO cells. bClonogenic survival assays of U2OS WT, PARP1-KO, and PARP2-KO cells upon UV-C irradiation. The data is depicted as mean +S.E.M. from n=4 independent experiments, except for PARP1-KO for which n=3. cRepresentation of the deletions in the C. elegans parp-1 (ok988) and parp-2 (ok344) strains. dGerm cell and embryo UV survival assays of PARP1-deficient parp-1(ok988) and PARP2-deficient parp-2(ok344) C. elegans.eGerm cell and embryo UV survival assays of XPCdeficient xpc-1 (tm3886), XPC and PARP1 double deficient (ok988; tm3886), or XPC and PARP2 double deficient (ok344; tm3886) C. elegans.d,eThe data are depicted as mean +S.E.M. from three independent experiments. NATURE COMMUNICATIONS | https://doi.org/10.1038/s41467-022-31820-4 ARTICLE NATURE COMMUNICATIONS | (2022) 13:4762 | https://doi.org/10.1038/s41467-022-31820-4 | www.nature.com /naturecommunications 9 UV-B tubes (40 W). Following 24 h recovery on OP50 E. coli culture plates, three adult animals were allowed to lay eggs for 4 h on 6 cm plates seeded with HT115 bacteria, in quintuple for each UV-B dose. The number of hatched and unhatched (dead) eggs was counted 24 h later and the survival percentage was calculated. Results are plotted as the average of three independent experiments. LacO-LacR system for detecting PARylated proteins. U2OS 2-6-3 cells containing 200 copies of a LacO-containing cassette were plated on an 18-mm glass coverslip. The next day the cells were co-transfected with lipofectamine 2000 (Invitrogen) and plasmid DNA for 6 h at 37 °C. Next the medium was replaced with DMEM +/+and incubated overnight at 37 °C. Prior to the UV-C microirradiation, the medium was replaced with CO 2 -independent Leibovitz L15 medium (Thermo Fisher Scientific) and cells were incubated with 10 µM of PARG inhibitor (Sigma) for 30 min. If indicated the cells were additionally incubated with 10 µM Olaparib. UV-C laser tracks were made using a diode-pumped solid-state 266-nm Yttrium Aluminum Garnet laser (average power 5 mW, repetition rate up to 10 kHz, and pulse length 1 ns). The UV-C laser is integrated into a UGA-42Caliburn/2 L Spot Illumination system (Rapp OptoElectronic). Micro-irradiation was combined with live-cell imaging in an environmental chamber set to 37 °C on an all-quartz widefield fluorescence Zeiss Axio Observer 7 microscope, using a ×100 (1.2 NA) ultrafluar glycerol-immersion objective (UV-C). The laser system is coupled to the microscope via a TriggerBox, and a neutral density (ND-1) filter blocks 90% of the laser light. An HXP 120-V metal-halide lamp was used for excitation. Images were acquired in Zeiss ZEN and quantified in ImageJ. Immunoblot. Immuno-dot and immuno-slotblot assays were performed as previously described61. DNA was extracted using DNeasy Blood & Tissue Kit (Qiagen 69504). DNA (300 ng per well, two to three technical replicates per sample) was vacuum-transferred to a nitrocellulose membrane using the Bio-Dot or Bio-Dot-SF apparatus (Bio-Rad, 1706542/5). Membranes were baked at 80 oC in a Bio-Rad’s Gel Dryer model 583, blocked in 5% milk in PBS with 0.1% Tween (PBST), washed three times in PBST, and incubated with 6-4PP or CPD antibodies (see Supplementary Table 5) overnight at 4 oC. Membranes were again washed in PBST and incubated with HRP-conjugated anti-mouse antibodies (ECL Mouse IgG, HRPlinked whole Ab (from sheep), Cytiva, NA931). Damage signal was detected using enhanced chemiluminescence (ECL™Prime Western Blotting System, GE Healthcare, RPN2232) and exposure in the Bio-Rad ChemiDocTM XRS +imaging system. Genomic DNA amount loaded onto the membrane was quantified using SYBR™Gold Nucleic Acid Gel Stain (1:5000 dilution, Invitrogen, S11494), by incubating the membrane with SYBR-Gold solution for 60 min, followed by three washes with PBST. Damage signal was normalized to SYBR-Gold signal using Image Lab version 6.0 from Bio-Rad. Immunofluorescence microscopy. Immunofluorescence staining was performed as described previously47. In brief, cells were fixed with 2% paraformaldehyde + 0.1% Triton X-100 for 15 min at room temperature, washed 3× with PBS +0.1% Triton X-100 and subsequently permeabilized with PBS +0.1% Triton X-100 for 2×10 min at room temperature. After blocking the cells in PBS +(PBS +0.5% BSA +0.15% glycine), cells were incubated with the primary antibody (see Supplementary Table 5), diluted in PBS +, overnight at 4 °C. Unspecific antibody staining was removed by washing the cells 5× in PBS +0.1% Triton X-100. Subsequently, cells were stained with Alexa Fluor 488/568-conjugated fluorescent secondary antibody (Thermo Fisher), diluted 1:500 in PBS+, for 1 h at room temperature. Finally, cells were washed 5× with PBS +0.1% Triton X-100, stained with Hoechst 33342 (Fisher Scientific, 1:5000 in PBS) for 10 min, and washed 3× in PBS. The immunofluorescence intensities were measured on a Zeiss Axio Observer Z1 confocal spinning-disk microscope equipped with an sCMOS ORCA Flash 4.0 camera (Hamamatsu), using a Plan-Apochromat ×40/0.95-KOrr air objective or a ×40 C-Apochromat/1.2-KOrr water objective. Nuclear PAR levels. Cells were grown for 24 h in 96 well SCREENSTAR plates (Greiner Bio-One) and incubated for 10 min with 1 µM of PARG inhibitor (PDD 00017273, Sigma) before irradiation with 20 J/m2UV-C light (Stratalinker 1800, Agilent Genomics). After UV-C treatment, the cells were incubated for 5 min at 37 °C in the presence of PARG inhibitor, and subsequently fixed and stained for immunofluorescence as described above. Nuclear PAR levels were quantified in ImageJ by thresholding cell nuclei using the Hoechst signal and subsequently measuring the mean fluorescent signal of poly-(ADP-ribose) in the nucleus. PAR levels at local UV damage. Cells were seeded on 18-mm glass coverslips in 12-wells plates in DMEM with 1% FBS. After 24 h, the medium was replaced with CO 2 -independent Leibovitz L15 medium (Thermo Fisher Scientific) and cells were incubated with 10 µM of PARG inhibitor (PDD 00017273, Sigma) for 10 min. Subsequently, cells were locally irradiated with 30 J/m2UV-C (TUV PL-S 9 W; Philips) through a polycarbonate mask with pores of 5 µm (Millipore) as described62. After UV-C treatment, the cells were incubated for 5, 10, or 20 min at 37 °C in the presence of PARG inhibitor, and subsequently fixed and stained for immunofluorescence as described above. The mean fluorescent signal of PAR levels at sites of local damage was quantified in ImageJ by thresholding cell nuclei using the DAPI signal and thresholding sites of local damage by using the signal of either DDB2 or XPC. XPC immunofluorescence at local UV-C lesions. Cells were grown on coverslips (Fisher Scientific). Before irradiation, the coverslips were covered by 5 µm nanopore filters (Millipore) to allow local UV-C irradiation. The cells were then irradiated with 100 J/m2UV-C light (Stratalinker, Agilent Genomics) and fixed 10 min after irradiation. The immunofluorescence was essentially done as described above, with the addition of a denaturation step with 0.07 M NaOH in PBS for 5 min and a second blocking step in PBS +before incubation with the primary antibody, to allow recognition of CPD lesions. The enrichment of XPC at CPD lesions was quantified in ImageJ. Nuclei and CPD lesions were recognized by thresholding the Hoechst and CPD signal, respectively. The mean fluorescent intensity of XPC and CPD was measured at CPD spots and in the rest of the nucleus. The enrichment of XPC/CPD at UV-C lesions was quantified as followed: mean fluorescence(spot)/ mean fluorescence(nucleus background)−1. Proteins. Recombinant XPC-RAD23B was expressed in Sf9 insect cells and purified as described previously63. XPC has 3xFLAG tag at the N-terminus. In vitro PARylation. In vitro PARylation was performed in 10 μlvolumeat 25 °C for 30 min as described previously64.ThePARylationmixturecontained buffer (100 mM Tris-HCl pH8.0, 10 mM MgCl2, 10% glycerol, 1.5 mM DTT, 100 μg/ml BSA), 20 μM NAD and 1 pmole of PARP1 protein. XPC-RAD23B was added at 0, 0.25, 0.5, and 1 pmole for every pmole of PARP1. The tubes containing the above ingredients were incubatedat25°Cfor 5min.Following this, 0.5 pmoles of plasmid DNA irradiated with 5000 J/m2of UV-C was added to each tube and they were incubated further for 30 min. The reaction was stopped by adding an equal volume of 2× Laemmli buffer and the samples were separated on denaturing 6% SDS-PAGE. The gel was transferred on nitrocellulose and probed for PAR after which blots were additionally proved for PARP1 and XPC. 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The PRIDE database and related tools and resources in 2019: improving support for quantification data. Nucleic Acids Res. 47, D442–D450 (2019). Acknowledgements The authors acknowledge Sylvie Noordermeer for the PARP inhibitor olaparib. We thank Artur Mayerhofer for kindly providing the Stratalinker UV device for irradiation. We thank Nicholas Lakin for providing PARP1-KO and PARP2-KO cells. We thank Susan Janicki for providing U2OS 2-6-3 cells. C. elegans strains were provided by the Caenorhabditis Genetics Center (funded by NIH Office of Research Infrastructure NATURE COMMUNICATIONS | https://doi.org/10.1038/s41467-022-31820-4 ARTICLE NATURE COMMUNICATIONS | (2022) 13:4762 | https://doi.org/10.1038/s41467-022-31820-4 | www.nature.com /naturecommunications 17 Programs P40 OD010440). We thank Masanao Miwa (National Cancer Center Research Institute, Tokyo) for providing 10H hybridoma cells obtained through the Riken cell bank. We thank Mihaela Robu for feedback on the manuscript. This research was supported by an LUMC Research Fellowship, ENW-M (OCENW.KLEIN.090), and ALW-VIDI grants (ALW.016.161.320) from the Dutch Research Council (NWO) to M.S.L. This research was further funded by the DFG (German Research Foundation) through Project-ID 213249687 - SFB 1064 and Project-ID 325871075 - SFB 1309, as well as LMU to A.G.L. C.B. was the recipient of a grant from the Stiftungskommission of the LMU Medical Faculty. R.G-P was the recipient of a Young Investigator Grant from the Dutch Cancer Society (KWF-YIG 11367). A.C.O.V. was funded by an ERC grant (310913). H.L. and M.v.d.W. were funded by the Netherlands Organization for Scientific Research (711.018.007 and CancerGenomiCs.nl) and the Oncode Institute, which is partly financed by the Dutch Cancer Society. O.D.S. was supported by the Korean Institute of Basic Science (IBS-R022-A1) and the National Cancer Insitute (USA, P01CA092584). G.M.S. was supported by grants from the Research Centre of CHU de Quebec Laval University as well as from the Natural Sciences and Engineering Research Council of Canada through the Discovery Grant (RGPIN-2016-05868) and the Discovery Accelerator Supplement Grant (RGPAS-492875-2016). S.A. was funded by the Israel Cancer Research Fund Research Career Development Award (3013004741), the Israel Cancer Association grant (20210078), and Israel Science Foundation grant (1710/17) administered by the Israeli Academy of Science and Humanities and is the recipient of the Jacob and Lena Joels memorial senior lectureship. H.v.A. was funded by a VICI grant from the Dutch Research Council (NWO-VICI grant VI.C.182.052). Author contributions C.B. generated plasmids, U2OS Flp-In cell lines, performed clonogenic survivals, co-IP experiments, western blot analyses, UV-C laser micro-irradiation experiments, local damage, and nuclear PAR immunofluorescence stainings. KA generated and validated U2OS singleKO cells, generated plasmids and U2OS Flp-In cell lines, performed co-IP experiments for western blot and mass spectrometry, UDS experiments, western blot analyses, UV-C laser micro-irradiation, LacR-based PARylation assays and prepared genomic DNA for immunoblot experiments. D.v.d.H. performed all PAR immunofluorescence at sites of local damage, C.G-L., T.T.K., and J.C. performed denaturing IP experiments, M.B.R. and H.v.A. performed chromatin expansion assays, M.v.d.W. and H.L. performed GGR sensitivity assays in C. elegans. R.G-P analyzed the mass spectrometry samples with support from A.C.O.V. A.Y., A.P., S.A. performed and analyzed all immunoblot experiments. D.E.C.B. performed clonogenic survivals and western blot analyses. A.K. performed RRS experiments and performed clonogenic Illudin S survivals. R.G.S. and G.M.S. performed the in vitro PARylation assay. H.S.K. and O.D.S. provided recombinant XPC-RAD23B complex. A.G.L. and M.S.L. conceived, coordinated, and supervised the project. C.B. and M.S.L. drafted the manuscript. All authors commented and edited the manuscript. Competing interests A.G.L. is a founder, shareholder, and managing director of Eisbach Bio, a biotech developing small molecule therapeutics. The remaining authors declare no competing interests. Additional information Supplementary information The online version contains supplementary material available at https://doi.org/10.1038/s41467-022-31820-4. Correspondence and requests for materials should be addressed to Andreas G. Ladurner or Martijn S. Luijsterburg. Peer review information Nature Communications thanks Brendan Price and the other, anonymous, reviewer(s) for their contribution to the peer review of this work. Peer reviewer reports are available. 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