# PARP1 auto-modification promotes faithful Okazaki fragment processing and limits replication fork speed

Jonas D Elsborg, Sebastian H N Munk, Alba Adelantado-Rubio, Ramóna Pék, Zita Fábian, Victor Imburchia, Siham Zentout, Ivo A Hendriks, Sara C Buch-Larsen, Rebecca Smith, et al.

► To cite this version:

Jonas D Elsborg, Sebastian H N Munk, Alba Adelantado-Rubio, Ramóna Pék, Zita Fábian, et al. PARP1 auto-modification promotes faithful Okazaki fragment processing and limits replication fork speed. Molecular Cell, 2025, 85 (19), pp.3562-3575.e10. 10.1016/j.molcel.2025.09.006. hal-05306029

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# Molecular Cell

## PARP1 auto-modification promotes faithful Okazaki fragment processing and limits replication fork speed

**Graphical abstract**

<figure><img src="image_2.png" /></figure>

**Authors**

Jonas D. Elsborg, Sebastian H. N. Munk, Alba Adelantado-Rubio, ..., Sébastien Huet, Apolinar Maya-Mendoza, Michael L. Nielsen

**Correspondence**

apomm@cancer.dk (A.M.-M.), michael.lund.nielsen@cpr.ku.dk (M.L.N.)

**In brief**

Using a separation-of-function mutant, Elsborg et al. explore the distinct roles of PARP1 auto-modification and trans-ADP-ribosylation. Auto-modification regulates DNA replication fork progression, prevents replication stress, and is required for PARP1 eviction from DNA-damage sites. Its loss is synthetic lethal with FEN1 inhibition, highlighting a key role in Okazaki fragment processing.

**Highlights**

* Mutation of four serines generates an auto-modification-deficient PARP1

* AM drives PARP1 eviction from DNA breaks but not repair factor recruitment

* AM controls replication fork speed and Okazaki fragment maturation

* Proteomics maps the ADP-ribosylation network at the replication fork

<figure><img src="image_3.png" /></figure>
## Article

# PARP1 auto-modification promotes faithful Okazaki fragment processing and limits replication fork speed

Jonas D. Elsborg,<fnref n="1" />,<fnref n="7" /> Sebastian H.N. Munk,<fnref n="2" /> Alba Adelantado-Rubio,<fnref n="2" /> Ramóna Pék,<fnref n="3" /> Zita Fábian,<fnref n="4" /> Victor Imburchia,<fnref n="3" /> Siham Zentout,<fnref n="3" /> Ivo A. Hendriks,<fnref n="1" /> Sara C. Buch-Larsen,<fnref n="1" /> Rebecca Smith,<fnref n="3" />,<fnref n="7" /> Jiri Bartek,<fnref n="5" /> Julien P. Duxin,<fnref n="4" />,<fnref n="6" /> Sébastien Huet,<fnref n="3" /> Apolinar Maya-Mendoza,<fnref n="2" />,\* and Michael L. Nielsen<fnref n="1" />,<fnref n="8" />,\*

## SUMMARY

Poly(ADP-ribose) polymerase (PARP) inhibitors have proven their efficacy for treating tumors defective in homologous recombination via synthetic lethality. In response to DNA breaks, PARP1 is the primary ADP-ribosylation writer, modifying itself (auto-modification) and other proteins to facilitate repair. However, enzymatic inhibition blocks both processes, making it difficult to dissect their distinct functional roles. Using proteomics and site-directed mutagenesis, we identified a PARP1 mutant deficient in auto-modification, yet it retains catalytic activity. This separation-of-function mutant revealed that PARP1 auto-modification slows DNA replication fork progression but is dispensable for repair factor recruitment. Instead, auto-modification promotes the timely release of PARP1 at DNA break sites and prevents the formation of replication stress. Simultaneous inhibition of FEN1 and loss of PARP1 auto-modification gives rise to synthetic lethality, implicating auto-modification in Okazaki fragment processing. Our results demonstrate that trapping of PARP at DNA breaks impedes repair factor accessibility, constituting an important dimension of PARP-inhibitor-driven cytotoxicity.

## INTRODUCTION

Cells defective in homologous recombination (HR) are hypersensitive to chemical inhibition of poly(ADP-ribose) polymerases (PARPs) 1/2,<fnref n="1" />,<fnref n="2" /> which suppresses their catalytic activity. Accordingly, PARP inhibitors (PARPi) have been developed as cancer therapies.<fnref n="3" /> PARPs catalyze ADP-ribosylation (ADPr), transferring ADP-ribose from NAD⁺ to target proteins as a post-translational modification (PTM). In the context of DNA repair, PARP1 binds various DNA lesions, including single-strand gaps/breaks (SSBs).<fnref n="4" />⁻<fnref n="6" /> This triggers a conformational change that promotes PARP1 activation<fnref n="7" />,<fnref n="8" /> and forms a composite active site with histone PARylation factor 1 (HPF1).⁹⁻¹³ Activated PARP1 catalyzes ADPr onto itself (auto-modification), histones, and other substrates,<fnref n="4" />⁻<fnref n="6" />,¹⁴ with auto-modification promoting the release of PARP1 from chromatin, after which the enzyme reverts to its inactive state.¹⁰,¹⁵,¹⁶

Accordingly, inhibition of PARP1 catalysis prevents auto-modification, inducing PARP trapping by retention at chromatin,¹⁷⁻¹⁹ and it also blocks trans-modification, disrupting repair factor recruitment and chromatin remodeling.<fnref n="4" />,²⁰⁻²² Catalytic activity is thus essential for recruitment of DNA repair factors such as XRCC1 and LIG3.²³ Beyond repair, PARP1 has roles in DNA replication. It senses single-stranded DNA (ssDNA) gaps on the lagging strand during Okazaki fragment processing (OFP), and PARPi prevents maturation of nascent DNA.²⁴⁻²⁶ PARPi also accelerates fork progression, causing replication stress,²⁷,²⁸ suggesting multiple roles for ADPr during S-phase.

Mechanistically, PARP1 is thought to act via ADPr-dependent recruitment of repair factors to its auto-modification.<fnref n="4" />,<fnref n="6" />,²⁹,³⁰ Whether this also applies during replication is unclear. A key challenge to understanding the functional relevance of PARP1 ADPr lies in the difficulty in distinguishing the contributions of self and non-self ADPr during these processes. As PARPi

<footnote n="1">¹Department of Cellular and Molecular Medicine, Novo Nordisk Foundation Center for Protein Research, Proteomics Program, Faculty of Health and Medical Sciences, University of Copenhagen, Copenhagen, Denmark</footnote>

<footnote n="2">²DNA Replication and Cancer Group, Danish Cancer Institute, Copenhagen, Denmark</footnote>

<footnote n="3">³Univ Rennes, CNRS, IGDR (Institut de génétique et développement de Rennes) - UMR 6290, BIOSIT, UMS3480, 35000 Rennes, France</footnote>

<footnote n="4">⁴Department of Cellular and Molecular Medicine, Novo Nordisk Foundation Center for Protein Research, Protein Signaling Program, Faculty of Health and Medical Sciences, University of Copenhagen, Copenhagen, Denmark</footnote>

<footnote n="5">⁵Genome Integrity Group, Danish Cancer Institute, Copenhagen, Denmark</footnote>

<footnote n="6">⁶Biotech Research and Innovation Centre, Faculty of Health and Medical Sciences, University of Copenhagen, Copenhagen, Denmark</footnote>

<footnote n="7">⁷Present address: Sir William Dunn School of Pathology, University of Oxford, Oxford, UK</footnote>

<footnote n="8">⁸Lead contact</footnote>

<footnote>\*Correspondence: apomm@cancer.dk (A.M.-M.), michael.lund.nielsen@cpr.ku.dk (M.L.N.)
https://doi.org/10.1016/j.molcel.2025.09.006</footnote>
A PARP1

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B

Overlap (ADPr sites)

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C

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D

Treatment

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E

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F

Characterization of PARP1 auto-modification by MS

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G

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H

PARP1 auto-modification
(all ADPr peptides)

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I

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J

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K

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prevents both auto-modification and *trans*-modification, it obscures the mechanism by which they induce cytotoxicity. To address this, we here describe a PARP1 mutant defective in auto-modification but proficient for *trans*-ADPr.

This separation-of-function mutant revealed that auto-modification is dispensable for repair factor recruitment but required for alternative OFP and for eviction of PARP1 at DNA breaks. We also show that auto-modification restricts replication fork speed and map ADPr targets at replication forks, identifying factors involved in OFP.

## RESULTS

### Site-specific characterization of ADPr reveals a novel fourth site within the PARP1 auto-modification loop

Our previous work identified several ADPr sites on activated PARP1, with auto-modification primarily occurring at three major sites: S499, S507, and S519.<sup>14</sup> To explore its regulatory role, we generated U2OS PARP1 knockout (KO) cell lines expressing GFP-tagged PARP1 wild type (WT) or a mutant with these three serines replaced by alanine (3SA) (Figure 1A). We activated PARP1 by treating cells with methyl methanesulfonate (MMS) or H<sub>2</sub>O<sub>2</sub><sup>31</sup> and then used our mass spectrometry (MS)-based Af1521-enrichment strategy to map ADPr at a site-specific level using electron-transfer dissociation (ETD)-based MS<sup>32</sup> (Figure S1A). We identified 3,621 ADPr sites on 1,203 proteins with high reproducibility (Figure S1B; Table S1), predominantly on serines, consistent with PARP1-HPF1 activity after DNA damage<sup>9,33</sup> (Figure S1C). PARP1 WT and 3SA mutants modified the same proteins, with near-complete overlap between sites (Figure 1C), including across treatments (Figure S1D).

Next, we asked which ADPr sites would be differentially regulated between PARP1 WT and 3SA mutant cells. In untreated cells where PARP1 activation is low, no sites were significantly regulated (Figure S1E). Following DNA damage, the three canonical serines were ADPr-modified only in WT, whereas in 3SA cells, a fourth serine, S504, became abundantly modified (Figure 1C). Since S504 ADPr was mutually exclusive with the canonical sites, this implies multi-site auto-modification on PARP1, with several modified serine residues coexisting simultaneously

on the same molecule (Figure 1D). The auto-modification loop (residues 483–530<sup>34</sup>) contains exactly four serines, all modified in either WT or 3SA cells. Therefore, we additionally mutated this fourth serine to alanine (4SA) to generate an auto-modification-deficient PARP1. Transient expression and immunoprecipitation of PARP1 confirmed robust suppression of auto-modification in 4SA compared with WT following DNA damage (Figure 1E).

Recently, the chemical stability of ester-linked modifications, such as Asp/Glu-linked ADPr, has been brought into question.<sup>35</sup> Since these chemical bonds may undergo hydrolysis at alkaline pH and with heating,<sup>36</sup> our Af1521-based enrichment strategy may underrepresent their relative abundance, as the protocol includes steps with such conditions. To characterize PARP1 auto-modification under conditions where all linkage types are stable, we additionally created stable PARP1 4SA cell lines and performed proteomic analysis using a modified protocol from Longarini and Matić,<sup>35</sup> which preserves ester-linked ADPr. We compared PARP1 WT, 4SA, and *HPF1* KO cells under DNA-damage conditions, as well as *in vitro* purified PARP1 variants (WT and E988K MARylation-specific mutant<sup>37</sup>) supplemented with HPF1 (Figure 1F).

From highly purified PARP1 (~95%), we identified 56 high-confidence ADPr sites after applying stringent filtering (Table S2), the highest number reported by any study, including Longarini and Matić<sup>35</sup> (~40 sites). We confirm that this improved method preserves ester-linked ADPr during MS analysis, with high-quality spectra localizing ADPr to glutamate in every condition (Figure 1G). However, while Glu-ADPr was stabilized, their relative contribution toward PARP1 auto-modification was low when compared with Ser-ADPr in HPF1-proficient samples. Our analysis corroborates that HPF1 is essential for directing PARP1 specificity toward Ser, in accordance with previous reports.<sup>9,11,12</sup> Short PARG treatments minimally affected PARP1 ADPr, indicating prevalent MARylation, but in E988K samples, PARG reduced glutamate- but not serine-linked ADPr (Figure 1G). This observation corroborates a recent report finding PARG to exhibit some hydrolase activity toward Asp and Glu MARylation.<sup>35</sup> Next, we wanted to directly quantify ADPr levels on PARP1 when all linkage types are preserved.

Figure 1. Site-specific characterization of ADPr reveals a novel fourth site within the PARP1 auto-modification loop

(A) Schematic of PARP1 domains.

(B) Scaled Venn diagram depicting the overlap between the ADPr-sites found in PARP1 WT and 3SA mutant cells.

(C) Volcano plot of ADPr sites enriched in PARP1 WT cells and 3SA mutant cells in response to H<sub>2</sub>O<sub>2</sub> exposure.

(D) Mean ADPr-site intensity of PARP1 serine residues 499, 504, 507, and 519 quantified across experimental conditions (n = 4).

(E) Western blot analysis following GFP-PARP1 immunoprecipitation in U2OS *PARP1* KO cells transiently expressing GFP-PARP1 WT or 4SA-mutant. Cells were treated using 0.24 mM MMS for 30 min.

(F) Experimental overview for mapping PARP1 auto-modification sites using ETD-based MS while preserving ester-linked ADPr.

(G) PARP1 ADPr-peptide intensity (scaled to median PARP1 expression) per experimental condition, stratified by PARG-treatment. Barplots reflect the mean intensity quantified within replicate conditions, stacked by their amino acid acceptor sites. The number of sites found within each condition is marked below the bars.

(H) Quantification of the relative PARP1 auto-modification levels for WT and 4SA mutant proteins. The graph shows the conditions where PARG was omitted during sample preparation. We allowed any ADPr peptide for quantification regardless of its localization score for peptide-resolution mapping. Significance was determined using Welch's t test, *p* < 0.001, *n* = 4.

(I) Top: the ADPr-peptide abundance is plotted against the amino acid positions for each ADPr site across the amino acid sequence of PARP1. Each data point represents the total intensity of a unique peptide identified for that condition. Bottom: the relative ADPr site density is displayed in red (frequency polygon, bin size = 25, area under each curve sums up to 1). The blue box annotates the auto-modification loop within the PARP1 sequence.

(J) AlphaFold model of PARP1, labeled with the auto-modification sites within the functional loop, including annotation of other domains.

(K) Volcano plot analysis of ADPr sites quantified in PARP1 4SA cells in response to 0.24 mM MMS treatment for 30 min.
<figure><img src="image_16.png" /><figcaption>B</figcaption></figure>

<figure><img src="image_17.png" /><figcaption>C</figcaption></figure>

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Quantification of PARP1-dependent repair factors

<figure><img src="image_21.png" /><figcaption>G</figcaption></figure>

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(4SA, recomb.)</figcaption></figure>

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Genotype

U2OS PARP1 KO + GFP-XRCC1

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UV damage (20J)
60 min recovery

(legend on next page)
We found that ADPr of the PARP1 4SA mutant was significantly reduced by ~99% compared with WT, confirming that these four mutations confer near-complete loss of auto-modification (Figure 1H).

ADPr acceptor sites preferentially localize within intrinsically disordered regions.¹⁴ Indeed, PARP1 from cells (both WT and *HPF1* KO) was modified almost exclusively within the disordered auto-modification loop (Figure 1I). This striking specificity is supported by structural studies showing that this flexible loop can extend into the PARP1 catalytic site to enable auto-modification in *cis*.¹⁵ Conversely, PARP1 *in vitro* reactions exhibited a completely different modification space, with a high density and intensity of ADPr near the N-terminal Zn fingers. This activity is likely a result of *trans*-modification between two neighboring PARP1 molecules, considering that this domain is far away from the active site.¹⁵ Thus, our data demonstrate that physiological auto-modification of PARP1 is confined to the auto-modification loop and predominantly targets serines under HPF1-proficient conditions (Figure 1J). Finally, global ADPr profiling of PARP1 4SA cells using Af1521 enrichment (Figure S1A) revealed 1,695 high-confidence sites on 739 proteins (Table S3), measured with high reproducibility (Figure S1F). Like 3SA, the 4SA mutant retained catalytic activity (Figure 1K) and targeted the same substrates as WT (Figure S1G), showing auto-modification does not alter substrate specificity.

In summary, we identified S504 as a novel auto-modification site, activated when canonical sites are mutated. Upon mutation of all four serines, the auto-modification of PARP1 is almost entirely lost, yet the catalytic activity and specificity of the enzyme are retained. Thus, this PARP1 variant is uniquely positioned for separating the functional consequences of PARP1 auto-modification from the catalytic activity of the enzyme.

## A specific role for auto-modification in PARP1 eviction and DNA repair

Following its activation, PARP1 is released from chromatin in an auto-modification-dependent manner.<sup>33,38–40</sup> With this in mind,

we asked if the PARP1 4SA mutation would affect its dissociation kinetics from sites of DNA damage. Using laser irradiation to induce DNA damage, we followed the recruitment of GFP-PARP1 WT and 4SA to damaged chromatin by live-cell microscopy (Figure S2A). We observed that PARP1 WT had a remarkably faster rate of dissociation than the 4SA mutant, confirming that PARP1 release is facilitated by its auto-modification (Figure 2A). The observed dissociation delay resembles niraparib inhibition,⁴¹ suggesting a robust suppression of auto-modification.

We next investigated the potential contribution of Glu auto-modification for PARP1 eviction, tracking the recruitment of PARP1 WT and PARP1 6EA mutant in HPF1-proficient and KO cells. The PARP1 6EA mutant lacks the proposed Glu acceptor sites identified in HPF1 KO cells,³⁵ which we also located within the auto-modification loop (Figure 1I). If substantial auto-modification would target glutamates, we would expect this mutant to dissociate at a slower rate than PARP1 WT; however, this was not the case (Figure 2B). Conversely, ablation of serine ADPr by HPF1 KO significantly increased the retention time of PARP1 WT, demonstrating their importance for eviction (Figure 2C). When 6EA release was assessed in HPF1 KO cells, both Ser and Glu-ADPr of PARP1 were prohibited with an additive effect for enhanced trapping. This result shows that Glu auto-modification may functionally contribute to PARP1 eviction, but only under HPF1 KO conditions where synthesis of Ser-ADPr is otherwise not possible.

Because the 4SA mutant experienced prolonged association with damaged chromatin, we asked if this would lead to enhanced activity. Indeed, PARP1 4SA became hyper-activated upon DNA-damage induction, abundantly ADP-ribosylating a wide range of proteins in the cell (Figure 2D). MS quantification confirmed that proteins from PARP1 4SA cells had significantly elevated levels of ADPr (>2-fold), compared with WT following MMS-induced damage (Figure 2E). The bulk of ADPr sites were uniformly upregulated in the PARP1 4SA mutant compared with the WT, demonstrating that this increase was not driven by a few individual sites (Figure S2B).

**Figure 2. A specific role for auto-modification in PARP1 eviction and DNA repair**

(A) Recruitment kinetics of U2OS *PARP1* KO cells transiently expressing GFP-PARP1 WT and 4SA mutant cells by live-cell microscopy following laser microirradiation. The data are normalized to the maximum value (recruitment peak) for each condition, mean ± SEM (n ≥ 14 cells per cell line).

(B) Recruitment kinetics of U2OS cells with single KO (PARP1) or double KO (PARP1 and HPF1) transiently expressing GFP-PARP1 WT and 6EA mutant by live-cell microscopy following laser microirradiation. The data are normalized to the maximum value (recruitment peak) for each condition, mean ± SEM (n ≥ 12 cells per cell line).

(C) Dissipation time for the cells shown in (B), showing the 50% decay of PARP1 signal intensity at the stripe. Significance was determined using the Welch's two-sample t test with Holm correction. *p* value thresholds are denoted as ***p* < 0.01 and ****p* < 0.0001.

(D) Western blot analysis of ADPr and PARP1 in parental U2OS cells and GFP-PARP1 WT and 4SA mutant cell lines. Cells were treated as indicated with 0.24 mM MMS for 30 min.

(E) Abundance of ADPr peptides across WT and 4SA mutant cells, and bars represent means. Samples were left untreated or exposed to 0.24 mM MMS for 30 min. Significance was determined by Welch's two-sample t test, *p* = 0.0019 (n = 4).

(F) Experimental design to identify PARP1-dependent repair factors using *Xenopus laevis* egg extracts by PP-MS.

(G) Heatmap displaying the regulation of selected replication and repair factors across various time points and conditions as indicated on the figure. Protein recruitment to the plasmid is indicated by Z score, and normalization was performed row wise (n = 4).

(H and I) Volcano plot of proteins recruited to the pGAP plasmid by PARP1 WT (H) or 4SA mutant (I) after 10 min of repair.

(J) Recruitment kinetics of mean ± SEM GFP-XRCC1 signal transiently transfected in a *PARP1* KO background. Co-transfected plasmids express PARP1 WT, 4SA mutant, or LW/AA mutant. The XRCC1 recruitment was quantified at its peak following laser microirradiation (30 s), and significance was determined using Welch's t test with Holm correction, *p* < 5% (n ≥ 14 cells per cell line).

(K) Western blot analysis showing S139 phosphorylation of histone H2AX (γH2AX) of the indicated cells following UV radiation (20 J). Cells were left to recover for 60 min following exposure.
As PARP1-dependent ADPr recruits repair factors,⁴ we wondered if this recruitment requires auto-modification. Using the *Xenopus laevis* cell-free system,⁴²,⁴³ we immunodepleted endogenous PARP1 and reintroduced purified human WT, 3SA, or 4SA. Supplementing extracts with a plasmid containing a single DNA lesion (pGAP), DNA repair is triggered, and recruited proteins are quantified by plasmid pull-down MS (PP-MS) after 5 or 10 min (Figure 2F; Table S4). The samples formed distinct clusters with good reproducibility, with the largest separation between intact control plasmids and the pGAP plasmids (Figure S2C), and we observed recruitment of numerous DNA-repair factors to the damaged plasmids (Figure S2D).

To visualize the overarching protein networks recruited to the plasmids, we highlighted the most defining features in a heatmap (Figure 2G). The depletion of *Xenopus* PARP1 was successfully achieved, as it was not detectable by MS following its depletion, while concomitantly we were only able to detect the purified PARP1 variants in the samples where they were reintroduced. We next investigated the PARP1-dependent factors recruited to the damaged plasmids. From this, we detected PARP1 itself, as well as XRCC1 and LIG3, involved in the repair of SSBs.⁴⁴ We also identified PNKP, a protein that modulates phosphorylation of the ends of the DNA backbone to make them compatible with ligation, and POLβ, which is the DNA polymerase extending the resected gap before LIG3-mediated ligation. Finally, we also identified APLF, a histone chaperone that is recruited to sites of DNA damage by an ADPr-dependent mechanism.⁴⁵ It binds to the Ku complex,⁴⁶ of which XRCC5/6 are core components (Figure 2H).

If the auto-modification of PARP1 is required for the recruitment of these DNA-repair factors, we would expect their recruitment to be abrogated in the reactions reconstituted with PARP1 4SA. However, this was not the case, and both 4SA and 3SA were proficient in the recruitment of repair factors (Figures 2I and S2E). On the other hand, recruitment of the MRN complex (which initiates DDR signaling for downstream repair⁴⁷) was specifically lost after addition of PARP1 4SA but not with WT. Since the 4SA mutation impedes PARP1 dissociation from DNA breaks, this suggests that trapping may prevent the loading of MRN at DNA lesions by means of competition, corroborating previous reports.⁴⁸

We also explored the recruitment of XRCC1 by live single-cell imaging coupled with laser irradiation as previously described. In PARP1 KO cells, XRCC1 recruitment to damaged chromatin was low (Figure 2J). Conversely, re-expressing either PARP1 WT or the 4SA mutant led to robust accumulation of XRCC1, demonstrating that its recruitment is contingent on PARP1 catalytic activity but not auto-modification, in agreement with our PP-MS results (Figures 2H and 2I). We also analyzed XRCC1 recruitment in PARP1-KO cells expressing the LW/AA mutant unable to bind HPF1,⁹,⁴⁹ mimicking HPF1 KO conditions leading to loss of serine ADPr and a near-complete suppression of histone ADPr.⁹,⁴⁹,⁵⁰ While XRCC1 was still recruited to damaged chromatin with the PARP1 LW/AA mutation, the intensity was significantly reduced compared with HPF1-proficient conditions (Figure 2J), suggesting ADPr of chromatin (e.g., histones) may play a larger role for DNA-repair factor recruitment than previously appreciated.

DNA-damage signaling was intact in WT and 4SA cells after ionizing radiation, as seen by strong γH2AX after 60 min (Figure 2K). However, 4SA cells showed elevated γH2AX even without damage, implying endogenous stress. Because 4SA impaired MRN recruitment (Figure 2I), we assessed the development of NBS1 phosphorylation. WT and parental U2OS induced NBS1 phosphorylation rapidly after laser irradiation, whereas PARP1 KO and 4SA mutants were delayed (Figures S2F and S2G), supporting that both PARP1 activity and its release from DNA-damage sites may influence the efficiency of DDR signaling.

Collectively, we show that the PARP1 4SA mutation switches the catalytic activity from auto- to trans-modification, with enhanced activity correlating with increased trapping potency. While auto-modification promotes eviction, it is not essential for repair factor recruitment when ADPr of chromatin or other factors can compensate for this loss. Still, impaired release may influence the timing of DDR signaling and increase endogenous DNA damage.

## PARP1 auto-modification modulates the speed of replication fork progression

The global homeostasis of Ser-ADPr is regulated by its writer and eraser enzymes, PARP1-HPF1 and ARH3, respectively,¹²,⁵¹,⁵² with PARP1 activity restricting the speed of the replication fork to prevent replication stress.²⁵,²⁷,²⁸,⁵³ To further probe the role of Ser ADPr in fork regulation, we combined genetic ablation of HPF1 or ARH3 with PARPi treatment and measured replication dynamics using the DNA fiber assay (Figure S3A). This approach, based on the incorporation of nucleotide analogs into nascent DNA, which enables calculation of replication fork speed by measuring the length of these replication tracts.⁵⁴ In parental U2OS cells, PARPi accelerated the replication fork speed (Figure 3A), consistent with previous findings.²⁷ In ARH3 KO cells, however, PARPi did not increase fork speed, likely due to persistently elevated serine ADPr in the absence of its eraser.⁵⁵ Conversely, HPF1 KO cells already displayed faster forks relative to parental cells, and when additionally treated with PARPi, their replication fork speed was unprecedentedly fast. When replication forks progress too fast, replication stress ensues.⁵⁶ Accordingly, we found that HPF1 KO cells treated with PARPi, but not ARH3 KO cells, accumulate γH2AX in S-phase, which persists into G2 (Figure 3B). Of note, replication forks across all cells had comparable stability and did not show signs of breakage or stalling (Figure S3B).

These findings suggest that PARP1 catalytic activity is involved in regulating the speed of the replication fork. Considering that PARP1 4SA mutant cells experienced elevated levels of γH2AX signaling compared with WT cells under basal conditions (Figure 2K), and they slightly increased CHK1 phosphorylation (Figure 3C), suggesting replication stress, we asked whether auto-modification directly modulates replication dynamics. Using DNA fiber analysis, we confirmed that PARP1 KO accelerated fork speed,²⁷ but this phenotype was rescued by reintroduction of PARP1 WT or 3SA (Figure 3D). However, this was not the case for the PARP1 4SA mutant cells, which showed significantly accelerated DNA replication forks in all cases, with fork speed and stability equivalent to PARP1 KO cells
<figure><img src="image_35.png" /><figcaption>Figure 3. PARP1 auto-modification modulates the speed of replication fork progression</figcaption></figure>

(A) DNA fiber analysis of replication fork speed in parental (U2OS), HPF1 KO, and ARH3 KO cell lines treated with 10 µM olaparib for 24 h as indicated (n = 250 forks per condition per cell line).

(B) QIBC analysis of the γH2AX level in different cell-cycle phases in the same conditions as in (B), *n* = 1,000 nuclei per cell line per condition.

(C) Western blot analysis of CHK1-phosphorylation and CHK1 in cells expressing the indicated PARP1 variants.

(D) DNA fiber analysis of replication fork speed in cell lines expressing the indicated PARP1 variants, and U2OS indicates the parental cell line (n = 200 forks per cell line).

(E) QIBC analysis of nuclear levels of phosphorylated replication protein A (pRPA) in the indicated cell lines, *n* = 1,000 nuclei per condition.

(F) Analysis of γH2AX level in S-phase with QIBC in PARP1 WT and 4SA mutant cells, *n* = 1,000 nuclei per condition.

Significance was determined using Welch's two-sample t test for all panels. Thresholds for *p* value cutoffs are as follows: \* *p* < 0.05, \*\* *p* < 0.01, \*\*\* *p* < 0.001, \*\*\*\* *p* < 0.0001. *p* values were adjusted with the Holm correction to account for multiple hypothesis testing.

(Figures 3D and S3C). To complement this result, we also measured phosphorylation of RPA, a prominent replication stress marker,⁵⁷ and found it to be elevated in cells with accelerated replication forks (Figure 3E). Accordingly, we also found that PARP1 4SA cells in S-phase experienced significantly elevated γH2AX (Figure 3F). Our findings collectively demonstrate that PARP1 auto-modification plays a crucial role in maintaining normal rates of genomic DNA synthesis and is regulated through the serine-ADPr axis. When this homeostasis is perturbed, cells experience replication stress and accumulate DNA damage.

Loss of PARP1 auto-modification impairs
alternative OFP

In the absence of exogenous DNA damage, ADPr has been
shown to increase during S-phase of the cell cycle.²⁴ Using
quantitative-image-based cytometry⁵⁸ (QIBC), we visualized
cell-cycle distributions by tracking replicating cells by quanti-

fying their incorporation of EdU, which combined with immuno-
fluorescence staining allows us to monitor nuclear ADPr levels
across large cell populations (Figure 4A). Loss of PARP1 drasti-
cally reduced nuclear ADPr in S-phase, while both WT and 4SA
mutant cells rescued this signal, confirming their activation dur-
ing DNA replication. However, the ADPr targets during DNA
replication have never been studied systematically and remain
largely uncharted.

To address this, we synchronized PARP1 WT and 4SA mutant
cells using a double-thymidine block. After release into S-phase,
ADPr-modified proteins were enriched and detected by MS
analysis (Figure 4B; Table S5). To promote the accumulation of
ssDNA gaps activating PARP1 during DNA replication, we addi-
tionally inhibited FEN1, a nuclease essential for canonical OFP.
Our analysis confirmed selective enrichment of ADPr-modified
proteins (Figure S4A), with a high degree of experimental repro-
ducibility (median CV < 15%; Figures S4B and S4C). Gene
<figure><img src="image_36.png" /><figcaption>**Figure 4.** Loss of PARP1 auto-modification impairs alternative OFP</figcaption></figure>

(A) QIBC analysis of chromatin-associated ADPr intensity throughout the cell cycle in the indicated cell lines treated with 10 µM PARG inhibitor for 30 min (n = 800 cells per cell line).

(B) Experimental overview to identify ADPr targets during S-phase by MS. Cells were synchronized by double-thymidine block and released into S-phase. 1 h after release into S-phase, 10 µM FEN1 inhibitor was added for 1 h before purification of ADPr-modified proteins with the Af1521-macrodomain and subsequent MS analysis.

(C) Gene set enrichment analysis displaying biological processes significantly enriched for ADPr-target proteins during S-phase.

(D) Analysis of proteins differentially modified by ADPr during S-phase in response to FEN1-inhibition as described in (B) in PARP1 WT and 4SA mutant cells.

(E) STRING network analysis of significant FEN1-inhibitor-dependent ADPr targets during S-phase. The network displays the functional interconnectivity (edges) between the targets (nodes) identified, with a PPI-enrichment *p* < 1E-10. Nodes are colored according to their significant enrichment of the indicated terms on the legend, with a false discovery rate (FDR)-corrected *p* < 0.05.

(F) Normalized growth curves of PARP1 WT and 4SA mutant cells after treatment with 20 µM FEN1 inhibitor on day 0. Data represent the mean cell area ± SE (n = 3).

(G) Model showing the role of PARP1 auto-modification for OFP. The model shows the presence of single-strand gaps/nicks on the lagging strand, which promote PARP1 recruitment. Proposed mechanism of action for PARP1 WT (top) and auto-modification-deficient mutant (bottom) are shown in the figure.
Ontology analysis highlighted biological processes related to protein PARylation and DNA replication as the top hits, but we also identified several proteins implicated in mitochondrial functions, which broadens the scope of S-phase ADPr beyond nuclear processes (Figure 4C).

We next examined modification of PARP1 itself during S-phase. Only PARP1 WT, but not the 4SA mutant, was significantly ADPr-modified (Figure S4D), validating that the 4SA mutant is auto-modification deficient. Considering the pivotal role of PARP1 during OFP, we sought to identify ADPr targets of PARP1 when this alternate repair pathway is activated with FEN1 inhibition (Figure 4D). We confirmed that FEN1 inhibition promoted the activation of PARP1 WT and its subsequent auto-modification (Figure S4E), unlike PARP1 4SA, which was not enriched (Figure S4F). The FEN1i-dependent proteins formed a highly interconnected network pinpointing ADPr targets relevant to OFP at the replication fork (Figure 4E). These proteins have roles in DNA replication and repair, underscoring their importance in sensing and repairing ssDNA gaps. For example, we identified the RFC complex, which continually loads PCNA at Okazaki fragments near the primer-template junction generated by POLα on the lagging strand,⁵⁹⁻⁶¹ together with the canonical (LIG1/FEN1) and non-canonical (PARP1/ XRCC1/LIG3) enzymes for OFP. We emphasize that there is a large overlap of ADPr targets between WT and 4SA mutant cells, consistent with the observation that PARP1 auto-modification does not influence the specificity of trans-ADPr (Figure S2D).

Our data support a role for PARP1 in the resolution of unligated Okazaki fragments during DNA replication. Thus, we wondered if the elevated levels of DNA damage we detected in PARP1 4SA cells during DNA replication could be an indication that they are impaired in facilitating alternative OFP (Figures 3E and 3F). To address this, we treated PARP1 WT and 4SA mutant cells with FEN1 inhibitor to promote a situation where cells must rely exclusively on the alternative OFP pathway mediated by PARP1 for repair. We found that PARP1 4SA mutant cells were highly sensitive to FEN1 inhibition, while WT cells were not (Figure 4F), despite comparable growth under unperturbed conditions (Figure S4G). Of note, both cell lines had a similar rate of proliferation during the first ~24 h following FEN1i treatment, supporting a model where the unrepaired ssDNA gaps are inherited by the daughter cells, overwhelming these cells in the subsequent S-phase where they may be converted to DSBs. Altogether, our proteomics investigation reveals the ADPr targets of the replication fork and highlights the critical role of PARP1 auto-modification for facilitating alternative OFP.

DISCUSSION

Since the discovery that PARPi is synthetic lethal with HR deficiency,¹,² significant efforts have been made to understand the underlying mechanism(s). One prevailing hypothesis is that PARP activity during DNA replication is critical for maintaining DNA synthesis fidelity. Specifically, PARP1 may sense unligated Okazaki fragments on the lagging strand, and its inhibition could disrupt lagging strand maturation, increasing ssDNA gaps.²⁴,²⁶,⁶²

In this study, we developed a PARP1 mutant deficient in auto-modification while preserving catalytic activity, allowing for a separation of those functional roles. Our findings reveal that PARP1 trapping may perturb DNA repair, even when catalytic activity is intact, suggesting that the cytotoxic effects of PARPi cannot be attributed solely to loss of catalytic function for downstream signaling. Our results indicate that PARP1 trans-modification is sufficient for recruiting repair factors, whereas auto-modification is required for eviction of PARP1 and the timely exchange of repair factors at damaged DNA (Figure 4G). Supporting this, catalytically inactive PARP1 is lethal in mice, while PARP1 KO is not.⁶³,⁶⁴ Furthermore, comparisons between different types of PARPi that either promote retention or release by allosteric regulation found that compounds increasing retention are more cytotoxic.¹⁹ Mechanistically, PARP1 binding to DNA breaks can block MRN complex access and prevent end resection.⁴⁸ Consistently, our PP-MS analysis showed that the PARP1 4SA mutant, but not WT, abrogates MRN complex binding to damaged DNA (Figures 2H and 2I), underscoring that free access to repair substrate is also necessary for faithful repair.

The observation that PARP1 auto-modification is not strictly required for recruitment of most DNA-repair factors is surprising, as this modification has generally been regarded as a key platform for repair-complex assembly. Interestingly, Hrychova et al. recently showed that HPF1-dependent ADPr is largely dispensable for most aspects of SSB repair.⁵⁰ Combined, these two observations imply that while recruitment of repair factors such as XRCC1 is PARP1-dependent, they do not specifically rely on ADPr of one single substrate. In support of this model, Kliza et al. used PARylated biotin probes to enrich readers of this modification and found XRCC1 to be the strongest binder.⁶⁵ Notably, this experiment was performed under conditions where the PAR chain was not anchored to any protein substrate, suggesting that XRCC1 cannot discriminate between the various factors modified by PAR. Thus, numerous chromatin targets could be employed for repair factor recruitment, including but not limited to PARP1, histones,⁴⁹,⁶⁶ DNA,⁶⁷ or even untethered PAR chains synthesized at DNA breaks.⁵⁸ We note that the prevalence of protein-free PAR chains is highest when HPF1 is absent, making it particularly relevant to consider this contribution under HPF1-deficient conditions.

We previously showed that PARP1 inhibition accelerates replication fork progression and induces replication stress.²⁷ Here, we expand on this concept by demonstrating that serine ADPr, controlled via the HPF1/ARH3 axis, regulates replication fork speed. Auto-modification-deficient PARP1 fails to restore fork speed to physiological levels despite maintaining trans-modification activity, highlighting a unique role for auto-modification in restraining replication. One possibility is that PARP1 auto-modification promotes its timely eviction at replication forks to allow exchange with other factors. Another is that it mediates interactions with fork-specific proteins not required during SSB repair. Supporting this, PARP1 and TIMELESS physically interact⁶⁹ and function together to safeguard the replisome by sensing transcription-replication conflicts, which are increased with inhibition of PARP1.⁷⁰ Disruption of this interaction leads to DNA damage during replication,⁷⁰ with TIMELESS binding auto-modified PARP1.⁷¹ The
TIMELESS-PARP1 interaction also facilitates repair of unil-
gated Okazaki fragments and suppresses lagging-strand
ssDNA gaps.⁷² Loss of PARP1 auto-modification could there-
fore decouple this pathway, explaining our observation that
4SA mutant cells experience replication stress and are syn-
thetic lethal with FEN1 inhibition. Similarly, CARM1/PRMT4,
another regulator of replication fork speed preventing ssDNA
gap formation,⁵³ interacts with PARP1. By contrast, this bind-
ing is strengthened with PARPi but lost with auto-modification.
Thus, our results are consistent with a model where PARP1
auto-modification enables factor exchange (e.g., TIMELESS
and CARM1) at replication forks to fine-tune fork speed, a pro-
cess potentially disrupted in 4SA mutant cells.

Beyond these mechanistic insights, our proteomics study also provides a resource for ADPr regulation during DNA replication. We mapped S-phase ADPr targets to pinpoint the proteins ADP-ribosylated by PARP1 during alternative OFP when the canonical pathway is blocked. We identify several lagging-strand-associated factors, such as FEN1/LIG1, which is associated with the lagging strand via interactions with PCNA,⁷³ and the entire RFC complex, which loads PCNA at the primer-template junction.⁷⁴ While the presence of DNMT1/UHRF1 at the replication fork to propagate DNA methylation is not a novel concept,⁷⁵ the notion that ADPr has regulatory functions for this process has never previously been described. Furthermore, we also found a substantial amount of chromatin remodeling factors present, and both the FACT complex and ALC1 are known to be recruited by ADP-ribosylated histones.⁷⁶,⁷⁷

When FEN1 activity is blocked, the number of ssDNA gaps on the lagging strand increases due to ineffective OFP,²⁴,²⁵ with a significant amount of gaps accumulating behind the replication fork.²⁶ Here we observe robust ADPr on HLTF, and this is unexpected, because the HLTF helicase typically is associated with the 3' end of the nascent leading strand,⁷⁸ where it facilitates fork reversal during replication stress and regulates fork progression.⁷⁹ Our data suggest that HLTF may also be recruited to the free 3' end of DNA gaps on the lagging strand some distance behind the replication fork; however, the functional implications of this are entirely unexplored.

In conclusion, we generated a PARP1 mutant deficient in ADPr auto-modification that simultaneously retains catalytic activity, revealing dual roles of PARP1 in DNA repair and repli-cation. Our data demonstrate an important regulatory role for PARP1 auto-modification in DNA-damage signaling and repli-cation fork progression, with loss of auto-modification as an important dimension for PARPi-driven cytotoxicity, separate from catalytic activity toward other downstream targets. This study serves as a valuable resource and furthers our under-standing of the underlying mechanisms of PARPi sensitivity for cancer therapy.

Limitations of the study

Currently, no bottom-up proteomics method can directly distin-
guish between MAR and PAR on peptides. Consequently, our
MS data report on the stoichiometry and relative regulation of
ADPr sites without directly resolving MAR versus PAR contribu-
tions. While western blotting can detect PAR directly, it lacks
the specificity for identifying and quantifying individual modified

protein targets. By contrast, MS provides site-specific resolution
but cannot discern chain length or branching, resulting in com-
plementary—but not fully overlapping—insights into ADPr
signaling.

While mutational analysis of the PARP1 4SA mutant reduced auto-modification by approximately 99%, a small residual signal remains. Further mutagenesis could potentially eliminate additional modification sites; however, considering the extensive auto-modification landscape of PARP1, introducing a large number of mutations (i.e., 56 or more) would likely affect overall protein function and confound interpretations regarding site-specific roles in ADPr signaling.

Finally, we observed activation of DNA-damage response and replication stress pathways in PARP1 4SA cells. However, we cannot definitively determine whether these phenotypes result from increased levels of unrepaired DNA lesions (e.g., ssDNA gaps) or from defective repair processes themselves. This limitation reflects a broader challenge in genome stability research, where damage accumulation and repair efficiency are interdependent variables. For instance, impaired repair can increase ssDNA gaps, while excessive damage may overwhelm repair capacity, prompting reliance on error-prone pathways.⁸⁰ Disentangling these effects will require more targeted mechanistic studies.

RESOURCE AVAILABILITY

Lead contact

Further information and requests for resources should be directed to and will be fulfilled by the lead contact, Michael L. Nielsen (michael.lund.nielsen@cpr.ku.dk).

Materials availability

Requests for generated materials should be directed to the corresponding
authors.

Data and code availability

* The MS data have been deposited to the ProteomeXchange Consortium via the PRIDE partner repository,⁸¹ with the dataset identifiers PRIDE: PXD052191 and PRIDE: PXD066033.

* Code availability: steps to follow to process data from raw MaxQuant data output are available on GitHub at https://github.com/Jonaselsborg/Peptide-modification-extractor.

* Any additional information required to reanalyze the data reported in this paper is available from the **lead contact** upon request.

ACKNOWLEDGMENTS

We thank the lab of Ivan Ahel for U2OS cell lines with *PARP1*, *ARH3*, and *HPF1* KOs and for providing purified HPF1 protein. We thank Pablo Hernandez-Varas and Thomas Hartig Braunstein from the Core Facility for Integrated Microscopy, Faculty of Health and Medical Sciences, University of Copenhagen, for training and support. We acknowledge the assistance and support by the Protein Production and Characterization Platform at the Novo Nordisk Foundation Center for Protein Research for aid in purifying variants of PARP1. We would like to thank Martin Rykær and Jeppe Madsen from the Mass Spectrometry Platform at the Novo Nordisk Foundation Center for Protein Research for excellent instrument support. We would like to extend our gratitude toward Sergey Vakhrushev for providing instrument access at a critical moment when our own Lumos instrument was undergoing repairs as the revised manuscript was under preparation. We thank Jiri Lukas and Gijs Zonderland for critical reading of the manuscript. R.P. was supported by a fellowship from the
Fondation pour la Recherche Médicale (ECO202406019133), and S.Z. received funding from the Fondation ARC pour la recherche sur le cancer (ARCDOC42022010004560). Z.F. was supported by the Novo Nordisk Foundation (grant no. NNF18CC0033876). This project received funding from the Danish Independent Research Fund (DFF, grant agreement 9060-00030B) and the Novo Nordisk Foundation (grant no. NNF22OC0074140). A.M.-M. was supported by The Danish Cancer Society, KVBU_R302-A17590 and KBVU_R374-A22693. J.B. was supported by the Danish Cancer Society, R322-A17482; the Novo Nordisk Foundation, NNF20OC0060590; and the Swedish CancerFonden: 170176. The Novo Nordisk Foundation Center for Protein Research is supported financially by the Novo Nordisk Foundation (grant agreement NNF14CC0001).

## AUTHOR CONTRIBUTIONS

J.D.E., S.H.N.M., R.P., Z.F., A.A.-R., S.Z., and A.M.-M. performed the experiments. J.D.E., M.L.N., A.M.-M., J.P.D., and S.H. designed the experiments. J.D.E. performed liquid chromatography-mass spectrometry (LC-MS) analysis. J.D.E., S.H.N.M., A.A.-R., R.P., I.A.H., V.I., and R.S. analyzed the data. J.D.E. wrote the paper with contributions from M.L.N., S.H., S.H.N.M., Z.F., I.A.H., S.C.B.-L., and A.M.-M.; J.B. secured funding and discussed the findings. All authors read and accepted the manuscript. M.L.N. conceived the study. M.L.N., A.M.-M., S.H., and J.P.D. supervised the research.

## DECLARATION OF INTERESTS

The authors declare no competing interests.

## STAR★METHODS

Detailed methods are provided in the online version of this paper and include the following:

### KEY RESOURCES TABLE

#### EXPERIMENTAL MODEL AND STUDY PARTICIPANT DETAILS

##### METHOD DETAILS

* SDS-PAGE and Western blotting

* Laser Microirradiation and Protein Recruitment Assays

* Cell growth assay

* DNA fiber assay

* Immunostaining and quantitative image based cytometry (QIBC)

* UV-A laser stripe ablation

* Cell treatment for site-specific ADP-ribosylome

* Expression and purification of GST-Af1521

* Purification of ADP-ribosylated peptides using Af1521 macro-domain

* Xenopus Egg Extracts

* Expression and purification of PARP1 variants

* Plasmid pulldowns for mass spectrometry

* Purification of ADPr-modified proteins during S-phase for MS analysis

* Purification of ADPr-modified PARP1 protein while preserving ester-linked modifications

* MS instrument analysis

* MS data processing

#### QUANTIFICATION AND STATISTICAL ANALYSIS

* Data filtering of site-specific ETD-based global ADPr data

* Data filtering of PARP1 ADPr sites recorded with ETD fragmentation while preserving ester-linked modifications

* Quantification and statistical analysis of MS data

## SUPPLEMENTAL INFORMATION

Supplemental information can be found online at https://doi.org/10.1016/j.molcel.2025.09.006.

Received: August 2, 2024

Revised: July 21, 2025

Accepted: September 3, 2025

Published: October 2, 2025

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## STAR★METHODS

### KEY RESOURCES TABLE

<table><thead><tr><th>REAGENT or RESOURCE</th><th>SOURCE</th><th>IDENTIFIER</th></tr></thead><tbody><tr><td colspan="3"><strong>Antibodies</strong></td></tr><tr><td>ADP-ribose (mono/poly) [E6F6A]</td><td>Cell Signaling</td><td>Cat#83732;RRID:AB_2749858</td></tr><tr><td>GAPDH</td><td>Abcam</td><td>Cat#ab9485;RRID:AB_307275</td></tr><tr><td>Vinculin</td><td>Sigma</td><td>Cat#V9131;RRID:AB_477629</td></tr><tr><td>GFP</td><td>Roche</td><td>Cat#11814460001;RRID:AB_390913</td></tr><tr><td>PARP1 [46D11]</td><td>Cell Signaling</td><td>Cat#9532S;RRID:AB_659884</td></tr><tr><td>PARP1 [ab32138]</td><td>Abcam</td><td>Cat#ab32138;RRID:AB_777101</td></tr><tr><td>PARP1 [for PP-MS immunodepletion]</td><td>Ryu et al.<sup>82</sup></td><td>From Yoshiaki Azuma</td></tr><tr><td>CHK1</td><td>Santa Cruz</td><td>Cat#sc-8408;RRID:AB_627257</td></tr><tr><td>p-CHK1 (S345)</td><td>Cell Signaling</td><td>Cat#2348;RRID:AB_331212</td></tr><tr><td>β-Actin</td><td>Sigma</td><td>Cat#A1978;RRID:AB_476692</td></tr><tr><td>p-H2A.X (S139)</td><td>Abcam</td><td>Cat#ab22551;RRID:AB_447150</td></tr><tr><td>p-RPA</td><td>Novus</td><td>Cat#NB100-544;RRID:AB_10002227</td></tr><tr><td>PARP1-Trap nanobody (agarose-linked)</td><td>Chromotek</td><td>Cat#xta;RRID:AB_2631384</td></tr><tr><td>p-NBS1 (S343)</td><td>Upstate</td><td>Cat#05-663</td></tr><tr><td>BrdU (rat)</td><td>Abcam</td><td>Cat#ab6326;RRID:AB_305426</td></tr><tr><td>BrdU (mouse)</td><td>Becton, Dickinson</td><td>Cat#347580;RRID:AB_10015219</td></tr><tr><td>DyLight 550 (anti-rat)</td><td>Invitrogen</td><td>Cat#SA5-10019;RRID:AB_2556599</td></tr><tr><td>Alexa Fluor 488 (anti-mouse)</td><td>Invitrogen</td><td>Cat#A21202;RRID:AB_141607</td></tr><tr><td colspan="3"><strong>Bacterial and virus strains</strong></td></tr><tr><td>Rosetta 2 (DE3) Competent E. coli</td><td>Sigma-Aldrich</td><td>Cat#71397</td></tr><tr><td colspan="3"><strong>Chemicals, peptides, and recombinant proteins</strong></td></tr><tr><td>PARPi - Olaparib (AZD2281)</td><td>Selleckchem</td><td>Cat#S1060</td></tr><tr><td>PARGi (PDD 00017273)</td><td>Tocris</td><td>Cat#5952</td></tr><tr><td>FEN1i (LNT1)</td><td>Tocris</td><td>Cat#6510</td></tr><tr><td>Geneticin (G418 Sulfate)</td><td>Gibco</td><td>Cat#11811023</td></tr><tr><td>Hoechst 33342, trihydrochloride trihydrate</td><td>Invitrogen</td><td>Cat#H3570</td></tr><tr><td>DAPI</td><td>Invitrogen</td><td>Cat#62248</td></tr><tr><td>Hydrogen Peroxide (30%)</td><td>Sigma-Aldrich</td><td>Cat#H1009</td></tr><tr><td>Methyl Methanesulfonate (MMS)</td><td>Sigma-Aldrich</td><td>Cat#129925</td></tr><tr><td>Lysyl Endopeptidase (Lys-C)</td><td>Fujifilm Wako</td><td>Cat#125-05061</td></tr><tr><td>Trypsin</td><td>Promega</td><td>Cat#V5117</td></tr><tr><td>PARG (purified protein)</td><td>Nowak et al.<sup>83</sup></td><td>From Michael O. Hottiger</td></tr><tr><td>Benzonase</td><td>Millipore</td><td>Cat#E1014</td></tr><tr><td>GST-Af1521 (agarose bound)</td><td>Anagho et al.<sup>32</sup></td><td>N/A (In house)</td></tr><tr><td>5-Chloro-2'-deoxyuridine (CldU)</td><td>Invitrogen</td><td>Cat#C6891</td></tr><tr><td>5-lodo-2'-deoxyuridine (ldU)</td><td>Invitrogen</td><td>Cat#I7125</td></tr><tr><td>Click-iT EdU Cell Proliferation Kit</td><td>Invitrogen</td><td>Cat#C10340</td></tr><tr><td>5-Bromo-2'-Deoxyuridine (BrdU)</td><td>Invitrogen</td><td>Cat#B23151</td></tr><tr><td>HPF1 (purified protein)</td><td>Suskiewicz et al.<sup>9</sup></td><td>From Ivan Ahel</td></tr><tr><td>ADP-HPD</td><td>Sigma-Aldrich</td><td>Cat#118415-60UG</td></tr><tr><td>cOmplete, EDTA-free Protease Inhibitor Cocktail</td><td>Roche</td><td>Cat#11873580001</td></tr><tr><td>PIPES</td><td>Sigma-Aldrich</td><td>Cat#P6757</td></tr></tbody></table>

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<table><thead><tr><th>REAGENT or RESOURCE</th><th>SOURCE</th><th>IDENTIFIER</th></tr></thead><tbody><tr><td colspan="3"><strong>Deposited data</strong></td></tr><tr><td>Raw MS data (ADP-ribosylome, PP-MS, S-phase ADPr)</td><td>This paper</td><td>PRIDE: PXD052191</td></tr><tr><td>Raw MS data (preserved ester-bonds, PARP1 PD, ADPr sites)</td><td>This paper</td><td>PRIDE: PXD066033</td></tr></tbody></table>

**Experimental models: Cell lines**

<table><tr><td>U2OS</td><td>ATCC</td><td>Cat#HTB-96</td></tr><tr><td>U2OS PARP1 KO</td><td>Gibbs-Seymour et al.<sup>13</sup></td><td>From Ivan Ahel</td></tr><tr><td>U2OS HPF1 KO</td><td>Gibbs-Seymour et al.<sup>13</sup></td><td>From Ivan Ahel</td></tr><tr><td>U2OS ARH3 KO</td><td>Fontana et al.<sup>51</sup></td><td>From Ivan Ahel</td></tr><tr><td>U2OS eGFP-PARP1 WT</td><td>This paper</td><td>N/A</td></tr><tr><td>U2OS eGFP-PARP1 3SA mutant</td><td>This paper</td><td>N/A</td></tr><tr><td>U2OS eGFP-PARP1 4SA mutant</td><td>This paper</td><td>N/A</td></tr></table>

**Oligonucleotides**

<table><tr><td>dsDNA 30 nucleotides (for PARP1 activation in vitro); Fwd:ATCAGATAGCATCTGTGCGGCCGCTTAGGG; Rev:CCCTAAGCGGCCGCACAGATGCTATCTGAT</td><td>This paper</td><td>N/A</td></tr></table>

**Recombinant DNA**

<table><tr><td>pcDNA3.1-eGFP-PARP1 WT (Uniprot: P09874)</td><td>Genscript</td><td>N/A</td></tr><tr><td>pcDNA3.1-mCherry-PARP1 WT (Uniprot: P09874)</td><td>This paper</td><td>N/A</td></tr><tr><td>pcDNA3.1-eGFP-PARP1 3SA (S499A, S507A, S519A)</td><td>Genscript</td><td>N/A</td></tr><tr><td>pcDNA3.1-eGFP-PARP1 4SA (S499A, S504A, S507A, S519A)</td><td>Genscript</td><td>N/A</td></tr><tr><td>pcDNA3.1-mCherry-PARP1 4SA (S499A, S504A, S507A, S519A)</td><td>This paper</td><td>N/A</td></tr><tr><td>PATagRFP-H2B</td><td>Sellou et al.<sup>84</sup></td><td>From Sébastien Huet</td></tr><tr><td>GFP-XRCC1</td><td>This paper</td><td>N/A</td></tr><tr><td>pmCherry-PARP1 LW/AA</td><td>Smith et al.<sup>49</sup></td><td>From Sébastien Huet</td></tr><tr><td>eGFP-PARP1 6EA</td><td>Prokhorova et al.<sup>33</sup></td><td>From Ivan Ahel</td></tr><tr><td>pET24-PARP1 WT</td><td>Langelier et al.<sup>85</sup></td><td>From John M. Pascal</td></tr><tr><td>pET24-PARP1 3SA</td><td>Fábián et al.<sup>86</sup></td><td>From Julien P. Duxin</td></tr><tr><td>pET24-PARP1 4SA</td><td>This paper</td><td>N/A</td></tr><tr><td>pET24-PARP1 E988K</td><td>Fábián et al.<sup>86</sup></td><td>From Julien P. Duxin</td></tr><tr><td>pMH<sup>ssDNA-PK</sup> (pGAP)</td><td>Larsen et al.<sup>87</sup></td><td>From Julien P. Duxin</td></tr></table>

**Software and algorithms**

<table><tr><td>MaxQuant (v1.5.3.30)</td><td>Cox et al.<sup>88</sup></td><td>https://www.maxquant.org/</td></tr><tr><td>Cytoscape (v3.10.1)</td><td>Shannon et al.<sup>89</sup></td><td>https://cytoscape.org/</td></tr><tr><td>Arivis Vision4d (v4.1.1)</td><td>Zeiss</td><td>https://www.zeiss.com/microscopy/en/products/software/arivis-pro.html</td></tr><tr><td>ZenBlue (v2.6)</td><td>Zeiss</td><td>https://www.micro-shop.zeiss.com/en/de/softwarefinder/software-categories/zen-blue/zen-blue-pro/</td></tr></table>

**Other**

<table><tr><td>Dulbecco's Modified Eagle Medium (DMEM)</td><td>Gibco</td><td>Cat#31966021</td></tr><tr><td>Leibovitz's L-15 media</td><td>Gibco</td><td>Cat#21083027</td></tr><tr><td>Penicillin-Streptomycin</td><td>Gibco</td><td>Cat#15070063</td></tr><tr><td>Fetal Bovine Serum</td><td>Gibco</td><td>Cat#10438026</td></tr><tr><td>Trypsin-EDTA (0.25%), phenol red</td><td>Gibco</td><td>Cat#25200056</td></tr><tr><td>Dulbecco's phosphate buffered saline (DPBS)</td><td>Gibco</td><td>Cat#14190-094</td></tr><tr><td>Lipofectamine 2000 Transfection Reagent</td><td>Invitrogen</td><td>Cat#11668019</td></tr><tr><td>X-tremeGENE 360</td><td>Roche</td><td>Cat#8724105001</td></tr><tr><td>SepPak C18 Classic (360mg sorbent)</td><td>Waters</td><td>Cat#WAT051910</td></tr></table>

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<table><thead><tr><td>REAGENT or RESOURCE</td><td>SOURCE</td><td>IDENTIFIER</td></tr></thead><tbody><tr><td>Empore C18 SPE discs (StageTip)</td><td>Supelco</td><td>Cat#66883-U</td></tr><tr><td>Ultrafree-MC 0.45µm spin filter</td><td>Millipore</td><td>Cat#UFC30HV</td></tr><tr><td>30 kDa MWCO Centrifugal Filter</td><td>Amicon</td><td>Cat#UFC5030</td></tr><tr><td>100 kDa MWCO Centrifugal Filter</td><td>Sartorius</td><td>Cat#VN01H42</td></tr><tr><td>Dynabeads, M-280 (Streptavidin)</td><td>Invitrogen</td><td>Cat#11206D</td></tr><tr><td>Glutathione Sepharose 4B beads (agarose)</td><td>Cytivia</td><td>Cat#17075601</td></tr></tbody></table>

EXPERIMENTAL MODEL AND STUDY PARTICIPANT DETAILS

Human U2OS osteosarcoma (ATCC HTB-96) cells were acquired from ATCC. U2OS PARP1 KO, U2OS HPF1 KO,¹³ and U2OS ARH3 KO⁵¹ cells were described previously. Cells were cultured at 37°C and 5% CO₂ in Dulbecco's Modified Eagle's Medium (DMEM, Gibco) supplemented with 10% fetal bovine serum (Gibco) and a penicillin/streptomycin mixture (100 U/mL; Gibco).

To generate cell lines stably expressing eGFP-PARP1 wild type (WT) or 3SA, and 4SA mutants, U2OS PARP1 KO cells were transfected with the plasmids pcDNA3.1-eGFP-PARP1 WT (UniProt ID P09874), 3SA (S499A, S507A, S519A), or 4SA (S499A, S504A, S507A, S519A), acquired via GenScript. Transfection was performed using Lipofectamine 2000 (Thermo) according to manufacturer's recommendations. To select for single clones that stably express the transfected plasmid, cells were selected using 0.8 mg/mL G418 Geneticin (Gibco), after which colonies were picked and expanded. All cells were routinely tested for mycoplasma.

Egg extracts were prepared using *Xenopus laevis* (Nasco Cat #LM0053MX, LM00715MX). All experiments involving animals were approved by the Danish Animal Experiments Inspectorate and conform to relevant regulatory standards and European guidelines.

METHOD DETAILS

SDS-PAGE and Western blotting

Cells were collected and lysed in SDS-lysis buffer (2% SDS, 50 mM Tris-HCl pH 8.0) or in Laemmli sample buffer (2% SDS, 50 mM Tris pH 6.8, 100 mM dithiothreitol, 10% glycerol and 0.1% bromophenol blue), denatured for 10 min at 95°C at 1400 RPM, then sonicated for 30 sec using a Sonic Dismembrator (Fisherbrand). Protein concentrations were determined using the BCA assay (Pierce) or with Qubit protein assay (Invitrogen). Samples lysed in SDS-lysis buffer were supplemented with 10 µM DTT and Bromophenol blue. All samples were then analyzed by SDS-PAGE. Proteins were transferred from the gel onto nitrocellulose membrane and detected using immunoblotting in combination with horseradish peroxidase-conjugated secondary antibodies. Peroxidase activity was detected by ECL reagent (GE Healthcare) and Amersham Hyperfilm ECL (GE Healthcare). Antibodies used: ADPr, mono/poly (E6F6A, Cell Signaling 83732, 1:1000), GAPDH (Abcam, ab9485, 1:5000), GFP (Roche, cat 11814460001, 1:1000), PARP1 (Figure 2D) (Cell signaling, 46D11, 1:2500), PARP1 (Figure 3C) (Abcam ab32138, 1:500), CHK1 (Santa Cruz sc-8408, 1:500), p-CHK1 S345 (Cell Signaling 2348, 1:50), β-Actin (Sigma A1978, 1:5,000), p-H2A.X S139 (γH2AX, Abcam ab22551, 1:500).

Laser Microirradiation and Protein Recruitment Assays

U2OS PARP1 knockout cells were seeded in 35 mm dishes with 170 µm glass-bottom coverslips and transfected 48 h prior to aging with either PARP1 WT or 4SA mutant plasmids, as previously described, along with PATagRFP-H2B⁸⁴ using X-tremeGENE 360 (Roche) according to the manufacturer's instructions. For experiments involving the PARP1 6EA mutant and XRCC1, transfection was performed independently in a separate laboratory under similar conditions. In both cases, cells were pre-sensitized with fresh medium containing 0.3 µg/mL Hoechst for 1 h at 37°C prior to laser microirradiation.

Immediately before imaging, the medium was replaced with CO₂-independent, phenol red-free Leibovitz's L-15 medium (Gibco) supplemented with 10% FBS. All live-cell imaging was performed at 37°C using a Zeiss LSM 980 confocal microscope. For the PARP1 WT and 4SA experiments, a C-Apochromat 40×/1.2 NA water immersion objective was used, while the PARP1 6EA and XRCC1 experiments employed a Plan-Apochromat 63×/1.4 NA oil-immersion objective. In both setups, the pinhole was set to 1 Airy unit (corresponding to 36 µm for 488 nm excitation), and fluorescence detection was carried out using GaAsP detectors. GFP was excited at 488 nm and detected using a 500–550 nm bandpass filter; PATagRFP and mCherry (where applicable) were excited at 561 nm and detected using a 579–705 nm filter. Pixel sizes were set to 99 nm or 110 nm, satisfying Nyquist sampling criteria.

Laser microirradiation was used to induce DNA damage in a defined region of interest (ROI). In the WT and 4SA experiments, a 2.5 × 16 µm rectangular ROI was drawn across each nucleus, and DNA damage was induced using a 405 nm laser set to 30% power with 25 iterations. For the 6EA and XRCC1 experiments, the ROI measured 10 µm in width and 1 µm in height, and was irradiated under equivalent conditions. Photoconversion of PATagRFP-H2B was used to mark the damage site in the WT and 4SA experiments. Zeiss Definite Focus 2 was used for focal stabilization during time-lapse acquisition.

To quantify protein recruitment to damage sites, mean fluorescence intensities were measured within the damaged region ($I_d$), across the entire nucleus ($I_n$), and in the background outside of the cell ($I_{bg}$). In the WT and 4SA experiments, these values were
extracted using a custom MATLAB routine based on H2B segmentation; in the 6EA and XRCC1 experiments, automatic segmentation in ImageJ was used following image registration to correct for nuclear motion. In both cases, protein accumulation at sites of damage ($A_d$) was calculated as:

$$A_d = \frac{l_d - l_{bg}}{l_n - l_{bg}}$$

For PARP1 WT, 4SA, and 6EA, intensity within the damaged region was normalized to the pre-damage signal and then was further normalized to the maximum value. The dissipation time was calculated as the time required to reduce signal intensity by 50% following peak recruitment. For XRCC1, intensity values were normalized to pre-damage levels.

## Cell growth assay

U2OS PARP1 WT and 4SA mutant cells were plated in triplicate with a seeding density of 10,000 cells per well in a 24-well plate, using the same cell culture conditions as described previously. After 24 h, the growth medium was supplemented with 20 µM FEN1-inhibitor (Tocris), which marks the first time point. Relative cell density was recorded at 6 h intervals using the Incucyte S3 Live-Cell Analysis System (Sartorius), and normalized to the density of each well at the start of the experiment. Images were recorded using phase-contrast in brightfield mode. Growth curves are represented as means of independent triplicate experiments from separate wells, and error bars represent the standard error.

## DNA fiber assay

Treatment with olaparib was purchased from Selleckchem (S1060). Cells were incubated for 20 min with 25 µM CldU (Sigma-Aldrich), washed in medium (pre-warmed), and incubated for 20 min with 250 µM IdU (Sigma-Aldrich). The pulse-labelled cells were then collected followed by preparation of DNA fibre spreads as previously described. DNA fibres were stretched on five microscope slides per experimental condition, and two to three slides were subsequently stained. First, detection of CldU was done with a rat anti-BrdU antibody (Abcam ab6326, 1:100) followed by detection of IdU with a mouse anti-BrdU antibody (Becton, Dickinson 347580, 1:500). The primary antibodies were then detected using anti-rat DyLight 550 (Invitrogen, SA5-10019, 1:400) and donkey anti-mouse Alexa Fluor 488 (Invitrogen A21202, 1:400) secondary antibodies. Image acquisition of evenly spread fibres was done on a LSM800 confocal microscope (Carl Zeiss) with a Plan-Apochromat 63x/1.4 numerical aperture objective (oil immersion, Carl Zeiss). Semi-automatic acquisition was performed using tile arrays and autofocus with the LSM acquisition software. The LSM ZEN lite (blue edition) software was used to manually analyze double-labelled DNA replication forks. Between 100 and 250 replication forks were analyzed per slide, and all fork measurements for the same experimental condition were pooled together. At least two independent experiments were performed with similar results. For statistical testing and visualization, an equal number of replication forks were randomly sampled from each condition.

## Immunostaining and quantitative image based cytometry (QIBC)

Cells grown in clear-bottom plates were supplemented for 30 min with 10 µM EdU at 37 °C, and left untreated or supplemented with PARP-inhibitor (Olaparib, Selleckchem S1060) 10 µM or PARG-inhibitor (Tocris, 5952). The cells were then washed in cold PBS, pre-extracted on ice in cold 0.2% Triton X-100 for 2 min, washed in cold PBS again, and fixed for 10 min in cold 4% formaldehyde at room temperature. Then, the cells were permeabilized for 15 min with 1% Triton X-100, washed in PBS, and blocked in PBS+ (PBS containing 1% bovine serum albumin and 0.1% Tween 20). Cells were then stained for EdU detection with Click-iT EdU Imaging Kits (Invitrogen) according to manufacturer's instructions, followed by immunostaining with primary antibody (1 hour) and Alexa Fluor-conjugated secondary antibody (Invitrogen), in combination with Hoechst 33342 (Invitrogen), for 45 min. The cells were washed in PBS and PBS+ in between, and after the last, staining incubations. Then, the cells were stored in PBS for acquisition. Primary antibodies used: p-RPA (Novus NB100-544, 1:500), p-H2A.X S139 (γH2AX, Abcam ab22551, 1:500). QIBC image acquisition was carried out on a ScanR high-content screening station (Olympus) with an IX83 motorized microscope (Olympus), a UPLSAPO 2 40× objective (air immersion), and an ORCA-Flash 4.0 V3 camera. Image analysis was done using ScanR analysis (v3.2.0) software. The data from image analysis was analyzed in R (version 4.3.1 "Beagle Scouts") which, in brief, included filtering out dead cells and duplets, performing cell cycle phase-specific analyses (based on EdU and Hoechst signals), and data visualization.

## UV-A laser stripe ablation

Cells grown on coverslips were incubated with 20 µM BrdU for 24 hours to pre-sensitize the DNA to UVA ablation. The coverslips with cells were then transferred to well chamber slides (Ibidi) containing Leibovitz L-15 media (Gibco). UVA-laser ablation was performed using a Zeiss Axiovert 200 microscope, with a 20X air-immersion objective (203/0.8), with a PALM UVA pulse nitrogen laser (30Hz, 337 nm, (P.A.L.M. Laser Technologies) at 37°C. The laser output was set to 42% and the laser energy to 40, and was used to ablate the coverslip in a grid pattern, taking 6 min from the first to the last position, using the PALMRobo software. Immediately after the ablation, the cells were washed in cold PBS, fixed in 4% formaldehyde for 10 min at room temperature, and washed again in PBS. Then, immunostaining was performed as described above to detect p-NBS1 (Upstate 05-663, 1:200). The stained coverslips were mounted with Fluoromount-G (Invitrogen) and acquired on a Celldiscoverer 7 (Zeiss) with a 20X objective (widefield,
0.95 numerical aperture). Image analysis was performed using Arivis Vision4d software (v4.1.1, Zeiss). Only stripes from nuclei containing one stripe were used for further analysis.

## Cell treatment for site-specific ADP-ribosylome

For site-specific ADPr experiments for mass spectrometry analysis, cells were mock-treated or ADP-ribosylation was induced by treatment with either 5 mM hydrogen peroxide (Sigma Aldrich) or 0.02% (v/v) methyl methanesulfonate (MMS, Sigma Aldrich) for 30 min by addition to the culture medium at 37°C. In total, approximately 120 million cells per biological replicate were cultured in 500 mm² square dishes (Corning).

The method for lysis and proteolytic digestion for was done essentially as described previously.32,90 Briefly, cells were washed twice with ice-cold PBS, then scraped at 4°C in PBS and pelleted using centrifugation at 500 × g at 4°C. Then the pellets were lysed using 10x pellet volumes of Lysis Buffer (6 M guanidine-HCl, 50 mM Tris-HCl pH 8.5) by vigorous shaking and vortexing, after which the lysates was snap-frozen in liquid nitrogen and stored at -80°C until further processing.

Lysates were thawed and sonicated at 30 W for 1 second per mL of lysate while kept on ice, using 10 sec pulses using a Sonic Dismembrator (Fisherbrand). Samples were reduced and alkylated by addition of tris(2-carboxyethyl)phosphine (TCEP) and chloroacetamide (CAA) to a working concentration of 5 mM, and were then incubated for 1 h at RT. Proteins were digested using lysyl endopeptidase (Lys-C, 1:200 w/w; Wako Chemicals) for 3 h, and then diluted with three volumes of dilution buffer (50 mM Tris-HCl pH 8.5). Samples were further digested overnight using modified sequencing grade trypsin (1:200 w/w; Promega trypsin MS grade). Digested samples were acidified by the addition of trifluoroacetic acid (TFA) to a final concentration of 0.5% (v/v), cleared by centrifugation, and purified using reversed-phase C18 cartridges (SepPak Classic, 360 mg sorbent, Waters) according to the manufacturer's instructions. Peptides were eluted from the cartridges by gravity flow using 30% acetonitrile (ACN) in 0.1% TFA, then peptides were frozen overnight at -80°C and then lyophilized for 96 h to ensure complete solvent sublimation. Dataset 3 was prepared as described above, except the samples were diluted prior to digestion with Lys-C, and trypsin was omitted.

## Expression and purification of GST-Af1521

GST-Af1521 plasmids were transformed into Rosetta 2 Competent *E.coli* cells (NEB). Then, colonies were picked to grow a culture to 0.6-0.8 at 37°C in LB broth, then moved to 16°C for overnight induction with 0.5 mM IPTG. Pellets were harvested by several rounds of centrifugation in 50mL tubes (3000 × g at RT), after which they were frozen at -80°C. Pellets were lysed by thawing them on ice, and then adding lysis buffer [50 mM Tris-HCl, pH 7.5, 150 mM NaCl, 1 mM MgCl₂, 1 mM DTT, 1x bug buster (Roche), 1 µL/mL benzonase, 200 µg/mL lysozyme, protease inhibitor cocktail (Roche)], vortexing the pellet to bring it into suspension. Lysates were then incubated at RT on a rolling mixer for 20 minutes, followed by sonication (80W for 60s total in 10 second intervals on ice; Sonic Dismembrator, Fisherbrand), after which debris were pelleted by centrifugation at 4500 × g at 4°C for 30 min. Cleared lysate was transferred to new tubes, and supplemented with sepharose 4B beads (2mL dry volume for every 100mL of culture; Cytiva), after which binding was facilitated by incubating the tubes for 3h on a rolling mixer at 4°C. Beads were pelleted by centrifugation (500 × g at 4°C for 3 min), and transferred to a new 15mL tube. Then the beads were washed 6 times in ice cold PBS, after which they were resuspended in storage buffer (PBS supplemented with 10mM sodium azide) and kept at 4°C until further use.

## Purification of ADP-ribosylated peptides using Af1521 macrodomain

Lyophilized peptides were redissolved in AP buffer (50 mM Tris-HCl pH 8.0, 1 mM MgCl₂, 250 µM DTT, and 50 mM NaCl), then ADP-ribose polymers were reduced to monomers by addition of PARG enzyme (kind gift from Prof. Dr. Michael O. Hottiger) in a 1:10,000 w/w ratio overnight at RT. Then, ADP-ribosylated peptides were purified using a GST-tagged Af1521 macrodomain,32 as described above. Briefly, the enrichment was performed by addition of 100 µL of dry volume of GST-Af1521 beads to each sample, after which binding was facilitated by tumbling end-over-end for 3 h at 4°C. Beads were transferred to 1.5 mL LoBind tubes (Eppendorf) and washed 4x in AP buffer, changing tubes between the 2nd and 4th washing steps. Peptides were eluted twice from the GST-Af1521 beads by addition of two bead-volumes of ice-cold 0.15% TFA, after which the supernatants were pooled and cleared by centrifugation at 5000 × g for 3 min through a 0.45 µm spin filter (Ultrafree-MC, Millipore). To separate the purified peptides from the macrodomain, samples were subsequently spun through 100 kDa cut-off filters (Vivacon 500, Sartorius) which retain the macrodomain. Finally, ADP-ribosylated peptides were purified using C18 StageTips at high pH.90 Briefly, samples were basified by adding ammonium hydroxide to a final concentration of 50 mM, and then loaded onto StageTips carrying four layers of C18 disc material (punch-outs from 47 mm C18 3 M extraction discs, Empore). For dataset 1; Peptides were eluted in 3 block fractions by addition of 4, 10 and then finally 25% acetonitrile in 50 mM ammonium hydroxide. The flow through from high-pH StageTip loading was acidified with to a final concentration of 1% TFA and purified using low-pH StageTips using 25% ACN in 0.1% formic acid. For dataset 3; following 100 kDa spinfilter purification, one-third of the sample was purified using low-pH C18 StageTips as single shots and eluted with 30% acetonitrile in 0.1% formic acid. Two-thirds was purified as described above, using high-pH StageTip purification, but eluted with 30% acetonitrile in 50 mM ammonium hydroxide. Solvents were removed by heating them to 60°C in a SpeedVac under low atmospheric pressure. Peptides were dried to completeness and then dissolved in 0.1% formic acid, and frozen at -20°C until MS analysis.
## Xenopus Egg Extracts

For high-speed supernatant (HSS) preparation, 6 female frogs (Nasco) were primed by injection with 80 IU of human chorionic gonadotropin (hCG, Sigma). 2-7 days after priming, frogs were injected with 625 IU of hCG and placed in individual tanks containing 100 mM NaCl. 18-20 h post injection, eggs were collected and used for extract preparation. Eggs were first dejellied in cysteine buffer for 7 min (2.2% cysteine-HCl, pH 7.7), washed three times in 0.5× MMR buffer (final concentration: 50 mM NaCl, 1 mM KCl, 0.25 mM MgSO₄, 1.25 mM CaCl₂, 2.5 mM HEPES, 0.05 mM EDTA, pH 7.8) and washed three times in ELB sucrose buffer (2.5 mM MgCl₂, 50 mM KCl, 10 mM HEPES, 250 mM sucrose, 1 mM DTT, 50 µg/ml cycloheximide, pH 7.8). Eggs were packed for 1 min at 176 × g and crushed for 20 min at 20,000 × g in a swing bucket rotor at 4°C in the presence of cytochalasin B (2.5 µg/ml), aprotinin (5 µg/ml) and leupeptin (5 µg/ml). Crude interphase extract was recovered post-centrifugation and spun in ultracentrifuge for 90 min at 260,000 × g at 2°C following addition of cycloheximide (50 µg/ml), DTT (1 mM), aprotinin (10 µg/ml), leupeptin (10 µg/ml) and cytochalasin B (5 µg/ml). Following centrifugation, the lipid layer on top was removed. The soluble HSS was harvested, snap frozen in 33 µl aliquots and stored at -80°C.

## Expression and purification of PARP1 variants

The plasmid for expressing human WT PARP1 was a kind gift from J. M. Pascal. PARP1 S499A, S504, S507A and S519A mutations were introduced via Quikchange mutagenesis (Agilent) and confirmed by Sanger sequencing. To purify WT and mutant PARP1, a modified 3-step purification protocol used.⁸⁵ Each PARP1 plasmid was transformed into Rosetta 2 Competent *E.coli* cells (NEB). Cells were grown at 37°C to O.D. 0.8-1.0 in LB broth supplemented with 0.1 mM ZnSO₄ and moved to 16°C for overnight induction with 0.2 mM IPTG. Bacteria were harvested by centrifugation and resuspended in 25 mL of lysis buffer (25 mM HEPES pH 8.0, 500 mM NaCl, 0.5 mM TCEP and 10 mM benzamide). Suspensions were sonicated and cleared by high-speed centrifugation at 40,000 × g for 1 h at 4°C. Cleared lysates were loaded on a HisTrap HP column (Cytiva) with binding buffer (25 mM HEPES pH 8.0, 500 mM NaCl and 0.5 mM TCEP) and bound proteins were washed with 50 mL of high salt buffer (25 mM HEPES pH 8.0, 1 M NaCl, 20 mM imidazole and 0.5 mM TCEP) followed by 50 mL of low salt buffer (25 mM HEPES pH 8.0, 1 M NaCl, 20 mM imidazole and 0.5 mM TCEP). Proteins were eluted with elution buffer (25 mM HEPES pH 8.0, 500 mM NaCl, 400 mM imidazole and 0.5 mM TCEP). Collected proteins were diluted with no salt buffer (50 mM Tris-HCl pH 7.0, 1 mM EDTA and 0.1 mM TCEP) to a final NaCl concentration of 375 mM and further purified with a HiTrap Heparin HP column (Cytiva) to remove contaminant DNA from the protein preparations. Proteins were washed with buffer A (50 mM Tris-HCl pH 7.0, 1 mM EDTA, 375 mM NaCl and 0.1 mM TCEP) and subsequently eluted with an increasing NaCl gradient. Fractions containing the target proteins were collected, concentrated to 3 mL using 30,000 MWCO centrifugal filters (Amicon) and further purified on a HiPrep 16/60 Sephadryl S-200 HR gel filtration column (Cytiva) with PARP1 storage buffer (25 mM HEPES pH 8.0, 150 mM NaCl, 1 mM EDTA and 0.1 mM TCEP). The proteins were then flash-frozen in liquid nitrogen and stored at -80°C.

## Plasmid pulldowns for mass spectrometry

Depletion of *Xenopus* PARP1 from HSS egg extracts was achieved by immunoprecipitation with a home-made PARP1 antibody,⁸² conjugated to Protein A Sepharose Fast Flow (PAS) (GE Health Care). Briefly, beads were washed in PBS buffer, after which antibody containing serum, or an equivalent control serum, was incubated overnight at 4°C tumbling end-over-end. Then, the conjugated beads were subsequently washed first in PBS, and then twice in ELB-sucrose (10 mM HEPES pH 7.7, 50 mM KCl, 2.5 mM MgCl₂, and 250 mM sucrose), twice with ELB. The depletion was performed by removing the supernatant, and adding the HSS extract onto the dry beads, incubating for 15 min at RT before harvesting. The depletion procedure was repeated three times.

The pMH<sup>ssDNA-PK</sup> (pGAP) plasmid was generated as previously described.⁸⁷,⁹¹ Briefly, plasmids were nicked with Nb.BbvCl (NEB), and ligated with the following fluorinated oligo: 5'-TGAGGTAC[C5-FlurodC]GGATTGAGTAGCTACCGGATGC-3'. The resulting plasmid was then cut with Nt.BbvCl (NEB) and the resulting 31bp fragment was melted off and captured by annealing to an excess complementary oligo. Excess oligos were then degraded by Exonuclease I (NEB) treatment. The gapped plasmid was then recovered by phenol/chloroform extraction. A DNA protein crosslink was formed by mixing M.Hpall-His₆ with fluorinated plasmid in reaction buffer (50 mM Tris-HCl pH 7.5, 5 mM 2-mercaptoethanol, 10 mM EDTA) supplemented with 100 µM of S-adenosylmethionine (NEB) for 12 hr at 37°C. Finally the plasmid was treated with Proteinase K (37C overnight in presence of 0.5% SDS; NEB) to degrade the crosslinked M.Hpall to make a 4 amino-acid peptide adduct.

To initiate protein recruitment to the damaged plasmid, pGAP was gently mixed with mock- or PARP1-depleted HSS at 10 ng/µL (final concentration), supplemented either with PARP1 storage buffer (25 mM HEPES pH 8.0, 150 mM NaCl, 1 mM EDTA and 0.1 mM TCEP) or with recombinantly purified PARP1 variants at a final concentration of 15 ng/µL. At 5 and 10 min, 8 µL of reaction was withdrawn and plasmids and associated proteins were recovered via plasmid pull-down using Lacl coated magnetic beads as described previously.⁹² Briefly, 6 µL Streptavidin-coupled magnetic beads (Dynabead M-280, Invitrogen) per pull-down reaction were washed and equilibrated using Lacl binding buffer (50 mM Tris-HCl pH 7.5, 150 mM NaCl, 1 mM EDTA, 0.02% Tween-20). The beads were then incubated with biotinylated Lacl at RT for 40 min, and then subsequently washed four times with Pull-Down Buffer (10 mM HEPES pH 7.7, 50 mM KCl, 2.5 mM MgCl₂, 250 mM sucrose, 0.25 mg/ml BSA, 0.02% Tween-20) and stored on ice until further use. Reaction samples at 5 and 10 min were gently mixed with the prepared Lacl coated magnetic beads and incubated for 30 min at 4°C with rotation. Then, samples were washed twice with wash buffer (10 mM HEPES pH 7.7, 50 mM KCl, 2.5 mM
MgCl2, 250 mM sucrose, 0.03% Tween-20), and once with wash buffer without detergent (10 mM HEPES pH 7.7, 50 mM KCl, 2.5 mM MgCl2, 250 mM sucrose). Samples were washed one additional time in 50 µL of wash buffer without detergent and transferred to a new LoBind tube (Eppendorf) to remove residual detergent. The supernatant was removed, and the plasmid-bound beads were re-suspended in 50 µL ice-cold 50 mM ammonium bicarbonate buffer supplemented with 500 ng modified sequencing grade trypsin (Promega) to facilitate on-bead digestion at 30°C overnight with mild shake. Beads were separated from peptides by filtering the digestion mix through a 0.45 µm spin filter (Ultrafree-MC, Millipore). Samples were then reduce-alkylated by the addition of TCEP and CAA to final concentrations of 5 mM. Peptides were purified for MS analysis by high-pH StageTips as essentially described above, then eluted with 25% acetonitrile in 50 mM ammonium hydroxide into LoBind tubes, and subsequently dried to completion using a SpeedVac for 2 h at 60°C, after which the dried peptides were dissolved in 11 µL of 0.1% formic acid and stored at -20°C until MS analysis.

## Purification of ADPr-modified proteins during S-phase for MS analysis

Using the double-thymidine method, we synchronized PARP1 WT and 4SA mutant cells in their G1/S boundary and released them into S-phase. To achieve this, cells seeded in quadruplicates in 10 cm dishes (corning), and arrested cells for 17 h by adding thymi-dine (Merck) to a total concentration of 2 mM in the culture medium. Then, cells were released for 9 h, and double-blocked with the addition of 2 mM thymidine for another 17 h. Cells were released for 1 h, at which point they received either DMSO or 10 µM FEN1-inhibitor (Tocris) for an additional 1 h. Two hours following their release into S-phase, cells were washed in ice-cold PBS two times, scraped and immediately lysed in 5 mL RIPA buffer (50 mM Tris-HCl pH 8.0, 100 mM NaCl, 1% Triton X-100) supplemented with 5 mM MgCl2, 0.1% Benzonase (Millipore), protease and phosphatase inhibitors (Roche), 2 µM Olaparib (Selleckchem), and 1 µM PARGi PDD00017273 (Tocris) for 30 min at 4°C. Then, samples were centrifuged a 4250 × g to pellet debris, and the supernatants were split evenly across either Af1521-linked glutathione Sepharose 4B beads or empty control beads (Cytiva). The purification of the GST-Af1521 macrodomain was performed essentially as described previously.32 The beads were incubated with the lysates for 3 h at 4°C tumbling end over end to facilitate binding between the ADP-ribosylated proteins and the Af1521 containing beads. After binding, the beads were transferred to new LoBind tubes 3 times and were washed 2 times between each tube change in RIPA buffer (described above) without supplements. Proteins were digested on-beads with RIPA buffer supplemented with 500 ng of trypsin (Promega, Sequencing Grade) per sample, and incubated over night at 30°C with mild shake. To separate the peptides from the beads, samples were spun through a 0.45 µM filter (Ultrafree-MC, Millipore), and then reduced and alkylated with 5 mM TCEP and 5 mM chloroacetamide for 30 min at RT. Peptides were subjected to StageTip cleanup by desalting and purifying them on C18 discs at high pH, as described above, and eluted at 25% acetonitrile in 50 mM ammonium hydroxide. All samples were dried to completion in protein LoBind tubes (Eppendorf), using a SpeedVac for 2 h at 60°C, after which the dried peptides were dissolved using 11 µL of 0.1% formic acid and stored at -20°C until MS analysis.

## Purification of ADPr-modified PARP1 protein while preserving ester-linked modifications

To enrich in vivo auto-modified PARP1, samples were prepared by culturing a 10 cm dish in 4 replicates of U2OS cells with following genetic backgrounds: PARP1 WT, PARP1 4SA, and HPF1 KO. Then, PARP1 activation for WT and 4SA mutants was achieved by treatment with either 0.02% (v/v) MMS (Sigma Aldrich) for 30 min by addition to the culture medium at 37°C. For HPF1 KO cells we activated PARP1 by addition of 2 mM hydrogen peroxide ($H_2O_2$, Sigma Aldrich) for 10 min to the culture medium at 37°C. We also prepared in vitro auto-modification reactions by combining 1 µM of purified PARP1 WT or PARP1 E988K mutant with a 30nt damaged DNA template at 1 µM, supplemented with 1 mM NAD⁺ to stimulate synthesis of ADP-ribosylation. The reaction was carried out in ADPr reaction buffer (25mM HEPES pH 8.0, 150 mM NaCl, 0.5mM TCEP). Purified HPF1 protein was added to half of the reactions at equimolar ratio to PARP1 enzyme; otherwise the reactions received ADPr reaction buffer as a control. Cells were washed two times with ice-cold PBS, and then scraped and pelleted at 500 x g at 4°C. Cells were subsequently lysed in modified RIPA buffer inspired by Longarini and Matić³⁵ (20mM HEPES pH 7.0, 150 mM NaCl, 2.5 mM MgCl2, 1% NP-40, 0.5% Sodium Deoxycholate, 0.1% SDS) supplemented with 2 mM Olaparip, 1 mM ADP-HPD, 750 U/mL Benzonase, and 1 pill of EDTA-free Protease Inhibitor Cocktail. For the sake of comparison, in vitro reactions and in vivo samples are both subjected to this lysis buffer, and are processed simultaneously while undergoing the same workflow as described below. The samples are lysed in 250 µL modified RIPA buffer on ice for 30 min, and then diluted with 1000 µL of the same modified RIPA lysis buffer, where MgCl2 and Benzonase was omitted in place of 30 mM EDTA to quench Benzonase reactivity. Samples were centrifuged for 10 min with 16,000 x g at 4°C and the supernatant was collected and transferred to pre-chilled tube containing 30 µL of pre-equilibrated PARP1-nanobody agarose beads (Chromotek). To facilitate protein binding, the tubes tumbled end-over-end for 1 hour at 4°C, after which bound beads were pelleted with 500 x g at 4°C and the supernatant was discarded. Beads were transferred to clean tubes, then washed 3x in ice-cold wash buffer (20 mM HEPES pH 7.0, 150 mM NaCl, 1 mM EDTA, 0.1 mM DTT), and then transferred to clean tubes to prevent carry-over of detergents and inhibitors. At this point, the samples were split into two conditions, and treated with 500 nM PARG enzyme or mock control in PARG reaction buffer (20 mM HEPES, pH 7.2, 50 mM NaCl, 5 mM MgCl2) at RT for 30 min. Then samples were washed stringently to remove non-specific and co-immunoprecipitated proteins. Firstly, two ice-cold washes in modified RIPA buffer (20mM HEPES pH 7.0, 150 mM NaCl, 30 mM EDTA, 1% NP-40, 0.5% Sodium Deoxycholate, 0.1% SDS) followed by a tube change. Secondly, two ice-cold washes in high-salt buffer (20 mM HEPES pH 7.0, 500 mM NaCl, 1 mM EDTA, 0.1 mM DTT) followed by a tube change. Lastly, two ice-cold washes with a low-salt buffer (20 mM HEPES pH 7.0, 1 mM EDTA, 0.1 mM DTT) followed by a tube change.
Proteins were then on-bead digested from dry beads with Lys-C at low pH in 200 µL digestion buffer (1.25 ng/µL Lys-C, 25 mM PIPES pH 6.5, 3M guanidine, 150 mM NaCl, 0.1 mM TCEP) prepared immediately prior to adding it to the samples, after which they were digested at 30°C for 1 hour. The addition of guanidine increases the missing cleavage rate, which is a prerequisite for increasing the number of ions with high charge-states eligible for ADPr localization using ETD fragmentation, and furthermore increases improves coverage of lysine-rich regions. The digestion was stopped by acidifying the samples to 0.5% TFA (v/v), and peptides were separated from beads by spinning them through a 0.45 µM filter (Ultrafree-MC, Millipore). Peptides were purified using low-pH StageTips with four-layer C18 SPE discs essentially as described above, except the washing and elution buffers contained 0.1% formic acid (pH ~3) instead of ammonia, and elution was performed using 30% acetonitrile in 0.1% formic acid to sufficiently elute longer peptides products resulting from Lys-C digestion. We note that alkylation of the thiol group of cysteines require incubation with CAA at pH > 8 using heating,⁹³ which would potentially remove ester-linked modifications and thus this step was omitted. However, the thiol groups are already reduced in our experimental setup and do not reform di-sulfide bridges at acidic pH since the -SH group remains protonated.⁹⁴ Eluted peptides were dried to completion at pH ~3 using a SpeedVac for 2 h at 30°C, after which the dried peptides were dissolved using 11 µL of 0.1% formic acid and stored at -20°C until MS analysis.

## MS instrument analysis

### Dataset overview

Dataset 1: ADP-ribosylome of PARP1 wild type and 3SA mutant cells in response to hydrogen peroxide and MMS, Lys-C and trypsin digestion. Related to Figure 1 and Table S1.

Dataset 2: ADPr sites on PARP1 with preservation of ester-linked PTMs. Related to Figure 1 and Table S2.

Dataset 3: ADP-ribosylome of PARP1 wild type and 4SA cells in response to MMS, Lys-C digestion. Related to Figures 1 and 2 and Table S3.

Dataset 4: *Xenopus laevis* plasmid pull-down for mass-spectrometry. Related to Figure 2 and Table S4.

Dataset 5: Targets of ADP-ribosylation during S-phase. Related to Figure 4 and Table S5.

### ETD-based mass spectrometric analysis

Samples for datasets 1, 2, and 3 were measured using and Orbitrap Fusion Lumos Tribrid mass spectrometer (Thermo), and analyzed on 20-cm long analytical columns with an internal diameter of 75 µm, packed in-house using ReproSil-Pur 120 C18-AQ 1.9 µm beads (Dr. Maisch). On-line reversed-phase liquid chromatography to separate peptides was performed using an EASY-nLC 1200 system (Thermo), and the analytical column was heated to 40°C using a column oven. Peptides were eluted from the column using a gradient of Buffer A (0.1% formic acid) and Buffer B (80% ACN in 0.1% formic acid). For dataset 1, high pH fractions; main gradient ranged from 3% buffer B to 32% buffer B over 62 min, followed by a brief increase to 40% buffer B to elute all peptides, and a subsequent wash block of 18 min. Total gradient length was 80 min. For dataset 1 low-pH fraction; main gradient ranged from 5% buffer B to 15% buffer B over 27.5 min, followed by a 7.5min ramp to 30% buffer B, and a subsequent wash block of 12 min. Total gradient length was 50 min. For dataset 2, single-shot elution; main gradient ranged from 3% buffer B to 24% buffer B over 50 min, followed by a 12 min ramp to 40% buffer B, and finally a wash block. Total gradient length was 80 min. For dataset 3, high-pH fractions we used the same gradient as described above, but compressed to a total length of 60 min.

Electrospray ionization (ESI) was achieved using a nanospray Flex Ion Source (Thermo). Spray voltage was set to 2 kV, capillary temperature to 275°C, and RF level to 30%. For dataset 1, full scans were performed at a resolution of 120,000 using a scan range of 300-1750 m/z with a maximum injection time of 50ms. The automatic gain control (AGC) was set to a target of 600,000 charges. For fragmentation, precursor ions were isolated in a 1.3 m/z window around the peak, with an AGC target of 200,000 charges and a maximum injection time of 180 ms, and the peptides were fragmented by electron transfer disassociation with supplemental higher-collisional disassociation (EThcD) at 20 NCE. To be eligible for fragmentation, we implemented a decision tree logic for selecting precursors for isolation; the intensity of the ion is larger than 110000. The charge state is 3, 4, or 5 prioritized in that order. A dynamic exclusion time of 60 s. Fragmented ions were analyzed in the Orbitrap with a resolution of 60,000 with a topN = 4 in data dependent acquisition (DDA) mode. For dataset 3, largely the same instrument parameters were used, except for the following: precursor scan range was set to 300-1300 m/z for all methods. For fraction 1, 2 and 3 the dynamic exclusion time was set to 45 s. For fraction 2 and 3, the intensity threshold for precursor isolation was lowered to 80,000 charges with a topN = 3, and a maximum injection time of 250 ms. For dataset 2 (PARP1 affinity purification), samples were eluted using a 66 min main gradient, ramping from 5% buffer B to 40% for 50 min, followed by a steeper gradient ranging from 40-50% buffer B over the course of 11 min, and finally followed by a wash block to make the total gradient 80 min. The instrument settings for dataset 2 were largely identical to what is described above, except dynamic exclusion time was set to 30 s, and quadrupole isolation window for precursor ions was set to 2 Da with a total injection time of 120 ms with topN = 5.

### HCD-based mass spectrometry analysis using Orbitrap Exploris 480

Plasmid pull-down samples for dataset 4 were measured using an Orbitrap Exploris 480 mass spectrometer (Thermo), using the same HPLC, column, and oven setup as outlined above. Peptides were analyzed as single-shot fractions, using gradients with a total length of 80 min. The main gradient was 60 min, ranging from 5% buffer B to 34% buffer B, with a 5 min ramp to 44% buffer B, followed by a 15 min wash block. Spray voltage was set to 2 kV, capillary temperature to 275°C, and RF level to 40%. For precursor scans, we used a scan range of 300-1300 m/z, with a maximum injection time of 60 ms for 200,000 charges, analyzed at 120,000 resolution using the Orbitrap. For precursor fragmentation, ions were isolated around a 1.3 m/z window, if their intensity was at least
170,000 charges with a charge state of 2–6. Dynamic exclusion time was set to 30 s. For fragmentation of ions, the instrument was set to data-dependent acquisition mode, running a topN = 7 method, and fragmentation was achieved using HCD with a collision energy of 25 % using a 58 ms injection time and an AGC target of 200 %. The fragmented MS/MS spectra were analyzed in the Orbitrap at 60,000 resolution.

HCD-based mass spectrometry analysis using Orbitrap Astral

Samples containing ADPr-modified proteins during s-phase for dataset 5 were measured using an Orbitrap Astral mass spectrom-
eter (Thermo), using the vanquish Neo UHPLC system (Thermo) for on-line reversed-phase liquid chromatography, with a column and
oven setup as described above. Peptides were analyzed as single-shot elution, using gradients with a total length of 40 min. The main
gradient was 30 min, ranging from 5- to 32% buffer B, after which a 3 min increase to 42 % buffer B was performed to elute all pep-
tides. The main gradient was followed by a 7 min wash block to clean the column between samples. Peptides were ionized using ESI
with a spray voltage of 2 kV, RF lens was set to 50%, and the capillary temperature to 275°C. For the full range master scan, we sam-
ple ions from the ion beam at a scan range of 300-1300 m/z, with a maximum injection time of 150 ms coupled with a 250% AGC
target, and analyze their m/z in the Orbitrap at 120,000 resolution. Ions eligible for fragmentation contained a charge state of 2-6,
and to prevent re-sampling we enabled dynamic exclusion with an exclusion window of 10 sec. We prioritize ions above 1e5 intensity,
but allow a sensitive scan priority for ions as low as 2.5e4. Fragmentation was achieved using HCD with a collision energy of 25%. The
resulting ion fragments were detected in the Astral mass analyzer with a scan range of 100-1500 m/z, normalized AGC target of 100%
and maximum injection times of 5 ms, or 20 ms in for the sensitive scan priority. The instrument recorded scans in data-dependent
acquisition mode, with a duty cycle of 300 ms.

MS data processing

Data processing of site-specific ETD-based ADPr data

Analysis of the MS raw data was performed using MaxQuant software,⁸⁸,⁹⁵ version 1.5.3.30. Datasets 1, 2, and 3 were analyzed separately. The search-space for dataset 1, 2, and 3 was generated from a fasta file containing all human proteins and their isoforms downloaded from UniProt on the 9th of December 2020 (DS1) and 25th of June 2023 (DS2+3). We also included the sequences of PARP1 mutants specific to each dataset as fasta files. MaxQuant default settings were used with the following exceptions as outlined below. For dataset 1, we allowed N-terminal acetylation, methionine oxidation, cysteine carbamidomethylation, and ADP-ribosylation on all amino acid residues known to potentially be modified (C, D, E, H, K, R, S, T, and Y), as variable modifications. We allowed up to 6 missed cleavages with trypsin digestion, and 4 variable modifications per peptide. To increase our confidence for correctly identifying modified peptides, the minimum delta score for modification-specific peptides was raised to 20. Matching between runs was enabled, but second-peptide search was not. For dataset 3, the same parameters as outlined above were used, except for the following changes: we restricted the in-silico digestion to LysC-specific peptides, allowing for 3 missed cleavages per peptide. We set the maximum peptide mass to 6000 for dataset to accommodate longer peptides generated with LysC. For dataset 2, similar parameters were used as described above, except we allowed 5 missing cleavages for LysC-specific peptides, with 4 variable modifications. Data were automatically filtered by posterior error probability to achieve a false discovery rate of <1% (default), at the peptide-spectrum match and the protein assignment levels.

Data processing of PP-MS data

The *Xenopus* egg extract plasmid pull-down for MS analysis was processed using MaxQuant (version 1.5.3.30, as described above), and searched with a FASTA file specific to *Xenopus laevis* downloaded on the 11th of February 2023, and additionally the sequences of the PARP1 mutants. We used the default MaxQuant settings, “Fast LFQ” disabled, and second peptide search was enabled. Variable modifications was permitted for N-terminal acetylation, methionine oxidation, while we restrict cysteine carbamidomethylation as a fixed modification. Stringent MaxQuant 1% FDR data filtering at the PSM- and protein-levels was applied (default). To fix missing or unannotated gene names from the *Xenopus* fasta database, we manually curated these genes where possible. To do so, we extracted the multiple gene names from the fasta file for each protein group, and collapsed short and long format versions of genes. Then we annotated missing gene names with the following by ranked priority: alternative gene names found in the fasta header, genes names (primary) as reported by the UniProt database accessed concomitantly with the fasta file, and finally a manually curated in-house database, as described previously.⁹¹ To construct this database, the UniProt ID from the fasta file served as key, and we drew value-pairs for missing or uninformative gene names still not annotated correctly from the following resources in order, where possible: UniProt, otherwise Xenbase, otherwise the Session et al. database,⁹⁶ otherwise RefSeq (via Xenbase), and otherwise via InterPro annotations.

Data processing of S-phase ADPr-target data

The S-phase ADPr-target raw files were processed using MaxQuant (version 2.4.8.0), and searched with a human FASTA file down-
loaded 23rd of June 2023 from the UniProt database, supplemented with the sequence for PARP1 4SA. The default MaxQuant set-
tings were used with the following exceptions: The minimum peptide length was 6 and the maximum peptide mass for the
Andromeda search space was increased to 6000, with a minimum delta score for modified peptides set to 50 and a minimum overall
Andromeda score of 100. Second peptide search was disabled and matching-between runs was enabled with matching window
of 30 s, and an alignment time window of 10 min. For the MS/MS spectra, we used a match tolerance to 15 FTMS top x mass window of
200 Da ensure stringency. All modified peptides were used for unrestricted protein quantification, and included deamidation of N/Q
and glutamine to pyroglutamic acid conversion as variable modifications. Stringent MaxQuant 1% FDR data filtering at the PSM- and
protein-levels was applied (default).
# QUANTIFICATION AND STATISTICAL ANALYSIS

## Data filtering of site-specific ETD-based global ADPr data

In addition to the automatic filtering and control of false discovery rate implemented by MaxQuant, dataset 1 and 3 were manually and stringently filtered to ensure proper identification, quantification and localization of the ADP-ribose onto peptides. Peptide spectrum matches (PSM) modified by more than one ADPr-moiety was omitted. PSMs was considered for identification of the localization probability was >0.9, and for quantification if it was at least 0.75. Flawed MaxQuant intensity assignments were manually corrected in the sites table and based exclusively on localized PSMs. We only considered MS/MS and multi-MS/MS for identification of sites, but we allowed multi-match for quantification.

## Data filtering of PARP1 ADPr sites recorded with ETD fragmentation while preserving ester-linked modifications

The raw MaxQuant output was further processed to ensure proper localization and quantification of sites found from ADPr-modified peptides detected in the experiment. Since experimental conditions could lead to different types of ADPr-linkages (i.e. HPF1 may promote serine-ADPr) it is particularly important that each experiment stands alone and does not extrapolate localization scores from other conditions to pass filtering. To this end, we extracted all individual localization probabilities within each modified peptide and discarded sites with a localization probability below 0.9. In case the peptide was multi-ADP-ribosylated, we considered each site within that peptide in isolation to prevent flawed intensity assignments to mislocalized sites. Next, we mapped intensities assigned by MBR, by matching the exact modified peptide sequence which passed our localization criteria. Furthermore, we only allowed transfer of MBR intensities if at least one MS/MS within that experimental group had been correctly localized for each modified peptide. This is important to prevent intensity mapping by MBR to modified peptides that are isobaric (i.e. identical mass), but actually have no underlying evidence to support a site for a particular condition, since such isobaric peptides elute with near-identical retention times. As an additional quality filter, we discard ions with MULTI-MATCH scores below 200 to only retain high-quality matches. To account for different expression levels of PARP1 and variable loading, we scaled the site-intensities by normalizing the total PARP1 intensity within each experiment to its global median. For a direct quantitative comparison between PARP1 WT and PARP1 4SA mutant auto-modification intensities, peptide-level localization is sufficient since no site-specific localization is required, meaning no additional filtering was applied. Statistical significance was determined using Student's T-test, *p* < 0.001.

## Quantification and statistical analysis of MS data

To further filter and analyze the processed raw files from MaxQuant, we used a combination of the statistical programming language R (version 4.3.1 "Beagle Scouts") and the Perseus software suite⁹⁷ Gene set enrichment analysis was performed using multiple Fisher's exact test with Benjamini-Hochberg multiple-hypothesis correction applied to test for enriched terms. Protein Gene Ontology annotations were downloaded concomitantly with the fasta files from the UniProt database. Volcano plots were generated using Perseus, implementing the S0 fudge-factor and multiple-hypothesis t-testing with FDR correction. We used the following significance thresholds: *p*-value < 0.01 applied with a fudge factor s0 = 0.5 (Figures 1C, S1E, and S2C); *p*-value < 0.05 applied with a fudge factor s0 = 0.1 (Figures 2H, 2I, 4D, S2G, S2H, and S4D-S4F). Principle component analysis was performed in R using the prcomp function. Heatmaps were generated using the plothmap package (version 1.0.12), after scaling the intensities row-wise to normalize them. Clustering was performed using Euclidian distances for both rows and columns, using the "average" clustering method. UpSet plots were generated using the UpSetR package (version 1.4.0),⁹⁸ using only ADPr sites identified directly by MS/MS as evidences for identification. Sequence context analysis for S-ADPr-sites were performed using the ice-Logo algorithm,⁹⁹ using the serine ADPr-modified sequence motifs as foreground, and all serine residues of the human proteome as a background, with theoretical charge-states of background peptides set to 3 or higher, accounting for MS acquisition settings. The protein network with annotations specific to Figure 4 was made using cytoscape (version 3.10.1), in combination with the STRING database and Omics Visualizer App.⁸⁹,¹⁰⁰,¹⁰¹